Handbook of Neurochemistry and Molecular Neurobiology: Neural Lipids [3rd ed.] 9780387303451, 0387303456

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Handbook of Neurochemistry and Molecular Neurobiology Neural Lipids

Abel Lajtha (Ed.)

Handbook of Neurochemistry and Molecular Neurobiology Neural Lipids Volume Editors: Guido Tettamanti and Gianfrancesco Goracci

With 120 Figures and 56 Tables

Editor Abel Lajtha Director Center for Neurochemistry Nathan S. Kline Institute for Psychiatric Research 140 Old Orangeburg Road Orangeburg New York, 10962 USA Volume Editors Guido Tettamanti Department of Medical Chemistry, Biochemistry and Biotechnology Via Saldini 50 20133, Milan Italy

Gianfrancesco Goracci Department of Internal Medicine Section of Biochemistry University of Perugia via del Giochetto 06122 Perugia Italy

Library of Congress Control Number: 2006922553 ISBN: 978‐0‐387‐30345‐1 Additionally, the whole set will be available upon completion under ISBN: 978‐0‐387‐35443‐9 The electronic version of the whole set will be available under ISBN: 978‐0‐387‐30426‐7 The print and electronic bundle of the whole set will be available under ISBN: 978‐0‐387‐35478‐1 ß 2009 Springer ScienceþBusiness Media, LLC. All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science+Business Media, LLC., 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. springer.com Printed on acid‐free paper

SPIN: 11416685 2109 – 5 4 3 2 1 0

Preface

The second Edition of the ‘‘Handbook of Neurochemistry’’ goes back to 1983. In that Edition, Brain lipids were distributed in different volumes, following the rationale underlying the Edition. Many of the chapters on lipids were outstanding and actually are ‘‘historical masterpieces’’ of scientific literature. After more then 25 years, the lipids of the nervous system were considered to deserve a separate volume. Many are the reasons for this decision. New methods have been developed for the structural analysis of lipids, for their quantification at the nano- and pico-mole levels, for the synthesis of analogs and derivatives suitable for biological investigations. Lipids entered into the ‘‘omic’’ era, too, and there is a consolidated ‘‘lipidomics’’. The metabolic pathways of lipids that 25 years ago appeared to be complex are presently in a way that is much more complex and intriguing, being intimately connected with the intricate network of intracellular molecular traffic. The impact of the new technologies for identifying genes, transfecting them into cells, and overexpressing or silencing them was tremendous, in terms of innovation and growing knowledge. Of course, this also applies to the lipid field. However, serious perplexities were also generated, again regarding lipids, too. A similar situation applies to the exponential development of the use of transgenic animals: many findings were obtained that validated previous hypotheses. But unexpected results also emerged, which presumably reflect the present incomplete knowledge of the regulation mechanisms of gene expression. A further field that blossomed magnificently in recent decades is membrane lipidology, ranging from the release of fragments from membrane lipids, having a bioactive role, to the separation of some lipids and few proteins into more rigid domains (lipid rafts) holding peculiar properties, and the discovery of lipid anchors to protein. A completely novel notion is also the occurrence of bioregulators of sphingoid nature, deriving from membrane sphingolipids. Just to finish, surprising findings concern the role of lipids in a number of neural diseases and the relationship between diet lipids and brain function. The ‘‘Neural Lipids’’ volume of the new Edition of the Handbook of Neurochemistry and Molecular Neurobiology was conceived to offer an update on present knowledge of neural lipids, evidencing the new advances and concepts but recalling the old basic ones in a perspective of continuity. Notwithstanding the efforts, the resulting view may probably be incomplete. However, it is surely sufficient to convince especially the newcomers to the field of the importance of structural and functional lipidology. It is remarkable that some of the authors of the chapters collected in this Edition were authors of the previous edition, too: this is an unequivocal sign of continuity of interest and dedication to lipid science. To finish on a sad note, two authors of this volume, Prof. L.A.Horrocks, and Prof. S.E.Pfeiffer, passed away before the publication of the volume. Prof. H.Moser, expert in peroxisomal- physiopathology, also left us at the beginning of his engagement. Through the kind mediation of his wife, four of his co-workers took care of continuing and terminating the work. Lloyd, Steve and Hugo continue to live in our memory and unchanged appreciation. This volume is dedicated to them. Gianfrancesco Goracci Guido Tettamanti

Table of Contents

Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xi

Biochemistry and Molecular Biology of Neural Lipids 1

Advances in Lipid Analysis/Lipidomics – Analyses of Phospholipids by Recent Application of Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 R. Taguchi

2

Choline and Ethanolamine Glycerophospholipids . . . . . . . . . . . . . . . . . . . . . . 21 A. A. Farooqui . L. A. Horrocks . T. Farooqui

3

Brain Phosphatidylserine: Metabolism and Functions . . . . . . . . . . . . . . . . . . . 39 R. Mozzi . S. Buratta

4

Metabolism and Enzymology of Cholesterol and Steroids . . . . . . . . . . . . . . . . 59 B. Stoffel-Wagner

5

Anandamide and Other Acylethanolamides . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 S. Petrosino . V. Di Marzo

6

Chemistry, Tissue and Cellular Distribution, and Developmental Profiles of Neural Sphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 G. Tettamanti . L. Anastasia

Cellular and Subcellular Localization of Neural Lipids 7

Nuclear Lipids and Their Metabolic and Signaling Properties . . . . . . . . . . . . 173 R. Ledeen . G. Wu

8

Lipids of Brain Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 199 L. Corazzi . R. Roberti

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Table of Contents

9

Neuronal Membrane Lipids – Their Role in the Synaptic Vesicle Cycle . . . . . 223 L. Lim . M. R. Wenk

10

Functional Dynamics of Myelin Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 239 S. N. Fewou . N. Jackman . G. van Meer . R. Bansal . S. E. Pfeiffer Function of Neural Lipids

11

The Phosphoinositides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 G. D’Angelo . M. Vicinanza . A. Di Campli . M. A. De Matteis

12

Lipid Mediators and Modulators of Neural Function: Lysophosphatidate and Lysolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 289 D. N. Brindley . A. U. Bra¨uer

13

Metabolism and Functions of Platelet-Activating Factor (PAF) in the Nervous Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 311 G. Goracci . M. L. Balestrieri . V. Nardicchi

14

Lipid Anchors to Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 353 N. N. Nalivaeva . A. J. Turner

15

Bioactive Sphingolipids: An Overview on Ceramide, Ceramide 1-Phosphate Dihydroceramide, Sphingosine, Sphingosine 1-Phosphate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 373 J. M. Kraveka . Y. A. Hannun

16

The Endocannabinoid System and its Manifold Central Actions . . . . . . . . . . 385 M. Maccarrone Diet, Brain Lipids and Brain Functions

17

Diet, Brain Lipids, and Brain Functions: Polyunsaturated Fatty Acids, Mainly Omega‐3 Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 409 J. M. Bourre

18

Choline and Its Products Acetylcholine and Phosphatidylcholine . . . . . . . . . 443 R. J. Wurtman . M. Cansev . I. H. Ulus

19

Alcohol and Neural Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 503 E. Cazzaniga . A. Bulbarelli . M. Masserini Lipids in Neural Disfunction and Diseases

20

Roles of Cytosolic and Secretory Phospholipases A2 in Oxidative and Inflammatory Signaling Pathways in the CNS . . . . . . . . . . . . . . . . . . . . . . . . . 517 G. Y. Sun . A. Y. Sun . L. A. Horrocks . A. Simonyi

Table of Contents

21

Lipids in Neural Tumors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 535 J. R. Van Brocklyn

22

Lipids in Alzheimer’s Disease Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 563 D. A. Butterfield . H. M. Abdul

23

Neural Lipids in Parkinson’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 583 M. Barichella . G. Pezzoli . A. Mauri . C. Savardi

24

Lipids in Multiple Sclerosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 593 L. Rinaldi . F. Grassivaro . P. Gallo

25

Brain Oxidative Stress from a Phospholipid Perspective . . . . . . . . . . . . . . . . 603 A. Brand-Yavin . E. Yavin

26

Peroxisomal Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 631 G. V. Raymond

27

Sphingolipid‐Inherited Diseases of the Central Nervous System . . . . . . . . . . 671 S. L. Hoops . T. Kolter . K. Sandhoff

28

Mouse Models with Gene Deletions of Enzymes and Cofactors Involved in Sphingolipid Synthesis and Degradation . . . . . . . . . . . . . . . . . . . 703 R. Jennemann . H.-J. Gro¨ne . H. Wiegandt . R. Sandhoff Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 743

ix

Contributors

Hafiz Mohmmad Abdul Sanders-Brown Center on Aging, University of Kentucky, Lexington, KY 40536, USA

Anja U. Bra¨uer Institute of Cell Biology & Neurobiology, Center for Anatomy, Charite´-Universita¨tsmedizin Berlin, Phillipstrasse 12, 10115 Berlin, Germany

Luigi Anastasia Department of Medical Chemistry, Biochemistry and Biotechnology Via Saldini 50, 20133, Milan, Italy

David N. Brindley Signal Transduction Research Group, Department of Biochemistry, University of Alberta, Edmonton, Alberta, T6G 2S2, Canada Email: [email protected]

Maria Luisa Balestrieri Department of Biochemistry and Biophysics, Second University of Naples, Via L. De Crecchio 7, 80138 Naples, Italy Rashmi Bansal Department of Neuroscience, University of Connecticut Medical School, 263 Farmington Avenue, Farmington, CT 06030-3401, USA Michela Barichella Parkinson Institute, Istituti Clinici di Perfezionamento, Via Bignami, 1 20126 Milan, Italy Email: [email protected] Jean Marie Bourre INSERM, U 705, CNRS, UMR 7157, 200 rue du Faubourg Saint Denis, 75745 Paris cedex 10, France Email: [email protected]

Annette Brand-Yavin IBCHN, London Metropolitan University, 166-220 Holloway Road, London N7 8DB, UK

James R. Van Brocklyn The Ohio State University Medical Center, 4164 Graves Hall, 333 W. 10th Ave., Columbus, OH 43210 , USA Email: [email protected] Alessandra Bulbarelli Department of Experimental Medicine, University of Milano Bicocca, Via Cadore, 48, 20052 Monza (MI), Italy Sandra Buratta Department of Internal Medicine, Biochemistry Section, University of Perugia, via del Giochetto, 06122 Perugia, Italy D. Allan Butterfield Department of Chemistry, Center of Membrane Sciences, and Sanders-Brown Center on Aging, University of Kentucky, Lexington KY 40506, USA Email: [email protected] Antonella Di Campli Department of Cell Biology and Oncology, Consorzio Mario Negri Sud, 66030 Santa Maria Imbaro (Chieti), Italy

xii

Contributors Mehmet Cansev Department of Brain and Cognitive Sciences, Massachusetts Institute of Technology, Cambridge MA, 02139, USA Department of Pharmacology and Clinical Pharmacology, Uludag University Medical School, Gorukle, Bursa, 16059, Turkey Emanuela Cazzaniga Department of Experimental Medicine, University of Milano Bicocca, Via Cadore, 48, 20052 Monza (MI), Italy Email: [email protected] Lanfranco Corazzi Department of Internal Medicine, Section of Biochemistry, University of Perugia, 06122 Perugia, Italy Email: [email protected] Giovanni D’Angelo Department of Cell Biology and Oncology, Consorzio Mario Negri Sud, 66030 Santa Maria Imbaro (Chieti), Italy Akhlaq A. Farooqui Department of Molecular and Cellular Biochemistry, 1645 Neil Avenue, Columbus, Ohio 43210-1218, USA Tahira Farooqui Department of Molecular and Cellular Biochemistry and Department of Entomology, The Ohio State University, Columbus, Ohio 43210, USA Simon Ngamli Fewou Department of Neuroscience, University of Connecticut Health Center, P.O. Box 3401, 263 Farmington Avenue, Farmington, CT 06030-3401, USA Email: [email protected], [email protected] Paolo Gallo Multiple Sclerosis Centre – Veneto Region, First Neurology Clinic – Department of Neurosciences, Via Giustiniani, 5 – 35128 Padova, Italy Email: [email protected]

Gianfrancesco Goracci Department of Internal Medicine, Section of Biochemistry, University of Perugia, via del Giochetto, 06122 Perugia, Italy Email: [email protected] Francesca Grassivaro Multiple Sclerosis Centre – Veneto Region, First Neurology Clinic – Department of Neurosciences, Via Giustiniani, 5 – 35128 Padova, Italy

Hermann-Josef Gro¨ne Department of Cellular and Molecular Pathology, German Cancer Research Center, Im Neuenheimer Feld 280, 69120 Heidelberg, Germany

Yusuf A. Hannun Department of Biochemistry and Molecular Biology, Medical University of South Carolina, Charleston, South Carolina 29425, USA Email: [email protected]

Silvia Locatelli Hoops Kekule´-Institut fu¨r Organische Chemie und Biochemie der Universita¨t Bonn, Gerhard-Domagk-Str. 1, 53121 Bonn, Germany

Lloyd A. Horrocks Department of Molecular and Cellular Biochemistry, The Ohio State University, Columbus, 1645 Neil Avenue, Columbus, Ohio 43210–1218, USA Email: [email protected]

Nicole Jackman Department of Neuroscience, University of Connecticut Medical School, 263 Farmington Avenue, Farmington, CT 06030-3401, USA Richard Jennemann Department of Cellular and Molecular Pathology, German Cancer Research Center, Im Neuenheimer Feld 280, 69120 Heidelberg, Germany Email: [email protected]

Contributors Thomas Kolter Kekule´-Institut fu¨r Organische Chemie und Biochemie der Universita¨t Bonn, Gerhard-Domagk-Str. 1, 53121 Bonn, Germany

Gerrit van Meer Membrane Enzymology Bijvoet Center / Institute of Biomembranes, Utrecht University, Padualaan 8, 3584 CH Utrecht, The Netherlands

Jacqueline M. Kraveka Division of Hematology/Oncology, Department of Pediatrics, Medical University of South Carolina, Charleston, South Carolina 29425, USA

Rita Mozzi Department of Internal Medicine, Biochemistry Section, University of Perugia, via del Giochetto, 06122 Perugia, Italy Email: [email protected]

Robert Ledeen New Jersey Medical School, UMDNJ, Dept. Neurology & Neurosciences MSB-H506, 185 South Orange Ave., Newark, NJ 07103, USA Email: [email protected] Lynette Lim Department of Biological Sciences, Centre for Life Sciences, 28 Medical Drive, #04-21, Singapore 117607 Mauro Maccarrone Department of Biomedical Sciences, University of Teramo, Teramo, Italy IRCCS C. Mondino, Mondino-Tor Vergata Center for Experimental Neurobiology, Rome, Italy Email: [email protected] Vincenzo Di Marzo Endocannabinoid Research Group at the Institute of Biomolecular Chemistry, National Research Council, Via Campi Flegrei 34, Comprensorio Olivetti, Bldg. 70, 80078 Pozzuoli (NA), Italy Email: [email protected] Massimo Masserini Department of Experimental Medicine, University of Milano Bicocca, Via Cadore, 48 20052 Monza (MI), Italy Maria Antonietta De Matteis Department of Cell Biology and Oncology, Consorzio Mario Negri Sud, 66030 Santa Maria Imbaro (Chieti), Italy Email: [email protected] Andrea Mauri Parkinson Institute, Istituti Clinici di Perfezionamento, Via Bignami, 1, 20126 Milan, Italy

Natalia N. Nalivaeva Institute of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK I.M. Sechenov Institute of Evolutionary Physiology and Biochemistry, Russian Academy of Sciences, 44 Moris Thorez avenue, 194223 St. Petersburg, Russia Email: [email protected] Vincenza Nardicchi Department of Internal Medicine, Section of Biochemistry, University of Perugia, via del Giochetto, 06122 Perugia, Italy Stefania Petrosino Endocannabinoid Research Group at the Institute of Biomolecular Chemistry, National Research Council, Via Campi Flegrei 34, Comprensorio Olivetti, Bldg. 70, 80078 Pozzuoli (NA), Italy Gianni Pezzoli Parkinson Institute, Istituti Clinici di Perfezionamento, Via Bignami, 1, 20126 Milan, Italy Steven E. Pfeiffer Department of Neuroscience, University of Connecticut Medical School, 263 Farmington Avenue, Farmington, CT 06030-3401, USA James Powers Department of Pathology, University of Rochester Medical Center, Rochester, NY, USA Gerald V. Raymond Department of Neurogenetics, Kennedy Krieger Institute, 707 North Broadway, Baltimore, MD 21205, USA Email: [email protected]

xiii

xiv

Contributors Luciano Rinaldi Multiple Sclerosis Centre – Veneto Region, First Neurology Clinic – Department of Neurosciences, Via Giustiniani, 5 – 35128 Padova, Italy

Albert Y. Sun Department of Medical Pharmacology and Physiology, University of Missouri, Columbia, MO 65211, USA

Rita Roberti Department of Internal Medicine, Section of Biochemistry, University of Perugia, 06122 Perugia, Italy

Ryo Taguchi Department of Metabolome, Graduate School of Medicine, The University of Tokyo, 7-3- 1 Hongo, Bunkyo-ku, Tokyo 113, Japan Email: [email protected]

Konrad Sandhoff Kekule´-Institut fu¨r Organische Chemie und Biochemie der Universita¨t Bonn, Gerhard-Domagk-Str. 1, 53121 Bonn, Germany

Guido Tettamanti Department of Medical Chemistry, Biochemistry and Biotechnology Via Saldini 50, 20133, Milan, Italy IRCCS Policlinico San Donato, Via Morandi 30, 20097 San Donato Milanese, Milan, Italy Email: [email protected]

Roger Sandhoff Department of Cellular and Molecular Pathology, German Cancer Research Center, Im Neuenheimer Feld 280, 69120 Heidelberg, Germany Chiara Savardi Parkinson Institute, Istituti Clinici di Perfezionamento, Via Bignami, 1, 20126 Milan, Italy Agnes Simonyi Biochemistry Department, M743, Medical Science Building, University of Missouri, Columbia, MO 65212, USA Steven Steinberg Department of Neurogenetics, Kennedy Krieger Institute, 707 North Broadway, Baltimore, MD 21205, USA Birgit Stoffel-Wagner Department of Clinical Biochemistry, University of Bonn, 53127 Bonn, Germany Email: [email protected] Grace Y. Sun Biochemistry Department, M743, Medical Science Building, University of Missouri, Columbia, MO 65212, USA Email: [email protected]

Anthony J. Turner Institute of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds, LS2 9JT, UK Email: [email protected] Ismail H. Ulus Department of Brain and Cognitive Sciences, Massachusetts Institute of Technology, Cambridge MA, 02139, USA Department of Pharmacology and Clinical Pharmacology, Uludag University Medical School, Gorukle, Bursa, 16059, Turkey Mariella Vicinanza Department of Cell Biology and Oncology, Consorzio Mario Negri Sud, 66030 Santa Maria Imbaro (Chieti), Italy Paul Watkins Department of Neurogenetics, Kennedy Krieger Institute, 707 North Broadway, Baltimore, MD 21205, USA Markus R. Wenk Department of Biological Sciences and Department of Biochemistry, Centre for Life Sciences, 28 Medical Drive, #04-21, Singapore 117607 Email: [email protected]

Contributors Herbert Wiegandt Department of Cellular and Molecular Pathology, German Cancer Research Center, Im Neuenheimer Feld 280 69120 Heidelberg, Germany

Richard J. Wurtman Department of Brain and Cognitive Sciences, Massachusetts Institute of Technology, Cambridge MA, 02139, USA Email: [email protected]

Gusheng Wu Department of Neurology and Neurosciences, University of Medicine and Dentistry of New Jersey, New Jersey Medical School, 185 So Orange Ave., Newark, NJ 07103, USA

Ephraim Yavin IBCHN, London Metropolitan University 166-220 Holloway Road, London N7 8DB, UK Email: [email protected]

xv

Section 1

Biochemistry and Molecular Biology of Neural Lipids

1

Advances in Lipid Analysis/ Lipidomics – Analyses of Phospholipids by Recent Application of Mass Spectrometry

R. Taguchi

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

2 2.1 2.2

Application of Soft Ionization by Mass Spectrometry for Lipidomics . . . . . . . . . . . . . . . . . . . . . . . . . . Several Practical Methods for Lipidomics by Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Untargeted and Global Analyses in Lipidomics by FTICRMS, UPLC-MS, or Shotgun LC‐MS/MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.1 Untargeted Lipidomics by FTICRMS with Flow Injection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.2 Untargeted Lipidomics by UPLC-MS with Highly Accurate MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.3 Untargeted and Shotgun Lipidomics by LC‐MS/MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Focused Lipidomics by Precursor ion Scanning or Neutral Loss Scanning by Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Targeted Methods for Lipidomics by Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

5 6 7 7 7 7 8 8

3 3.1 3.2 3.3

Application Results of Several Mass Spectrometric Methods for Lipidomics . . . . . . . . . . . . . . . . . . . 9 Application Results of Untargeted and Shotgun Analysis by LC‐MS/MS . . . . . . . . . . . . . . . . . . . . . . . . . 9 Application Results of Focused Lipidomics by Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Targeted Method using Expanded MRM for Lipidomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14

4

Future Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_1, # Springer ScienceþBusiness Media, LLC 2009

4

1

Advances in lipid analysis/lipidomics – analyses of phospholipids by recent application of mass spectrometry

Abstract: Mass spectrometry (MS) has become a most useful tool in the analysis of phospholipids. Analysis of molecular species of phospholipids adding to that of their classes and subclasses is necessary to elucidate their physiological functions. As analytical methods for lipidomics, basically three different types of approaches in the identification of phospholipid molecular species can be selected. The first one is shotgun LC‐MS/MS analysis with data‐dependent scan, the second one is structure‐related focused methods such as precursor ion scanning or neutral loss scanning. Both types of data can be subjected to our search engine, ‘‘Lipid Search’’ (http://lipidsearch.jp), and most probable molecular species can be obtained with their compensated ion intensities. The lipid database for this search engine was constructed theoretically from their structure similarities and variations in polar head groups and fatty carbonyl chains. And identified individual molecular species can be automatically profiling according to their compensated ion intensities. The third method, such as multiple reaction monitoring, is also important for detecting very small amounts of targeted molecules such as lipid mediators or oxidized lipid metabolites. The choice of these three different kinds of methods seems to be very important for neurochemical research for detecting different kinds of lipid metabolites such as unknown lipid ligands, focused class of lipids, or targeted minor lipid mediators. List of Abbreviations: CID, collision‐induced dissociation; ESI, electrospray ionization; HPLC, high‐performance liquid chromatography; LC, liquid chromatography; MS, mass spectrometry; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PI, phosphatidylinositol; PS, phosphatidylserine; SM, sphingomyelin; UPLC, ultra performance liquid chromatography

1

Introduction

Lipids are a class of molecules thought to be very important, not only as energy source or constituents of biological membrane, but also as functional molecules concerning the many regulation steps in biological process (Di Paolo et al., 2004). Furthermore, recent research has revealed the roles of lipids, such as mediators of signal transduction and ligands receptors. And these functionally important lipid metabolites seem to be extremely rich in nerve system. Lipidomics is an important field in metabolomics, and is growing very rapidly by the recent advance in mass spectrometry (Han and Gross, 1994 and 2005; Pulfer and Murphy, 2003). In lipidomics, techniques of mass spectrometry become very important. Furthermore, recent advances in mass spectrometry make it possible to get comprehensive analyses of lipid metabolites within the cells and tissues. Studies on lipidomics are essential to get further understanding of each physiological and biological function of proteins concerning lipid metabolism. In this process, studies on comprehensive profiling on lipid metabolites in the cells should be inevitable. In particular, to identify real lipid substrates for enzyme proteins, lipid ligands for receptor proteins, and lipid metabolites for its carrier proteins, lipidomics by mass spectrometry is very useful. Another aim of lipidomics is to identify lipid molecules from mass spectrometry (MS) data and get profiling patterns of alteration of these molecules under specific circumstances. In these analytical processes of profiling, elucidation of unknown pathway or exact lipid substrate specificity of new enzyme proteins can be investigated. Before the use of MS, phospholipids were mainly detected by identifying radioisotopes after thin layer chromatography, or by applying gas chromatography (GC) after derivatization (Yokoyama et al., 2000; Nor Aliza et al., 2001; Sana et al., 2002; Tserng and Griffin, 2003). But these methods can not be applied to identification of all molecules in a phospholipid mixture. By using classical ionization methods in mass spectrometry such as electron impact (EI) and chemical impact (CI), it has been very difficult to get molecular‐related ions without any collapse. In these ionizations, fragment patterns of each molecule are basically used for criteria of identifications. Because of this reason, these methods were exclusively used for the mass measurement of purified single molecules. For the mixture, such as GC‐MS were used after derivatization for effective separation and analytical sensitivities. But for the molecules difficult to be evaporated and ionized, useful methods such as GC‐MS were not available. Thermospray ionization and atmospheric pressure chemical ionization (APCI) were also used in combination with high‐performance

Advances in lipid analysis/lipidomics – analyses of phospholipids by recent application of mass spectrometry

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liquid chromatography (HPLC) separation. In these process, partially effective methods such as fast atom bombardment (FAB) have been reported until common usage of electrospray ionization (ESI) or matrix‐assisted laser desorption/ionization (MALDI). ESI and MALDI make MS to be able to detect very small level of biological molecules. Recently, ESIMS has been used for the analyses of lipids. Within recent 10 years, there have been many improvements in MS, such as introduction of ESI‐TOFMS and ion‐trap MS. Molecular diversity of glycerophospholipids arises from the nature of the linkage and from the identity of the fatty side chain that is linked to the sn‐1 and sn‐2 carbon atom. In the analytical methods for lipidomics by mass spectrometry, adding to the comprehensive and untargeted analysis, focused or targeted analyses for categorical components are very important. It is very difficult to obtain exact identification of all metabolites even in the limited classes of molecules such as lipid metabolites. This is caused by different extraction efficiency of individual metabolites, different solubility in analytical solvents, different ionic efficiency, and broad dynamic ranges of their existence in biological samples. Even in the case of proteomics, it is very difficult to detect small amounts of peptides or proteins in mammalian plasma because of very wide dynamic ranges of protein contents in plasma. This is exactly the same in lipid metabolites in most of biological samples. For detecting minor but physiologically important lipid molecules in the nerve system, specified technical strategies should be applied in selecting the detection methods including choice of HPLC system with most effective columns and that of the most suitable MS system and collision conditions. In this chapter, recent applications of mass spectrometry for the analyses of lipids, mainly on phospholipids and their metabolites, are addressed.

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Application of Soft Ionization by Mass Spectrometry for Lipidomics

Since ESI is a soft ionization method, each molecule in a mixture can be detected without any fragmentation (Han and Gross, 1994; Kerwin et al., 1994). However, in general only the major peaks will be detected if the sample is injected as a mixture without any LC separation. One of the solutions to this problem is to use specific detecting methods, such as precursor ion scanning and neutral loss scanning (Heller et al.,1988; Lehmann et al., 1997; Domingues et al., 1998); these scanning modes are often used for measurement of particular focused phospholipids (Brugger et al., 1997; Hsu and Turk, 2003). ESI (Fenn et al., 1989) and MALDI is very mild ionization methods compared with previous ionization methods ever used. Soft ionization in mass spectrometry has induced some paradigm changes in the applications of mass spectrometry in biological studies. Effective insight can be obtained by comprehensive analyses of metabolic molecules under genetically, environmentally, or physiologically different conditions. MALDI is essentially used as off‐line methods, whereas ESI can be used as a flow system, and is easily combined with on‐line separation systems such as HPLC or capillary electrophoresis (CE). Sensitivity of detection by ESI essentially depends on the concentration of molecules in the sample solution. Thus, for obtaining a highest sensitivity, it is very important to use low elution rate with small size of column. For this purpose, capillary or nana‐LC system combined with ESI has been used. Concerning metabolic molecules as target of metabolome, individual molecular structures are mostly known and relations of each metabolite are well studied. Thus, we can easily imagine their metabolic linkage from our former knowledge. From these circumstances, we will be able to get effective data from global (i.e. comprehensive) analysis of metabolites by mass spectrometric analyses, for elucidating new function of enzyme proteins including substrate specificities. By ESIMS, selective analyses of individual molecules in the mixture can be effectively obtained. Further, by using FTICRMS, more than several hundreds of different molecules in the mixture eluted at same retention time can be effectively and separately identified by its high resolution and accurate mass values (Marto et al., 1995; Fridriksson et al., 1999; Ivanova et al., 2001; Jones et al., 2003). Recent advances in this field made many hybrid types of MS systems such as ion‐trap and TOFMS. The most important feature of these ionization methods are that the individual natural molecules can be ionized without any fragmentation. Further, they made us possible to obtain very sensitive measurement such as

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pico‐ or femto‐mol level of metabolites. Thus this method is suitable to determine biological molecules with very small amounts.

2.1 Several Practical Methods for Lipidomics by Mass Spectrometry There are several different approaches for lipid analyses. Here, comprehensive analysis of mass spectrometry is classified into three different categories. These are essentially classified such as untargeted, focused, and targeted analyses. 1. Untargeted and global lipidomics Relatively major component can be identified with this method (1) FTICRMS with flow injection (or with LC) (2) UPLC-MS by highly accurate MS (3) LC‐MS/MS with data‐dependent scanning 2. Focused lipidomics More sensitive method than the first one for focused categories of lipids Precursor ion scanning and neutral loss scanning with flow injection (or with LC), Head group survey, fatty acyl survey. 3. Targeted lipidomics Most sensitive methods within these three methods Multiple reaction monitoring (MRM) with LC The individual molecular species of lipid metabolites can be essentially defined as pairs of m/z (a charged mass value/a number of charges) value of molecular‐related ions and their fragment ions. The first one is an untargeted, so‐called global or comprehensive method for detecting all molecules contained within extracted lipid samples without preliminary information for molecular‐related ions or their fragments (Taguchi et al., 2000; Houjou et al., 2005). In a case of the untargeted and comprehensive method, no preliminary expected data of m/z and fragments are used. The strategy of this method is all the detected peaks of molecular‐related ions were subjected to further analysis (with or without exact identification for such as principal component analysis). By using mass spectrometer with high resolution such as FTICRMS, this method can be also used without LC separation (Ishida et al., 2004), but in most cases this method is effectively applied to quadrupole or TOFMS, with using LC‐MS or LC‐MS/MS. Thus combination with proper HPLC separation and mass spectrometer with high resolution such as FTICRMS of TOFMS is preferred. But even in the cases of identification only by mass values from molecular‐related ions, information of the retention time is important to obtain separate detection of isobaric molecular ions. Another important factor is information of fragment ions obtained by MS/MS. Thus application of LC‐MS/ MS with data‐dependent scanning mode is useful and practical method same as in proteomics. Data‐ dependent MS/MS spectra can be obtained as a shotgun strategy for molecular‐related ion peaks with relatively high intensity (Houjou et al., 2005). The usage of fragmentation is applicable as several different situations or methods. To obtain the information of fragment ions from targeted molecular‐related ions is commonly applied in the cases such as structural confirmation of suspected molecules or structural identification of unknown molecules. The second one is a focused method detecting some categories of molecules comprehensively using specific fragments or neutral losses caused from specific feature of their chemical structures (Houjou et al., 2004, 2005; Taguchi et al., 2005; Ishida et al., 2005a, b). In this case, peaks of molecular‐related ions lower than detection limit of s/n value in the mass spectrum can be effectively identified separately from noise peaks. This method also can be effectively performed by the combination with separation by LC, in this case detection limit of minor components or molecules with low ionic efficiency can be highly improved with lowering the ion suppression by separate elution from other major ions with high ion efficiency (Taguchi et al., 2005). In the case of the focused method, data of target fragments or target neutral losses for these surveys are used, but m/z of molecular‐related ions are surveyed comprehensively (Houjou et al., 2004, 2005; Taguchi et al., 2005; Ishida et al., 2005a, b).

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The third one is a so‐cold targeted method which can be applied to the targeted molecules by using the information both for molecular‐related ions and their specific fragment ions. Thus each combination of m/z values of these ions should be selected for detecting each individual molecule. This method is very popular and commonly used as a quantitative method for target drugs or drug metabolites in pharmaceutical company. In the case of the targeted method, individual m/z data pairs of molecular‐ related ions and characteristic fragment ions are used. But even in this method, scanning for many theoretically expected m/z value pairs can be surveyed comprehensively. For the application of this method in lipidomics, comprehensively collected data for hypothetical pairs of characteristics fragment ions and individual molecular‐related ions including hypothetically constructed molecular‐related ions are used. Recent triple stage quadrupole MS can detect up to 300 hundred pairs of MRM (SRM) at only one LC runs. Thus we selected this method as one of comprehensive analytical methods to detect minor individual oxidized phospholipid molecular species. In this case, the detection limit of MRM is ten times higher than the case by using precursor ion scanning of oxidized fatty acid fragments.

2.2 Untargeted and Global Analyses in Lipidomics by FTICRMS, UPLC-MS, or Shotgun LC‐MS/MS 2.2.1 Untargeted Lipidomics by FTICRMS with Flow Injection In the case of FTICRMS, accurate mass less than 2 ppm is used as effective annotation. And in the case of connecting with LC separation, retention times of individual molecular‐related ions are additive information. Several different molecular‐related ions might have close m/z vales within 0.5 mass unit, and these peaks can not be effectively separated by quadrupole MS. Thus, the identification of these ions is obtained only after separation by HPLC in the case of quadrupole‐ and TOFMS. While mass resolution of higher than 100,000 and mass accuracy of less than 2 ppm is easily obtained by FTICRMS, thus two molecular‐ related ions containing different atomic compositions can be effectively identified (Ishida et al., 2004). But even by FTICRMS, isobaric ions with exactly same atomic composition but with different structure can not be efficiently identified without LC separation.

2.2.2 Untargeted Lipidomics by UPLC-MS with Highly Accurate MS More than a thousand molecular species of phospholipids and neutral lipids were effectively detected by using high separation UPLC system even with a highly accurate single MS system. Subtraction data from such as control and disease samples with different lipid profiles can be effectively obtained. Automated profiling software for detecting highly increased or decreased MS peaks are now undergoing in our laboratory for next version of Lipid Search. Succeeding 2nd LC-MS/MS for selected targeted MS peaks were further applied to identify these peaks. Combination of target discovery by untargeted LC-MS and target confirmation by succeeding LC-MS/MS for targeted molecules is very effective approach in metabolomics (will be reported elsewhere).

2.2.3 Untargeted and Shotgun Lipidomics by LC‐MS/MS Combination of LC and data‐dependent shotgun MS/MS is another approach in untargeted lipidomics (Houjou et al., 2005). Separation of phospholipids on a normal‐phase column mainly depends on the character of the polar head group. For example, elution follows the order phosphatidylinositol (PI), phosphatidylethanolamine (PE), phosphatidylserine (PS), phosphatidylcholine (PC), and sphingomyelin (SM) (Taguchi et al., 2000; Houjou et al., 2005). Although PC species, which are the major molecules in many cells and have a high ionization efficiency in ESI, prevent the detection of other classes, the number of molecules that can be identified increases remarkably by separation into separate classes using normal‐ phase liquid chromatography (NPLC)‐ESI/MS (Taguchi et al., 2000; Houjou et al., 2005). Separation was

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observed by creating a two‐dimensional (2D) map, in which the molecular mass was set along the vertical axis and the retention time along the horizontal axis (Taguchi et al., 2000; Houjou et al., 2005). We normally used the negative ion mode to detect fatty acids as fragment ions from phospholipids by MS/MS, and performed data‐dependent scanning which automatically identifies molecular peaks with high intensity for subsequent product ion scanning. However, even after separation for every class by NPLC before MS, only relatively major molecular species in each class were identified. Thus, a C30 reverse‐phase column was also applied to separate minor molecular species. When this column was used, the influence of the fatty acyl chains was greater than that of the polar head group. Different molecules with the same mass value (i.e., the total number of carbons of the two fatty acyl chains and unsaturated bonds are the same) were detected at different elution times. As a result of application of C30 reverse phase (RP) LC‐MS to top‐down and shotgun lipidomics using data‐ dependent scanning, much better separation of individual molecular species in a phospholipid mixture is obtained than in the case using NPLC. The one advantage of this method is a possibility that unexpected or new lipid metabolites might be detected with this method.

2.3 Focused Lipidomics by Precursor ion Scanning or Neutral Loss Scanning by Mass Spectrometry The second method is applied to detect focused and specified category of lipid metabolites for obtaining more effective and sensitive identification of lipids. For this purpose, selected surveys for precursor ions or neutral losses are performed by the triple quadrupole mass spectrometry. The feature of these methods is to obtain comprehensive detection within the focused categorical metabolites in the samples. We normally performed focused analysis on individual classes of phospholipids, or on phospholipids which contained specific fatty acyl moieties. In addition, we applied an automated search tool named ‘‘Lipid Search’’ (http:// lipidsearch.jp) for their identifications. The comprehensive analysis of lipids by soft ionization is essentially used for a crude lipid mixture containing many different lipid metabolites, and whole molecules existing in the sample were expected to be identified as much as possible. In this case, without preliminary structural information for the metabolites before mass analysis, the significant profiling data can be obtained. But even in this case, some focuses in the molecules are effective to detect important metabolites. For this purpose, a precursor ion scanning method and a neutral loss scanning method are both very important for lipid analyses by MS (Brugger et al., 1997; Lehmann et al., 1997; Ramanadham et al., 1998; Khaselev and Murphy, 2000; Ekroos et al., 2002; Han et al., 2004). These methods are used for comprehensive analysis of the categorical metabolites with structural similarities. The important factor is that by focusing in some limited categories of molecules, a detection limit is greatly enhanced, thus minor but important molecules can be possible to detect. We tried to make up optimal collision conditions for individual molecules to use these methods for the detection of specified class of phospholipids.

2.4 Targeted Methods for Lipidomics by Mass Spectrometry For detecting minor focused molecules, we normally use the second method such as precursor ion scanning or neutral loss scanning without LC separation, and then applied the third method as expanded MRM which can be applied to identify lipid metabolites, structurally related to the targeted molecule, comprehensively at best. We now using this method for detection oxidized phospholipids having specified oxidized fatty acyl chains (in preparation). In addition, this method is applied to analyze detection of molecular species of PI within mature glycosyl‐phosphatidylionositol (GPI) anchored protein as posttranslational lipid structure or their glycolipid precursors (will be reported elsewhere).

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Application Results of Several Mass Spectrometric Methods for Lipidomics

3.1 Application Results of Untargeted and Shotgun Analysis by LC‐MS/MS Phospholipid mixtures were analyzed by LC‐MS/MS with data‐dependent scanning. When using a C30 reverse‐phase column, phospholipids were eluted in order from the hydrophilic molecules to the hydrophobic molecule (> Figure 1-1). In phospholipids, the length and the unsaturated number of fatty acyl chains mainly influenced to the elution order in each molecular species. In this experiment, we used the negative ion mode to detect fatty acids as fragment ions from phospholipids by MS/MS, and performed data‐dependent scanning which automatically identifies molecular peaks with high intensity for subsequent product ion scanning (> Figure 1-2). The ion intensities of each molecular‐related ion were used for quantitative profiling of molecular species of phospholipids after proper compensation by standard phospholipids of same classes with our identification and a profiling tool ‘‘Lipid Search’’ (http://lipidsearch.jp) (> Table 1-1). The description of this search engine will be open to public soon.

. Figure 1‐1 Two-dimensional (2D) map. The 2D-map has the m/z value of [M þ HCO2] ions along the vertical axis and the retention time along the horizontal axis. When using a reverse-phase column, phospholipids elute in order to from the hydrophilic to the more hydrophobic molecules. In phospholipids, the length of fatty acyl chains mainly influence the elution order, i.e., in the order 32:1, 34:1, and 36:1. In addition, the number of double bonds in fatty acyl chains also mainly influence the elution order, i.e., in the order 34:3, 34:2, 34:1, and 34:0. (a) total ion chromatogram, (b) 2D-map (Houjou T et al., 2005. Rapid Commun Mass Spectrom 19: 654-666)

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. Figure 1‐2 MS/MS spectra of [M þ HCO2] ions of PCs from pig liver obtained using RPLC-ESI/MS/MS in data-dependent scanning mode. When different molecules with the same mass value were separated, the MS/MS spectra of the other molecular species are shown with a prime, (a)’, (b)’, etc. (Houjou T et al., 2005. Rapid Commun Mass Spectrom 19: 654-666)

3.2 Application Results of Focused Lipidomics by Mass Spectrometry > Table

1-2 shows m/z values of specific fragment ions and values of neutral loss for precursor ion scanning and neutral loss scanning of polar head groups and fatty acyl chains. Precursor ion scanning and neutral loss scanning were operated on a 4000Qtrap with flow injection at 2 mL/min. Optimum conditions for collision‐induced dissociation (CID) were selected by individual fragments or neutral loss (Larsen et al., 2001; Ekroos et al., 2003; Hsu and Turk, 2003; Wenk et al., 2003). Optimal conditions to detect the proper precursor ions and neutral losses were obtained by MS/MS analyses of each phospholipid class preliminary. Automatic and programmed scanning for each class of phospholipids was operated. In the positive ion mode, precursor ion scanning at m/z 184 was used for choline‐containing phospholipids. Neutral scanning of 141, 185, 189, and 277 Da were used for PE, PS, phosphatidylglycerol (PG), and PI respectively. In the negative ion mode, neutral loss scanning of 60 Da (loss of HCOOþCH3) and 87 Da (loss of serine–H2O) were used for choline‐containing phospholipids and serine‐containing phospholipids, respectively. And precursor ion scanning at m/z 153 and m/z 241 in the negative ion mode were used for glycerol‐containing phospholipids, and inositol‐containing phospholipids, respectively. Quantitative profiling of same phospholipids molecular species by focused methods with specified precursor scanning or neutral loss scanning of head group related mass were also obtained (Houjou et al., 2004, 2005; Ishida et al., 2005a, b; Taguchi et al., 2005). (> Figure 1-3) shows total ion spectrum and a spectrum obtained by neutral loss scanning of 141Da (phosphoryl ethanolamine) of phospholipids extracted from rat spleen in the positive ion mode. More sensitive detection results were obtained in the positive ion mode. > Table 1-3 shows identification results obtained by ‘‘Lipid Search.’’ Major molecular species of PE were effectively identified. (> Figure 1-4) shows a total ion spectrum and spectra of precursor ion scanning and neutral loss scanning of each polar head groups of total phospholipids extracted from THP‐1 cells in the positive ion

Retention time (min) 28.3 45.8 17.4 21.3 21.4 33.2 25.6 37.6 76.7 16.4 16.9 19.2 22.9 29.4 53.8 24.9 18.9 18.6 40.5 17.8 27.9 13.6 35.0 17.6 21.5 21.7 25.1 28.1

Source: Houjou et al. (2005)

828.59

818.58 820.61 826.53

816.57

804.54 806.69 810.55 812.56 814.53

792.62 798.54 800.57

778.56 790.62

774.57 776.55

m/z 764.55

Intensity 117 200 334 184 1897 2515 1592 363 4684 112 685 1172 24008 29028 1090 129 167 293 83 296 2023 232 6550 1066 12237 3140 2010 703

Molecular species (1‐alk‐2‐acyl,18:0‐14:0) (1‐alk‐2‐acyl,16:0‐16:0) (1‐acyl‐2‐acyl,16:1‐16:1) (1‐acyl‐2‐acyl,14:0‐18:1) (1‐acyl‐2‐acyl,16:0‐16:1) (1‐acyl‐2‐acyl,16:0‐16:0) (1‐alk‐2‐acyl,16:0‐18:1) (1‐alk‐2‐acyl,18:1‐16:0) (1‐alk‐2‐acyl,18:0‐16:0) (1‐acyl‐2‐acyl,14:0‐20:4) (1‐acyl‐2‐acyl,16:1‐18:2) (1‐acyl‐2‐acyl,16:0‐18:3) (1‐acyl‐2‐acyl,16:0‐18:2) (1‐acyl‐2‐acyl,16:0‐18:1) (1‐acyl‐2‐acyl,16:0‐18:0) (1‐alk‐2‐acyl,16:1‐20:4) (1‐alk‐2‐acyl,16:0‐20:4) (1‐alk‐2‐acyl,18:0‐18:3) (1‐alk‐2‐acyl,18:1‐18:2) (1‐alk‐2‐acyl,18:0‐18:2) (1‐alk‐2‐acyl,18:1‐18:1) (1‐alk‐2‐acyl,18:0‐18:1) (1‐alk‐2‐acyl,18:0‐18:0) (1‐acyl‐2‐acyl,18:2‐18:2) (1‐acyl‐2‐acyl,16:0‐20:4) (1‐acyl‐2‐acyl,18:1‐18:2) (1‐acyl‐2‐acyl,16:0‐20:3) (1‐acyl‐2‐acyl,18:0‐18:3)

. Table 1-1 Molecular species of PC mixture from pig liver identified by RPLC‐ESI/MS/MS

880.69 882.60 884.64

858.60 860.58 870.63 878.61

856.60

852.66

840.56 848.59 850.59

832.62

m/z 830.60

Retention time (min) 26.2 31.9 31.9 43.1 44.2 25.8 21.1 16.1 20.8 20.3 22.4 26.1 32.7 25.9 27.0 29.3 35.2 67.4 19.2 11.5 28.4 31.5 38.0 31.0 Intensity 2586 15362 15362 179 23995 1054 204 1034 531 1708 1998 562 22888 119 92 3191 261 124 95 99 623 1348 742 123

Molecular species (1‐acyl‐2‐acyl,18:0‐18:2) (1‐acyl‐2‐acyl,16:0‐20:2) (1‐acyl‐2‐acyl,18:1‐18:1) (1‐acyl‐2‐acyl,16:0‐20:1) (1‐acyl‐2‐acyl,18:0‐18:1) (1‐alk‐2‐acyl,18:0‐20:4) (1‐acyl‐2‐acyl,18:0‐20:7) (1‐acyl‐2‐acyl,16:0‐22:6) (1‐acyl‐2‐acyl,18:2‐20:4) (1‐acyl‐2‐acyl,18:1‐20:4) (1‐acyl‐2‐acyl,16:0‐22:5) (1‐acyl‐2‐acyl,18:0‐20:5) (1‐acyl‐2‐acyl,18:0‐20:4) (1‐acyl‐2‐acyl,16:0‐22:3) (1‐acyl‐2‐acyl,18:1‐20:2) (1‐acyl‐2‐acyl,18:0‐20:3) (1‐acyl‐2‐acyl,18:0‐20:2) (1‐acyl‐2‐acyl,18:0‐20:1) (1‐acyl‐2‐acyl,20:5‐20:5) (1‐acyl‐2‐acyl,18:1‐22:5) (1‐acyl‐2‐acyl,18:0‐22:6) (1‐acyl‐2‐acyl,18:0‐22:5) (1‐acyl‐2‐acyl,18:0‐22:4) (1‐acyl‐2‐acyl,18:0‐22:3)

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. Table 1-2a Precursor ion scanning and neutral loss scanning of individual classes of phospholipids in the positive and negative ion modes PC PE PS PI PG

Positive Pre‐m/z 184 (phosphoryl choline) N‐loss 141 Da (phosphoryl ethanolamine) N‐loss 185 Da (phosphoryl serine) N‐loss 277 Da (phosphoryl inositol þ NH4) N‐loss 189 Da (phosphoryl glycerol þ NH4)

Negative N‐loss 60 Da (HCOOþCH3) Pre‐m/z 196 (glycero phosphoryl ethanolamine–H2O) N‐loss 87 Da (serine–H2O) Pre‐m/z 241 (phosphoryl inositol–H2O) Pre‐m/z 153 (phosphoryl glycerol–H2O)

Note: Pre, precursor ion; N‐loss, neutral loss

. Table 1-2b The m/z values of carbonic anions for precursor ion scanning of phospholipids Carbonic anions 14:0 16:1 16:0 18:3 18:2 18:1 18:0

Values of m/z 227 253 255 277 279 281 283

Carbonic anions 20:5 20:4 20:3 20:2 20:1 22:6 22:5 22:4

Values of m/z 301 303 305 307 309 327 329 331

Source: Taguchi et al. (2005)

mode using automatic programmed scanning. In the positive ion mode, more sensitive detection of each class of phospholipids was obtained than that in the negative ion mode. (> Figure 1-5) shows a total ion spectrum and spectra of precursor ion scanning of carbonic anions of total phospholipids extracted from THP‐1 cells in the negative ion mode using automatic programmed scanning. Most of the molecular species of phospholipids with indicated fatty acyl chains were selectively identified (data not shown). Because one fatty acyl moiety was selected as a fragment ion, another fatty acyl moiety and the polar head are most often identified from the molecular‐related mass value with a help of other information. As a result of the detection in the search window of ‘‘Lipid Search’’ (http:// lipidsearch.jp) (> Figure 1-6), most probable individual lipid molecular species are indicated as a pair of fatty acyl chains simultaneously. Our search engine “Lipid Search” (http://lipidsearch.jp) is available via the web. We constructed this engine for the identification of individual lipids law mass data through collaboration with Mitsui Knowledge Industry. Our automated search engine can indicate the most probable candidates for each MS data. The database was constructed with theoretical m/z data of molecular weightrelated ions and their fragment ions for each molecular species of phospholipids, fatty acids, glycerolipids, and their metabolites. These databases are theoretically constructed using the fragment data obtained from the commercially available standards. User has to select the proper descriptions such as database, a MS type used, mass tolerance, a positive or negative ionization, an analytical condition in MS, and minimum intensity of MS peaks to be analyzed, in each box of the indicated search condition. This tool was newly revised in September 2007. In this search window, different classes of phospholipids containing specified fatty acyl chains such as an arachidonic acid can also be effectively identified. We also found that neutral loss scanning of fatty acid or carbonyl keten are also very effective to identify glycerolipids with specified fatty acyl chains.

. Figure 1‐3 Detection of phosphatidylethanolamine in the lipid mixture extracted from rat spleen by neutral loss scanning of 141 Da in the positive ion mode. (a) A total ion spectrum of phospholipids extracted from rat spleen in the positive ion mode. (b) A mass spectrum of neutral loss scanning of 141 Da (phosphorylethanolamine). (Taguchi R et al., 2005. J Chromatog B 823: 26-36)

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. Table 1-3 Identification of molecular species of phosphatidylethanolamine by the data from neutral loss scanning Intensity 109,920 75,862 177,432 86,785 225,015 106,054 62,093 52,149 112,740 216,364 892,332 70,242 30,041 57,267 160,293 132,254 108,036

m/z 716.5 718.5 740.5 742.5 744.6 746.6 752.6 754.6 764.5 766.5 768.6 770.6 782.6 790.5 792.6 794.6 796.6

Molecular species PE,1‐acyl,34:2 PE,1‐acyl,34:1 PE,1‐acyl,36:4 PE,1‐acyl,36:3 PE,1‐acyl,36:2 PE,1‐acyl,36:1 PE,1‐alk,38:5 PE,1‐alk,38:4 PE,1‐acyl,38:6 PE,1‐acyl,38:5 PE,1‐acyl,38:4 PE,1‐acyl,38:3 PE,1‐alk,40:4 PE,1‐acyl,40:7 PE,1‐acyl,40:6 PE,1‐acyl,40:5 PE,1‐acyl,40:4

Source: Taguchi et al. (2005) Note: PE phosphatidylethanolamine; alk, alkyl or alkenyl

3.3 Targeted Method using Expanded MRM for Lipidomics A targeted method using expanded MRM was applied to analyze lysophosphatidic acids or oxidative lipids (data not shown). Most popular quantitative methods have been used in metabolic analysis were single ion monitoring (SIM) and MRM. These methods were normally used in combination with HPLC as LC‐MS. The individual molecules were identified from their retention time and m/z value. Further, in the case of MRM, essentially the combination with the detection of precursor ions and major fragment ions were used. Even in this analysis, the ESI makes it possible to detect more than hundred molecules by a single LC run. MRM is commonly used in the quantitative analysis by mass spectrometry. But in MRM analysis, the target molecules to be analyzed are needed to be defined in advance, and the data of their molecular masses and their fragments should be preliminarily required to set the analytical conditions. I think even in this method some level of comprehensive approach can be possible to expand the detecting target to probable molecular masses by using theoretically constructed data sets calculated by their structural similarities. Such approaches were applied for detecting very low amounts of oxidized lipids. With this method, oxidized species of polyunsaturated fatty acids (PUFA) and phospholipids containing these oxidized PUFA ere effectively detected (will be reported elsewhere).

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Future Aspects

In lipidomics, innovations in soft ionization such as ESI, and tandem mass spectrometry make us possible to detect more than hundred molecular species of lipids comprehensively by the combination with LC. The next important factor in lipidomics is construction of search engines with practical and theoretical database to identify individual molecular species of lipids from individual mass data. Concerning the classification and database, a literature (Fahy et al., 2005) on basic agreement in identification (ID) number for lipids was published in combination work of Lipid MAPS (Metabolites and Pathways Strategy) (http://

. Figure 1‐4 Identification of individual molecular species of focused phospholipid classes by precursor ion scanning and neutral loss scanning of their head groups in the positive ion modes. Extracted total lipid mixture from THP-1 cells was subjected to precursor ion scanning of m/z 184 and neutral loss scanning of 141, 185, 189, and 277 Da. (a) EMS mode analysis of total lipids in the positive ion modes. (b) Precursor ion scanning at m/z 184 for PC and SM. (c) Neutral loss scanning of 141 Da for PE. (d) Neutral loss scanning of 185 Da for PS. (e) Neutral loss scanning of 189 Da for PG. (f) Neutral loss scanning of 277 Da for PI. (Taguchi R et al., 2005. J Chromatog B 823: 26–36)

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. Figure 1‐5 Identification of individual molecular species of THP-1 cell phospholipids having focused specified carbonic anions by precursor ion scanning in the negative ion mode. (a) EMS mode analysis of total lipids in the negative ion modes. (b) Precursor ion scanning at m/z 255 for 16:0 fatty acid. (c) Precursor ion scanning at m/z 281 for 18:1 fatty acid. (d) Precursor ion scanning at m/z 283 for 18:0 fatty acid. (e) Precursor ion scanning at m/z 303 for 20:4 fatty acid. (f) Precursor ion scanning at m/z 329 for 22:5 fatty acid. (Taguchi R et al., 2005. J Chromatog B 823: 26–36)

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. Figure 1‐6 A window of “lipid search.” A law mass data obtained from several different kinds of mass spectrometer of Applied Biosystems, Thermo electron, Waters and Shimadzu can be directory analyzed by lipid search on web. (http://lipidsearch.jp)

www.lipidmaps.org) in USA, European Lipid Initiative (ELI) (http://www.lipidomics.net), and The Japanese Conference on The Biochemistry of Lipids (JCBL) (http://lipidbank.jp). In addition, first meeting in mass spectrometry for lipidomics were held in Dresden at May in 2005. There several practical problems other than database in lipidomics were also discussed. Adding to the real database, we think theoretical database constructed by using structural similarities to basic fragmentation pattern of core standards is important. This theoretical database is induced for ‘‘Lipid Search.’’ Intensity data indicated in this chapter were only used for profiling relative intensities of individual molecular species of lipids. In addition, it is necessary to construct a tool for handling automatically a large number of quantitative data obtained from law mass data. But still there are several factors to solve this problem. One is how to compensate individual peaks with different intensities caused from different ionization efficiency. At present, quantitative profiling data within the same group of phospholipids can be rather effectively obtained by using one specific molecular species as an internal standard. But for exact quantitative analysis, several different level of compensation are needed such as compensation of isotope effect, and compensation for differences in ionization caused by differences in head groups, fatty acyl lengths and number of double bonds. Automated handling of these data and practical viewers for profiling data for each molecular

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Advances in lipid analysis/lipidomics – analyses of phospholipids by recent application of mass spectrometry

. Figure 1‐7 Database and analytical tools to be constructed for lipidomics Lipid Search and Lipid Navigator can be accessed by http://lipidsearch.jp) ARM, Atomic reconstruction of metabolism, is organized by Dr. Trita. (http://www. metabolome.jp). Lipid bank is accessed by http://lipidbank.jp, and Lipid base is accessed by http://www. lipidbase.jp

species in different samples are strongly needed for fast analytical process. For further understanding in metabolic alteration within different circumstances, construction of multivariate analytical tool is important. (> Figure 1-7) summarize these basic tools in our group to be constructed for high throughput analyses by mass spectrometry for lipidomics. As future techniques in lipidomics, a flux analysis of lipids by using stable isotope labeling and analysis of localization of lipids by imaging MS should be important for clarifying the physiological function and changes of individual lipid species in local.

Acknowledgment This work was supported by special Coordination fund from the Ministry of Education, Culture, Sports, Science and Technology of the Japanese Government, and a fund from Core Research for Evolutional and Technology (CREST) of Japan Science and Technology Agency (JST).

References Di Paolo G, Moskowitz HS, Gipson K, Wenk MR, Voronov S, Obayashi M, Flavell R, Fitzsimonds RM, Ryan TA, De Camilli P. 2004. Impaired PtdIns(4,5)P2 synthesis in nerve terminals produces defects in synaptic vesicle trafficking. Nature 431: 415-422. Domingues P, Domingues MR, Amado FM, Ferrer‐ Correia AJ. 2001. Characterization of sodiated glycerol

phosphatidylcholine phospholipids by mass spectrometry. Rapid Commun Mass Spectrom 15: 799-804. Ekroos K, Chernushevich LV, Simons K, Shevchenko A. 2002. Quantitative profiling of phospholipids by multiple precursor ion scanning on a hybrid quadrupole time‐of‐flight mass spectrometer. Anal Chem 74: 941-949.

Advances in lipid analysis/lipidomics – analyses of phospholipids by recent application of mass spectrometry Ekroos K, Ejsing CS, Bahr U, Karas M, Simons K, et al. 2003. Charting molecular composition of phosphatidylcholines by fatty acid scanning and ion trap MS3 fragmentation. J Lipid Res 44: 2181-2192. Fahy E, Subramaniam S, Brown HA, Glass CK, Merrill AH Jr, et al. 2005. A comprehensive classification system for lipids. J Lipid Res 46: 839-861. Fenn JB, Mann M, Meng CK, Wong SF, Whitehouse CM. 1989. Electrospray ionization for mass spectrometry of large biomolecules. Science 246: 64-71. Fridriksson EK, Shipkova PA, Sheets ED, Holowka D, Baird B, et al. 1999. Quantitative analysis of phospholipids in functionally important membrane domains from RBL‐2H3 mast cells using tandem high‐resolution mass spectrometry. Biochemistry 38: 8056. Han X, Gross RW. 1994. Electrospray ionization mass spectroscopic analysis of human erythrocyte plasma membrane phospholipids. Proc Natl Acad Sci USA 91: 10635-10639. Han X, Gross RW. 2005. Shotgun lipidomics: Electrospray ionization mass spectrometric analysis and quantitation of cellular lipidomes directly from crude extracts of biological samples. Mass Spectrom Rev 24: 367-412. Han X, Yang J, Cheng H, Ye H, Gross RW. 2004. Toward fingerprinting cellular lipidomes directly from biological samples by two‐dimensional electrospray ionization mass spectrometry. Anal Biochem 330: 317-331. Heller DN, Murphy CM, Cotter RJ, Fenselau C, Uy OM. 1988. Constant neutral loss scanning for the characterization of bacterial phospholipids desorbed by fast atom bombardment. Anal Chem 60: 2787-2791. Houjou T, Yamatani K, Nakanishi H, Imagawa M, Shimizu T, et al. 2004. Rapid and selective identification of molecular species in phosphatidylcholine and sphingomyelin by conditional neutral loss scanning and MS3. Rapid Commun Mass Spectrom 18: 3123-3130. Houjou T, Yamatani K, Nakanishi H, Imagawa M, Shimizu T, et al. 2005. A shotgun tandem mass spectrometric analysis of phospholipids with normal‐phase and/or reverse‐phase liquid chromatography/electrospray ionization mass spectrometry. Rapid Commun Mass Spectrom 19: 654-666. Hsu FF, Turk J. 2003. Electrospray ionization/tandem quadrupole mass spectrometric studies on phosphatidylcholines: The fragmentation processes. J Am Soc Mass Spectrom 14: 352-363. Ishida M, Imagawa M, Shimizu T, Taguchi R. 2005a. Specific detection of lysophosphatidic acids in serum extracts by tandem mass spectrometry. J Mass Spectrom Soc Jpn 53: 25-32. Ishida M, Imagawa M, Shimizu T, Taguchi R. 2005b. Effective Extraction and analysis for lysophosphatidic acids and their precursors in human plasma usng electrospray ionization mass spectrometry. J Mass Spectrom Soc Jpn 53: 217-226.

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Ishida M, Yamazaki T, Houjou T, Imagawa M, Harada A, et al. 2004. High‐resolution analysis by nano‐electrospray ionization Fourier transform ion cyclotron resonance mass spectrometry for the identification of molecular species of phospholipids and their oxidized metabolites. Rapid Commun Mass Spectrom 18: 2486-2494. Ivanova PT, Cerda BA, Horn DM, Cohen JS, McLafferty FW, et al. 2001. Electrospray ionization mass spectrometry analysis of changes in phospholipids in RBL‐2H3 mastocytoma cells during degranulation. Proc Natl Acad Sci USA 98: 7152-7157. Jones JJ, Stump MJ, Fleming RC, Lay JO Jr, Wilkins CL. 2003. Investigation of MALDI‐TOF and FT‐MS techniques for analysis of Escherichia coli whole cells. Anal Chem 75: 1340-1347. Kerwin JL, Tuininga AR, Ericsson LH. 1994. Identification of molecular species of glycerophospholipids and sphingomyelin using electrospray mass spectrometry. J Lipid Res 35: 1102-1114. Khaselev N, Murphy RC. 2000. Electrospray ionization mass spectrometry of lysoglycerophosphocholine lipid subclasses. J Am Soc Mass Spectrom 11: 283-291. Lehmann WD, Koester M, Erben G, Keppler D. 1997. Characterization and quantification of rat bile phosphatidylcholine by electrospray‐tandem mass spectrometry. Anal Biochem 246: 102-110. Marto JA, White FM, Seldomridge S, Marshall AG. 1995. Structural characterization of phospholipids by matrix‐assisted laser desorption/ionization Fourier transform ion cyclotron resonance mass spectrometry. Anal Chem 67: 3979-3984. Nor Aliza AR, Bedick JC, Rana RL, Tunaz H, Hoback WW, et al. 2001. Arachidonic and eicosapentaenoic acids in tissues of the firefly, Photinus pyralis (Insecta: Coleoptera). Comp Biochem Physiol A Mol Integr Physiol 128: 251-257. Pulfer M, Murphy RC. 2003. Electrospray mass spectrometry of phospholipids. Mass Spectrom Rev 22: 332-364. Ramanadham S, Hsu FF, Bohrer A, Nowatzke W, Ma Z, et al. 1998. Electrospray ionization mass spectrometric analyses of phospholipids from rat and human pancreatic islets and subcellular membranes: Comparison to other tissues and implications for membrane fusion in insulin exocytosis. Biochemistry 37: 4553-4567. Rugger B, Erben G, Sandhoff R, Wieland FT, Lehmann WD. 1997. Quantitative analysis of biological membrane lipids at the low picomole level by nano‐electrospray ionization tandem mass spectrometry. Proc Natl Acad Sci USA 94: 2339-2344. Taguchi R, Hayakawa J, Takeuchi Y, Ishida M. 2000. Two‐ dimensional analysis of phospholipids by capillary liquid chromatography/electrospray ionization mass spectrometry. J Mass Spectrom 35: 953-966.

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Taguchi R, Houjou T, Nakanishi H, Yamazaki T, Ishida M, et al. 2005. Focused lipidomics by tandem mass spectrometry. J Chromatogr B Anal Technol Biomed Life Sci 823: 26-36. Tserng KY, Griffin R. 2003. Quantitation and molecular species determination of diacylglycerols, phosphatidylcholines, ceramides, and sphingomyelins with gas chromatography. Anal Biochem 323: 84-93.

Wenk MR, Lucast L, Di Paolo G, Romanelli AJ, Suchy SF, et al. 2003. Phosphoinositide profiling in complex lipid mixtures using electrospray ionization mass spectrometry. Nat Biotechnol 21: 813-817. Yokoyama K, Shimizu F, Setaka M. 2000. Simultaneous separation of lysophospholipids from the total lipid fraction of crude biological samples using two‐dimensional thin‐layer chromatography. J Lipid Res 41: 142-147.

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Choline and Ethanolamine Glycerophospholipids

A. A. Farooqui . L. A. Horrocks . T. Farooqui

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22

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Biosynthesis of Choline and Ethanolamine Glycerophospholipids in Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23

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Incorporation of Choline and Ethanolamine Glycerophospholipids into Neural Membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25

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Effects of Structural Variation of Choline and Ethanolamine Glycerophospholipids on Neural Membrane Structure and Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26

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Catabolism of Choline and Ethanolamine Glycerophospholipids in Brain . . . . . . . . . . . . . . . . . . . . . . 27

6 Roles of Choline and Ethanolamine Glycerophospholipids in Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30 6.1 Choline and Ethanolamine Glycerophospholipids Provide Precursors for Second Messengers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30 6.2 PLA2, PLC, and PLD-Generated Second Messengers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30 6.3 Plasmalogens as Antioxidants and Membrane Fusion Molecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32 6.4 Choline and Ethanolamine Glycerophospholipids in Apoptotic Cell Death . . . . . . . . . . . . . . . . . . . . . . . 32 6.5 PAF and its Role in Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 6.6 Modulation of Enzymic Activities by Choline and Ethanolamine Glycerophospholipids . . . . . . . . . 33 7

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_2, # Springer ScienceþBusiness Media, LLC 2009

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Abstract: Choline and ethanolamine glycerophospholipids are amphipathic molecules that are asymmetrically distributed in the bilayer. They provide the neural membranes with a suitable environment, fluidity, and ion permeability. The degree of saturation and the length of glycerophospholipid-acyl chains are important determinants of neural membrane properties. Choline and ethanolamine glycerophospholipids are synthesized at the endoplasmic reticulum and are transported to other membranous structures by phospholipid exchange and transfer proteins. Glycerophospholipids undergo base-exchange, methylation, and decarboxylation reactions for interconversion. These reactions and activities of phospholipases A2, C, and D are involved in the turnover, compositional maintenance, and rearrangements of glycerophospholipids in membranes. Glycerophospholipids are a storage depot for precursors for second messengers, and may be involved in membrane fusion, apoptosis, and regulation of the activities of membrane-bound enzymes and ion-channels. List of Abbreviations: AA, arachidonic acid; cAMP, cyclic adenosine monophosphate; CDP, cytidinediphospho; CTP, cytidine triphosphate; DAG, 1,2-sn-diacylglycerols; DHA, docosahexaenoic acid; ER, endoplasmic reticulum; FFA, free fatty acids; NFκB, nuclear transcription factor κD; NPDR, neuroprotectin D receptor; PAF, platelet-activating factor; PKC, protein kinase C; PtdCho, phosphatidylcholine; PtdEtn, phosphatidylethanolamine; PtdH, phosphatidic acid; PLA, phospholipase A; PLC, phospholipase C; PLD, phospholipase D; RER1, resolvin E receptor; ResoDR1, resolvin D receptor; ResoER1, resolvin E receptor; TNF-α, tumor necrosis factor-α

1

Introduction

Choline and ethanolamine glycerophospholipids (also known as phosphoglycerides) constitute a biologically important group of molecules that along with serine and inositol glycerophospholipids, sphingomyelins, glycolipids, and cholesterol form the backbone of neural membranes. In neural membranes, these glycerophospholipids are organized in bilayers and held together by hydrophobic, coulombic, and van der Waal forces and hydrogen bonds (Farooqui et al., 2000b; Ivanova et al., 2004). In phospholipid bilayers, the polar regions orient toward the aqueous phase with the hydrophobic regions sequestered away from water. These regions interact with each other, creating the essential milieu of the membrane. The distribution of choline and ethanolamine glycerophospholipids is normally asymmetric across the plane of the plasma membrane. Less fluid glycerophospholipids such as ethanolamine glycerophospholipids and serine glycerophospholipids concentrate in the inner leaflet and more fluid glycerophospholipids such as choline glycerophospholipids and sphingomyelin concentrate in the outer leaflet (Farooqui et al., 2000b; Tillman and Cascio, 2003). This distribution of the neural membrane glycerophospholipids is quite stable. When the glycerophospholipids are redistributed so that ethanolamine glycerophospholipids or serine glycerophospholipids are in the outer leaflet of the bilayer, an aminophospholipid translocase (flippase) restores the normal phospholipid distribution (Pomorski et al., 2004). The differential packing of phospholipids, glycolipids, cholesterol, and proteins leads to the formation of microdomains, which can diffuse laterally. These microdomains (lipid rafts) serve as mobile platforms for signal transduction. They cluster and organize bilayer constituents including receptors, enzymes, and ion channels that protrude differentially through the membrane or are localized predominantly on the intracellular or extracellular membrane surface. In neural membranes, normal glycerophospholipid homeostasis is balanced between glycerophospholipid catabolism and resynthesis through the reacylation/deacylation cycle and de novo synthesis pathways (Farooqui et al., 2000a, b). The interaction of an agonist with its receptor results in the enhancement of glycerophospholipid metabolism. This not only regulates the activities of membrane-bound enzymes and ion channels, but also modulates many physicochemical properties of neural membranes such as fluidity, lateral pressure profile, bilayer thickness, and permeability (Farooqui et al., 2000b; Tillman and Cascio, 2003). Choline and ethanolamine glycerophospholipids also play a key role in signal transduction pathways. Lipid mediators generated from these glycerophospholipids transduce signals from the surface of the neural cell to the interior, influencing intracellular metabolism, ion transport, and gene expression. Because of the remarkable importance of choline and ethanolamine glycerophospholipids in membranes of the brain, it is crucial for neural cells to maintain and preserve the content and composition of their choline and

Choline and ethanolamine glycerophospholipids

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ethanolamine glycerophospholipids (Farooqui et al., 2000a). This provides neural membranes with structural and functional integrity that facilitates appropriate interactions with integral membrane proteins. They not only modulate the regular function of the membranes, but also control adaptive responses (Ivanova et al., 2004). Neural membranes are not simply inert physical barriers, but are complex, wellorganized, and highly specialized structures involved in receiving, processing, transporting, and transmitting information from one cell to another. Two unresolved fundamental questions related to choline and ethanolamine glycerophospholipid metabolism in neural membranes still remain unanswered. First, what mechanisms regulate the composition and total content of choline and ethanolamine glycerophospholipids in membranes of the neurons, astrocytes, oligodendrocytes, and microglia? To respond to this question, one must have complete knowledge of the biosynthesis of each molecular species of choline and ethanolamine glycerophospholipids in various pools of brain tissue. Second, how do choline and ethanolamine glycerophospholipids move between the membranes of different subcellular organelles in neurons, astrocytes, and oligodendrocytes? For example, the synthesis of choline and ethanolamine glycerophospholipids largely occurs in endoplasmic reticulum (ER). The membranes of other subcellular organelles are not capable of synthesizing their own choline and ethanolamine glycerophospholipids. Therefore, the intracellular transport and sorting of glycerophospholipids from the site of synthesis (ER) to their final destination is an essential event in glycerophospholipid metabolism. Although several mechanisms have been proposed for glycerophospholipid transport and sorting including carrier proteins, transport vesicles, and contact zones between donor and acceptor membranes (Valle´e et al., 1999; Oram et al., 2003; Voelker, 2003), the specificity and modulation of mechanisms associated with transport processes and their regulation by specific genes remain unknown (Van Meer and Sprong, 2004). The purpose of this study is to examine the metabolism and role of choline and ethanolamine glycerophospholipids and their lipid mediators in brain. With the development of lipidomics (Forrester et al., 2004), this discussion should initiate more studies on specific genes involved in the regulation of choline and ethanolamine glycerophospholipid synthesis and also on genes related to the sorting, transport, and modulation of second messenger generation from choline and ethanolamine glycerophospholipids in neural membranes.

2

Biosynthesis of Choline and Ethanolamine Glycerophospholipids in Brain

Choline and ethanolamine glycerophospholipids are synthesized mainly via the CDP–choline or CDP–ethanolamine pathways (Kennedy cycle). This pathway involves three enzymic steps catalyzed by choline or ethanolamine kinases, choline or ethanolamine phosphate cytidylyltransferases, and CDP–choline or CDP–ethanolamine:1,2-diacylglycerol choline or ethanolamine phosphotransferases (Kent and Carman, 1999; Kent, 2005) (> Figure 2-1). These enzymes have intracellular localization: choline or ethanolamine kinase is localized in the cytosol; cytidylyltransferases are distributed between the cytosol and membrane fractions, and phosphotransferases are integral membrane proteins that are predominantly present in endoplasmic reticulum (Golfman et al., 2001; Bleijerveld et al., 2004). Among these, the cytidylyltransferase reaction is the rate-limiting step for the CDP–choline or CDP–ethanolamine pathways. Cytidylyltransferases are regulated by a novel mechanism that involves translocation of the enzyme between the cytosol and endoplasmic reticulum. The cytosolic cytidylyltransferase is inactive until its translocation to the endoplasmic reticulum resulting in its activation (Clement and Kent, 1999; Kent and Carman, 1999). The phosphorylation of a cytidylyltransferase by cAMP-dependent kinase releases the enzyme from the membrane and renders it inactive (Carter et al., 2003). Subsequent dephosphorylation of the cytidylyltransferase results in its binding to endoplasmic reticulum membrane and renders it active (Carman and Kersting, 2004). All these enzymes have been purified, characterized, and cloned from several nonneural sources (Sugimoto et al., 2003; Banchio et al., 2004; Vance and Vance, 2004). Choline or ethanolamine phosphotransferases are localized in the endoplasmic reticulum (Wright and McMaster, 2002). They catalyze the transfer of phosphocholine or phosphoethanolamine to 1,2-diacylglycerol from CDP–choline or CDP–ethanolamine, with the release of CMP. Under physiological

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Choline and ethanolamine glycerophospholipids

. Figure 2-1 Reactions involved in the biosynthesis of glycerophospholipids. Glycerol-3-phosphate acyltransferase (1); lysophosphatidic acid acyltransferase; ethanolamine kinase (3); ethanolamine cytidylyltransferase (4); CDP–phosphoethanolamine cytidylyltransferase (5); phosphatidic acid phosphatase (6); CDP–phosphocholine cytidylyltransferase (7); phosphatidylethanolamine methyltransferase (8); phosphatidylserine decarboxylase (9); phosphatidylserine synthase (10); phospholipase A2 (11); phospholipase C (12); phospholipase D (13); PAFsynthesizing enzymes (14); ethanolamine glycerophospholipid (EtnGpl); choline glycerophospholipid (ChoGpl); serine glycerophospholipid (SerGpl); choline lysoglycerophospholipid (lyso-ChoGpl); phosphatidic acid (PtdH); diacylglycerol (DAG); free fatty acid; and protein kinase C (PKC)

conditions, the synthesis of choline or ethanolamine glycerophospholipids is favored as a result of the very rapid rephosphorylation of CMP to CTP, which requires ATP. Based on topographical studies, the choline phosphotransferase (Henneberry and McMaster, 1999) is likely located on the outer leaflet and the ethanolamine phosphotransferase is situated on the inner leaflet or it has transmembrane localization in the microsomal vesicle. Choline glycerophospholipids are also synthesized by the repeated methylation of ethanolamine glycerophospholipids by S-adenosylmethionine (S-AdoMet) (Shields et al., 2001). PtdEtn N-methyltransferase activity is associated with endoplasmic reticulum and mitochondria. Purification and characterization of two methyltransferases have been reported from rat liver (Shields et al., 2001). A minor pathway for the synthesis of PtdEtn involves the decarboxylation of serine glycerophospholipid. This enzyme has been purified, characterized, and cloned. Choline and ethanolamine glycerophospholipids contain more than one kind of fatty acid per molecule, so that a given class of these glycerophospholipids from any tissue actually represents a family of molecular species (DeLong et al., 1999; Farooqui et al., 2002). Choline glycerophospholipids contain palmitic acid or stearic acid at the sn-1 position and unsaturated fatty acids, such as arachidonic acid, oleic acid, linoleic acid, or linolenic acid at the sn-2 position of glycerol moiety. Ethanolamine glycerophospholipids contain palmitic acid or stearic acid at the sn-1 position with long-chain polyunsaturated fatty

Choline and ethanolamine glycerophospholipids

2

acids (arachidonic, oleic, or docosahexaenoic acid) at the sn-2 position. The base-exchange reaction between ethanolamine glycerophospholipids and serine is initiated by attack on the phosphodiester bond of ethanolamine glycerophospholipid or choline glycerophospholipid by the hydroxyl group of serine. There are differences between choline glycerophospholipids generated by the Kennedy pathway and by the methylation of the ethanolamine glycerophospholipids (DeLong et al., 1999). Choline glycerophospholipids produced from the Kennedy pathway mainly contain medium chain, saturated (e.g., 16:0/18:0), or monounsaturated (e.g., 16:0/18:1) species. In contrast, the methylation pathway generates much more diverse glycerophospholipids with significantly more long-chain polyunsaturated (18:0/20:4) fatty acid containing species. In mitochondrial membranes, choline glycerophospholipids are the major substrates for the synthesis of serine glycerophospholipids. Since there is no net change in the number or kind of bonds, this reaction is reversible and energy-independent. The base-exchange reaction is catalyzed by Ca2þ-dependent phosphatidylserine synthases I and II, which are localized in endoplasmic reticulum (Vance and Vance, 2004). Another class of choline or ethanolamine glycerophospholipids is represented by plasmalogens and platelet-activating factor (PAF) (Maclennan et al., 1996; Farooqui and Horrocks, 2001; Nagan and Zoeller, 2001). In contrast to other choline or ethanolamine glycerophospholipids, plasmalogens contain a vinyl ether linkage rather than an ester linkage at the sn-1 position of the glycerol moiety. Plasmalogens are synthesized from dihydroxyacetone phosphate, which combines with acyl-CoA to form 1-acyldihydroxyacetone phosphate (Lee, 1998). This reaction is catalyzed by acyl-CoA: dihydroxyacetone phosphate acyltransferase. An exchange reaction between the acyl group and the long-chain alcohol produces 1-Oalkyldihydroxyacetone phosphate, which in the presence of NADPH is converted to 1-O-alkylglycerol 3-phosphate. After acylation at the sn-2 position, the resulting 1-O-alkyl-2-acyl-sn-glycerol is hydrolyzed by 1-alkyl-2-acyl-sn-glycerol 3-phosphate phosphohydrolase. In the last step, the action of alkyl-2-acyl sn-glycerol choline or ethanolamine phosphotransferase in the presence of CDP–choline or CDP– ethanolamine results in the formation of plasmalogens (Lee, 1998; Nagan and Zoeller, 2001). PAF is a unique glycerophospholipid with lipid mediator properties (Maclennan et al., 1996; Ishii et al., 2002). PAF synthesis takes place either via the de novo pathway, which involves the transfer of phosphocholine from CDP–choline to 1-O-alkyl 2-acetyl-sn-glycerol, or via the remodeling pathway. In this pathway, 1-O-alkyl-2-acyl-sn-glycero-3-phosphocholine, present in membranes, is hydrolyzed by a phospholipase A2 generating 1-O-alkyl-2-lyso-sn-glycero-3-phosphocholine (lyso-PAF), which is then acetylated to PAF by an acetyltransferase. Another pathway for the synthesis of PAF is the oxidative fragmentation of choline glycerophospholipids (Farooqui and Horrocks, 2004b). All these pathways collectively modulate the levels of PAF under normal and pathological conditions.

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Incorporation of Choline and Ethanolamine Glycerophospholipids into Neural Membranes

The de novo synthesis of glycerophospholipids occurs at the endoplasmic reticulum. The newly synthesized glycerophospholipids self-assemble into thermodynamically stable bilayer vesicles, which detach from the endoplasmic reticulum and travel to other sites for donation of their glycerophospholipids to other membranous structures. This process involves spontaneous transfer of glycerophospholipids to other membranes and transport of glycerophospholipid molecules by glycerophospholipid transfer proteins. Some glycerophospholipid transfer proteins are specific and others are not. Multiple vesicular carriers with distinct mechanisms exist for the transfer of glycerophospholipids between distinct subcellular compartments. Studies dealing with the transfer of glycerophospholipids from the endoplasmic reticulum to the mitochondria also indicate the importance of the spatial organization of the endoplasmic reticulum and the existence of a specific proximity between various organelles (Kent and Carman, 1999). Other important factors that affect the transfer of glycerophospholipid to other membranes are the occurrence of specific membrane domains and the sorting mechanism for glycerophospholipids (Birner and Daum, 2003). The active transfer of glycerophospholipids between outer and inner leaflets occurs against electrical and concentration gradients by an enzymic mechanism (aminophospholipid transferase). This process uses

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Choline and ethanolamine glycerophospholipids

ATP to overcome the gradients. Nonenzymic movement of glycerophospholipids from one side to the other (flip-flop movement) also occurs, but it is slow and is measured in days or weeks. Thus, the distribution of glycerophospholipids in membranes is regulated not only by the activities of enzymes involved in their metabolism but also by the transport and incorporation processes into the membrane. Very little is known about intracellular transport of choline and ethanolamine glycerophospholipids in neurons. Pulse-chase radiolabeling studies in explant cultures of rat superior cervical ganglia and rat sciatic nerve indicate that both anterograde and retrograde transports of choline and ethanolamine glycerophospholipid vesicles occur between the cell body and distal axons (Negretti et al., 2000). The rate of anterograde transport of newly synthesized glycerophospholipids from cell bodies to axon was approximately several hundred millimeters per day. For axonal elongation, synthesis of choline and ethanolamine glycerophospholipids occurs in distal axons with some glycerophospholipid transport from axons to myelin membrane. Most of the glycerophospholipids in myelin are synthesized in the oligodendroglia. Very little is known about the genes that modulate biosynthesis, transport, and sorting of glycerophospholipids in various subcellular organelles. In the last decade, gene-targeted mice with defective glycerophospholipid synthesis and transport have been used to understand metabolic insight into CTP: phosphocholine cytidylyltransferase genes (Pcyt 1a and Pcyt 1b), the ethanolamine glycerophospholipid N-methyltransferase gene (Pemt), and changes in choline and ethanolamine glycerophospholipids in nonneural tissues (Watkins et al., 2003; Zhu et al., 2003; Vance and Vance, 2005). Gene-targeted mice can survive without certain glycerophospholipid biosynthesis genes; in each case, an alternative pathway or enzyme exists for synthesizing that glycerophospholipid (Vance and Vance, 2005). Studies on Pcyt 1a, Pcyt 1b, and Pemt genes in mouse brain have not been performed.

4

Effects of Structural Variation of Choline and Ethanolamine Glycerophospholipids on Neural Membrane Structure and Function

Choline and ethanolamine glycerophospholipids perform important functions in membranes. Certain sets of glycerophospholipids are selected for each membrane to give it the unique characteristics suited to its role. These characteristics include fluidity, permeability, local curvature, molecular packing or hydration, charge, and reactivity to support activities of membrane-bound enzymes and ion channels (Farooqui et al., 2000b). These characteristics are not the properties of individual glycerophospholipid classes, but those of an organized membrane as a whole. As all membranes possess a typical composition with more or less the same classes of glycerophospholipids, it is the ratio between these classes and their molecular species that are unique and provide specific characteristics to membranes from different organelles (Tillman and Cascio, 2003). In addition, most membranes from different organelles have some glycerophospholipidsynthesizing activities (interconversion reactions). This results in differences in glycerophospholipid composition. Studies on the composition of glycerophospholipids in various membranes are still in a developing state. In order to reconstruct a membrane composition that is found in vivo under physiological conditions, one has to determine the lipid composition expressed in terms of mole per surface area for a specific membrane in a particular subcellular organelle of a specific tissue. Moreover, studies to date have been performed mostly on membrane fractions that are contaminated with membranes of one organelle with another (Farooqui et al., 2000b). The head group determines the surface charge on glycerophospholipids: serine and inositol glycerophospholipids are strongly anionic, ethanolamine glycerophospholipids are slightly anionic, and choline glycerophospholipids and sphingomyelins are zwitterionic at neutral pH. The ratio of the strongly anionic to zwitterionic glycerophospholipids varies widely between cell types, but is usually constant for a particular cell type in different species. Cations like Ca2þ and Mg2þ bind to anionic head groups at the inner half of the lipid bilayer and can increase bilayer rigidity and induce lateral segregation of glycerophospholipids. Membranes therefore may act as a sink for these cations, which can be released as a result of membrane perturbation (Farooqui et al., 2000b). This process may be involved in many disease processes that are characterized by alterations in the properties of membranes and the levels of cations, such as Ca2þ, Mg2þ, and Fe3þ.

Choline and ethanolamine glycerophospholipids

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Catabolism of Choline and Ethanolamine Glycerophospholipids in Brain

Brain tissue actively catabolizes choline and ethanolamine glycerophospholipids. Each portion of the glycerophospholipid molecule turns over at a different rate, including the phosphate group, the nitrogenous base, and the acyl groups at the sn-1 and sn-2 positions (Porcellati, 1983). Glycerophospholipids are hydrolyzed by a group of enzymes called phospholipases. Thus, phospholipase A1 (PLA1) catalyzes the hydrolysis of ester bond at the sn-1 position forming free fatty acids and 2-acyl lysoglycerophospholipids. Phospholipase A2 (PLA2) acts on the fatty acid ester bond at the sn-2 position liberating free fatty acids and 1-acyl lysoglycerophospholipids that in turn can be acylated by acyl-CoA in the presence of an acyltransferase (deacylation/reacylation cycle). Alternatively, 1-acyl lysoglycerophospholipids can be hydrolyzed by a lysophospholipase forming fatty acids and the glycerophosphobase. Phospholipase C (PLC) hydrolyzes the phosphodiester bond at the sn-3 position of choline glycerophospholipids forming 1,2-diacylglycerols and phosphocholine. Finally, phospholipase D (PLD) cleaves glycerophospholipids into phosphatidic acid (PtdH) and freebase (Farooqui et al., 2000b). Phospholipases A1, A2, C, and D have been purified and characterized from brain tissue (Hirashima et al., 1992; Ross et al., 1995; Negre-Aminou et al., 1996; Exton, 1997; Banno, 2002; Fukami, 2002; Strokin et al., 2003; Zhang et al., 2004; Jenkins and Frohman, 2005). All these enzymes occur in brain tissue in multiple forms and are linked to various receptors such as glutamate receptors, dopamine receptors, cytokine receptors, and growth factor receptors (Attucci et al., 2001; Shen et al., 2001; Vitale et al., 2004; Farooqui et al., 2006). Many isoforms of PLC and PLD have been cloned from brain (Banno, 2002; Fukami, 2002; Stillwell and Wassall, 2003; McDermott et al., 2004). PLA1 and PLA2 from brain have not been cloned. The properties of PLA2, PLC, and PLD are shown in > Table 2-1. Neural membrane choline and ethanolamine glycerophospholipids are hydrolyzed by multiple forms of PLA2 activities (Farooqui and Horrocks, 2004a). Lysoglycerophospholipids that are generated by the action of PLA1 and PLA2 are either hydrolyzed by lysophospholipase or used to regenerate glycerophospholipids in the reacylation/deacylation cycle (Farooqui et al., 2000a). Plasmalogens are hydrolyzed by . Table 2-1 Substrate Specificity, Molecular Mass, and Effect of Calcium on Brain Phospholipases A1, A2, C, D, and Lysophospholipases from Brain Molecular mass (kDa) 112 95 200–500 100 88 40

Effect of calcium Stimulated Stimulated Inhibited Stimulated No effect No effect

Enzyme PLA1 PLA1 cPLA2

Substrate Choline glycerophospholipid

iPLA2 PlsEtn-PLA2

Choline glycerophospholipid Ethanolamine glycerophospholipid

PtdCho-PLC PLD-I

Choline glycerophospholipid Choline glycerophospholipid

62–65 124

Stimulated Stimulated

PLD-II Lysophospholipase

Choline glycerophospholipid Lysocholine glycerophospholipid Lysocholine glycerophospholipid

106 95

No effect No effect

References Pete et al. (1994) Pete et al. (1994) Yoshihara and Watanabe (1990) Yang et al. (1999) Mouton and Arendash (1990); Farooqui and Horrocks (2001) Fukami (2002) Klein et al. (1995); Exton (1997); McDermott et al. (2004) Exton (1997) Pete and Exton (1996)

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No effect

Farooqui et al. (1985)

Lysophospholipase

Choline glycerophospholipid

27

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plasmalogen-selective PLA2 and lysoplasmalogenase (Farooqui and Horrocks, 2001, 2004b). Choline and ethanolamine glycerophospholipids are also hydrolyzed by multiple forms of PLC and PLD (Banno, 2002; Fukami, 2002), generating many molecular species of diacylglycerol (DAG). Complexities of choline and ethanolamine glycerophospholipid degradation by multiple forms of PLC and PLD become obvious when one considers the participation and stimulation of as many as 12 distinct protein kinase C (PKC) isoforms grouped into three subfamilies by the different molecular species of DAG (Cook et al., 1998; Morishita et al., 2005). PKC-mediated protein phosphorylation is one of the most versatile posttranslational modifications used by neural cells. It plays a crucial role in the continuous remodeling of different transcriptional regulators. It remains to be seen whether or not DAG derived from the PLD-mediated hydrolysis of choline glycerophospholipids can activate PKC isoforms similarly to the DAG derived from inositol glycerophospholipids. In vitro studies indicate that the DAG derived from inositol glycerophospholipids is more effective in stimulating PKC isoforms than the DAG derived from the hydrolysis of choline glycerophospholipids (Marignani et al., 1996). During signal transduction, the transient generation of lipid mediators, such as DAG, arachidonic acid, and eicosanoids, involves a small portion of total neural membrane lipid. Regeneration of the hydrolyzed glycerophospholipids via the reacylation/deacylation cycle is necessary, not only for maintaining membrane integrity, but also for restoring future participation of neural membranes in signal transduction (Farooqui et al., 2000a). Multiple forms of PLA2, C, and D are part of a signal transduction network and cross-talk between receptor-regulated effector systems through the generated second messengers (> Figure 2-2) that is essential

. Figure 2-2 Diagram showing the receptor-mediated degradation of choline and ethanolamine glycerophospholipids by phospholipase A2 (PLA2), phospholipase C (PLC), and phospholipase D (PLD). A, agonist; R, receptor; phosphatidic acid (PtdH); protein kinase C (PKC); arachidonic acid (AA); docosahexaenoic acid (DHA), and plateletactivating factor (PAF)

Choline and ethanolamine glycerophospholipids

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for maintaining normal neuronal and glial cell growth (Farooqui et al., 1992). Suggestions on cross-talk among multiple forms of intracellular PLA1, A2, C, and D are supported by the observation that several forms of these enzymes are stimulated by the same agonist and the products of one phospholipase can participate in the activation of others. Thus, the stimulation of PLC generates DAG, which translocates and activates PKC-a and PKC-b leading to the activation of multiple forms of both PLA2 and PLD (Di Marzo et al., 1991). The action of DAG-lipase on DAG generates 2-arachidonoylglycerol (2-AG). This metabolite is an agonist for cannabinoid receptors (Sugiura et al., 1999). These receptors mediate many neurological effects through the stimulation of PLA2 and PLD isoforms mediated by the activation of PKC (Farooqui et al., 2000b). Similarly, the activation of PLA2 generates arachidonic acid and eicosanoids. These second messengers activate multiple forms of PLD (Klein et al., 1995; Kim et al., 1999). Thus multiplicity and cross-talk among PLA2, PLC, and PLD, along with multiple forms of PKC and other kinases in brain tissue, provides diversity in the function and specificity of various isoforms in regulating enzymic activities in response to a wide range of extracellular signals. However, at the same time it complicates the analysis of PLA2, PLC, and PLD functions in brain tissue (Farooqui and Horrocks, 2005). The complexity of this problem becomes obvious when one considers the coupling of various isoforms of PLA2, PLC, and PLD with different receptors in a single neural cell and tries to associate PLA2, PLC, and PLD activities with neuronal function in normal cells and in disease processes. Isoforms of PLA2, PLC, and PLD do not function interchangeably, but act in parallel to transduce signals (Farooqui and Horrocks, 2004a). It is likely that various isoforms of PLA2, PLC, and PLD act on different molecular species in various cellular pools of glycerophospholipids located in different types of neural cells. These isoforms of PLA2, PLC, and PLD may be regulated by different receptors through different coupling mechanisms (with and without G proteins) involving common second messengers (> Table 2-2). . Table 2-2 Receptors and PLA2, PLC, and PLD Associated with the Degradation of Choline and Ethanolamine Glycerophospholipids in Neural Membranes Receptor Glutamate Biogenic amine

Enzyme PLA2 PLA2, PLC

Coupling mechanism Without G-protein G-protein-linked

Muscarinic acetylcholine

PLA2, PLC, PLD

G-protein-linked

Retinoid Cannabinoid

PLA2, PLC, PLD PLA2, PLC, PLD

Without G protein G-protein-linked

Cytokine Growth factor

PLA2 PLA2

– –

References Kontos et al. (1990); Kolko et al. (1996) Vial and Piomelli (1995); Panchalingam and Undie (2001); Ross (2003) Zian and Drewes (1991); Bayo´n et al. (1997); Hou et al. (2001) Farooqui et al. (2004a) Hashimotodani et al. (2005); Ueda et al. (2005) Atsumi et al. (1998) Jupp et al. (2003)

This process may provide great versatility in ensuring that neural cells use arachidonic and docosahexaenoic acids and their metabolites efficiently. In brain tissue, the activity of PLA2, PLC, and PLD isoforms may depend not only on the structural, physicochemical, and dynamic properties of neural membranes but also on the interaction of extracellular signals with neural cell receptors. The activation of PLA2, PLC, and PLD isoforms in neural cells is the rate-limiting step for the production of lipid mediators such as eicosanoids, docosanoids, and PAF that are involved in inflammatory and anti-inflammatory activities in brain tissues (Bazan, 2005; Phillis et al., 2006). Therefore, a tight regulation of PLA2, PLC, and PLD isozymes is very important for normal neural membrane function. Isoforms of PLA2 play important roles in neuritogenesis, regeneration, apoptosis, inflammation, and neurodegeneration (Farooqui et al., 1997). Isoforms of PLC are involved in the regulation of diverse

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processes such as cell proliferation, contraction, secretion, and phototransduction (Fukami, 2002). Isoforms of PLD are implicated in membrane trafficking, cell proliferation, mitogenesis, oncogenesis, inflammation, neuronal plasticity, secretion, and diabetes (Fukami, 2002; Zhang et al., 2004).

6

Roles of Choline and Ethanolamine Glycerophospholipids in Brain

Besides being integral components of membranes, choline and ethanolamine glycerophospholipids have many other important functions (> Figure 2-3). They are regarded as dynamic molecules whose specific distribution and catabolism are the result of highly regulated processes that can lead to a host of important biological responses during signal transduction. Choline and ethanolamine glycerophospholipids are involved in the following processes. . Figure 2-3 Roles of choline and ethanolamine glycerophospholipids in neural membranes

6.1 Choline and Ethanolamine Glycerophospholipids Provide Precursors for Second Messengers Choline and ethanolamine glycerophospholipids are a reservoir for the generation of many bioactive mediators (Farooqui et al., 1997, 2000b). The activation of a single membrane receptor by an agonist (hormone, growth factor, or neurotransmitter) results in the stimulation of PLA2, PLC, and PLD (Farooqui et al., 2000b; Banno, 2002; Fukami, 2002; Jenkins and Frohman, 2005). This initiates a complex intracellular signaling cascade, as evidenced by the generation of various lipid second messengers (> Table 2-3). This complexity is further enhanced by the fact that the synthesis of one messenger, depending on the time interval following receptor activation, involves different glycerophospholipid substrates and metabolic pathways.

6.2 PLA2, PLC, and PLD-Generated Second Messengers Choline and ethanolamine glycerophospholipids are reservoirs of precursors for the generation of many bioactive mediators. The activation of a single membrane receptor by agonist (hormone, growth factor, or neurotransmitter) results in the stimulation of PLA2, C (PLC), and D (PLD) and initiates a complex intracellular signaling cascade, as evidenced by the generation of various lipid second messengers. Arachidonic and docosahexaenoic acids (AA and DHA), which are liberated by the action of cPLA2 and

Choline and ethanolamine glycerophospholipids

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. Table 2-3 Second Messengers Produced by the Action of Phospholipases on Choline and Ethanolamine Glycerophospholipids Glycerophospholipid Choline and ethanolamine glycerophospholipids

Phospholipase A2

Choline glycerophospholipid Choline and ethanolamine plasmalogens

C

Choline glycerophospholipid

D

A2

Second messenger Arachidonate, eicosanoids, PAF, lysocholine, and ethanolamine glycerophospholipids Diacylglycerol, acetylcholine Arachidonate, eicosanoids, PAF, docosahexaenoate, docosatrienes, resolvins PtdH, lysoPtdH

References Farooqui et al. (2000b)

Fukami (2002) Farooqui et al. (2000b); Bazan (2005); Serhan (2005) Banno (2002); Jenkins and Frohman (2005)

PlsEtn-PLA2 on choline and ethanolamine glycerophospholipids, have been implicated both in physiological and pathological processes. For example, these fatty acids modulate receptors and ion channels, and regulate the activities of many enzymes including protein kinases A and C, NADPH oxidase, DAG-kinase and Naþ, Kþ-ATPase (Farooqui and Horrocks, 2006). AA also inhibits glutamate uptake that is mediated by excitatory amino acid transporters in intact cells, tissue slices, synaptosomes, and various types of neuronal and glial cultures. High concentrations of AA produce a variety of detrimental effects on membrane structure. Thus, it has a profound adverse effect on the capacity of mitochondria to produce ATP. It uncouples oxidative phosphorylation and induces efflux of Ca2þ and Kþ from mitochondria (Farooqui and Horrocks, 2006). AA is metabolized to prostaglandins, leukotrienes, and thromboxanes. These metabolites are collectively called eicosanoids. Their action is mediated through specific cellular receptors called eicosanoid receptors (Phillis et al., 2006). They differ from other intracellular second messengers in one important way—they can cross the cell membrane and leave the cell in which they are generated to act on neighboring cells because of their amphiphilic nature (Farooqui and Horrocks, 2006). DHA affects not only the physicochemical properties (fluidity, permeability, fusion behavior, and elastic compressibility) of neural membranes (Stillwell and Wassall, 2003), but also modulates gene expression of many enzyme proteins involved in signal transduction processes (Horrocks and Farooqui, 2004). DHA modulates dopaminergic, noradrenergic, glutamatergic, and serotonergic neurotransmission, activities of membrane-bound enzymes, ion channels, receptors, learning and memory processes, inflammation and immunity, apoptosis, and gene expression (Horrocks and Farooqui, 2004; Farooqui and Horrocks, 2006). The action of an enzyme resembling 15-lipoxygenase on DHA produces 10, 17S-docosatrienes, 17Sresolvins, and neuroprotectins (Marcheselli et al., 2003; Serhan, 2005). These second messengers are collectively called docosanoids. They act through their specific receptors. These receptors include the resolvin D receptor (ResoDR1), the resolvin E receptor (ResoER1 and RER1), and the neuroprotectin D receptor (NPDR) (Serhan et al., 2004). The docosanoids not only antagonize the effects of eicosanoids, but also modulate leukocyte trafficking and downregulating the expression of cytokines (Marcheselli et al., 2003). Neuroprotectin D1 upregulates the antiapoptotic proteins, Bcl-2 and Bcl-xl and downregulates the expression of the proapoptotic proteins, Bax and Bad expressions (Mukherjee et al., 2004). Collective evidence suggests that the generation of docosanoids is an endogenous protective mechanism against neuroinflammation and neurodegeneration. Lysoglycerophospholipids, the other product of the PLA2-catalyzed reaction, have many effects on various systems. They are precursors for PAF. Choline lysoglycerophospholipid (Lyso-PtdCho) stimulates phenylalanine hydroxylase, alkaline phosphatase, cyclic 3,5-nucleotide phosphodiesterase, protein kinase C, and glycosyl- and sialyl-transferases. It also inhibits activities of acyl-CoA: lysophosphatidylcholine acyltransferase, lysophospholipase, guanylate cyclase, and adenylate cyclase (Farooqui and Horrocks, 2006).

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The action of synaptosomal phosphatidylcholine-specific PLC (PtdCho-PLC) on the sn-3 position of choline glycerophospholipids generates DAG and glycerophosphocholine. DAG stimulates PKC and glycerophosphocholine is either metabolized to free choline by alkaline phosphatase or used by cytidylyltransferase for the synthesis of CDP–choline. PtdCho-PLC is associated with the activation of nuclear transcription factor ĸB (NFĸB) in response to the tumor necrosis factor-a (TNF-a) (Schu¨tze et al., 1992) and the activation of transcription 6 (STAT-6) induced by interleukin-4 (Ramoni et al., 2001). Studies on this topic have suffered from the lack of purified PtdCho-PLC, which has not been cloned from brain tissue. In response to various extracellular stimuli, PLD hydrolyzes choline glycerophospholipids into PtdH and choline. PtdH stimulates the activities of kinases including PKCz, monoacylglycerol acyltransferase, phosphatidylinositol 4-kinase, and PLCg and increases the GTP-bound form of Ras. PtdH can also serve as the precursor for lysophosphatidic acid (lyso-PtdH), which has paracrine or autocrine signal properties (Feng et al., 2003). It increases the activity of tyrosine kinases and Ras-Raf-MAP kinase, PLCg and PLD. Further, both lyso-PtdH and PtdH inhibit the activity of adenylate cyclase through a pertussis-toxin sensitive mechanism, thereby lowering cAMP levels. LysoPtdH also stimulates a heterotrimeric G-protein receptor linked to Gi that initiates tyrosine kinase activation and stimulates the Ras-Raf-MAP kinase. PtdH can also be converted to DAG, which can stimulate PKC. PLD is also involved in membrane trafficking and various transport processes in which acidic glycerophospholipids may facilitate membrane budding and/or fusion (Klein et al., 1995).

6.3 Plasmalogens as Antioxidants and Membrane Fusion Molecules Because of the reactivity of the vinyl ether linkage of plasmalogens with singlet oxygen and other reactive oxygen species (ROS), plasmalogens may act as antioxidants. Thus, plasmalogens not only protect cellular membranes of Chinese hamster ovary cells against certain oxidative stresses, but also play an important role in defending low-density lipoprotein particles (LDL) against oxidative damage (Engelmann et al., 1994; Nagan and Zoeller, 2001). Thus, enrichment of LDL with plasmalogen phospholipids increases their oxidative resistance in vitro. Because of the greater propensity of ethanolamine plasmalogens to undergo lamellar to hexagonal phase transition, vesicles containing ethanolamine plasmalogens undergo fusion six times more rapidly than vesicles containing PtdEtn. This suggests that ethanolamine plasmalogen may be involved in membrane fusion, a process that occurs during synaptic transmission, hormone release, and membrane trafficking (Farooqui and Horrocks, 2001).

6.4 Choline and Ethanolamine Glycerophospholipids in Apoptotic Cell Death Apoptosis is a form of programmed cell death, which is characterized morphologically by nuclear condensation, cell shrinkage, and bleb formation (Sastry and Rao, 2000). Neurochemically, apoptotic cell death is characterized by the stimulation of caspases, a group of endoproteases with specificity for aspartate residues in proteins. During the execution of apoptosis, the cell changes the phospholipid asymmetry of the plasma membrane by rapidly translocating ethanolamine and serine glycerophospholipids to the outer leaflet where the serine glycerophospholipids function as a tag on the dying cell for recognition and removal by phagocytes (Farooqui et al., 2004b). The mechanism of this process is not known. However, a specific inside-outside ethanolamine and serine glycerophospholipid translocase may be involved in the loss of membrane asymmetry during apoptosis (Emoto et al., 1997; Williamson and Schlegel, 2002). This mechanism can explain the extremely rapid kinetics of ethanolamine and serine glycerophospholipid externalization on apoptotic cells. The disruption of glycerophospholipid asymmetry during the execution phase of apoptotic cell death leads to looser glycerophospholipid packing in the outer leaflet, thus allowing Ca2þ entry. The mild alteration in Ca2þ homeostasis and its short duration may lead to neuronal degeneration by the activation of caspases and PLA2 resulting in apoptotic cell death (Farooqui et al., 2004b). The detection of ethanolamine and serine glycerophospholipids on the cell surface can be made with a fluorescent conjugate of annexin V, a Ca2þ- and glycerophospholipid-binding protein that inhibits PLA2. Recent studies also indicate that Ro09-0198, a peptide that specifically recognizes ethanolamine

Choline and ethanolamine glycerophospholipids

2

glycerophospholipids, is a useful probe for studying transbilayer movement in cell membranes. Ro09-0198 can recognize ethanolamine glycerophospholipid exposure in CTLL-2 cells undergoing apoptosis. The exposure of ethanolamine glycerophospholipids correlates well with serine glycerophospholipid exposure on the outer leaflet. A complete loss of the asymmetric distribution of plasma membrane glycerophospholipids may occur during apoptosis. Ethanolamine glycerophospholipid-binding proteins promote cellular resistance to TNF-a-induced apoptosis by inhibiting the activation of the Raf-1/MEK/ ERK pathway, JNK, and ethanolamine glycerophospholipid externalization (Wang et al., 2004). Inhibition of choline glycerophospholipid synthesis also leads to apoptotic cell death (Cui and Houweling, 2002). For example, choline deficiency-induced apoptosis in PC12 cells is associated with a decreased content of choline glycerophospholipids in membranes. Furthermore, the inhibition of CDP–choline: 1,2-diacylglycerol choline phosphotransferase and overexpression of the ethanolamine glycerophospholipid methylation pathway also results in apoptotic cell death. Thus, collective evidence suggests that alterations in the glycerophospholipid metabolism of neural membranes are closely associated with apoptotic cell death (Zweigner et al., 2004).

6.5 PAF and its Role in Brain PAF is a short-lived biologically active ether lipid with diverse physiological and pathophysiological activities (Honda et al., 2002; Ishii et al., 2002). It is involved in inflammation, allergic reactions, and immune responses. It is a potent inducer of gene expression in CNS. It acts as a retrograde messenger for long-term potentiation, a modulator of glutamate release, and an upregulator of memory formation (Bazan, 2003). PAF is released by a wide variety of cells including macrophages, platelets, endothelial cells, mast cells, neutrophils, and neural cells. PAF exerts its biological effects by activating PAF receptors that consequently activate leukocytes, stimulate platelet aggregation, and induce the release of cytokines and expression of cell-adhesion molecules (Maclennan et al., 1996; Honda et al., 2002; Ishii et al., 2002). PAF receptors are linked to G-proteins and activate a variety of intracellular messenger systems such as calcium mobilization, arachidonic acid release, glycerophospholipid turnover, generation of cAMP, and tyrosine phosphorylation (Honda et al., 2002; Ishii et al., 2002). PAF receptors have been cloned from a number of sources including pig lungs and human leukocytes (Honda et al., 2002). PAF activates a wide variety of cells including neurons, neutrophils, eosinophils, monocytes, platelets, and endothelial cells. The physiological activity of PAF is not limited to its proinflammatory effects. PAF is also involved in a variety of other settings including neuronal migration (Tokuoka et al., 2003), gene expression, allergic reactions, and circulatory system disturbances (Fuentes et al., 2002).

6.6 Modulation of Enzymic Activities by Choline and Ethanolamine Glycerophospholipids Many enzymes require glycerophospholipids for their activities. Any hydrophobic molecule can sometimes meet this requirement, but some enzymes are highly specific for a particular glycerophospholipid. For example, neutral protease is regulated by choline and serine glycerophospholipids, phosphatidic acid, and lysophosphatidic acid (Chauhan et al., 2005). Certain serine proteases are inhibited by ethanolamine glycerophospholipid-binding proteins (Hengst et al., 2001). PAF induces the activation of a matrix metalloproteinase associated with endothelial cell invasion and migration (Axelrad et al., 2004). b-Hydroxybutyrate dehydrogenase, an enzyme found in the inner membrane of mitochondria, has an absolute requirement for choline glycerophospholipids. Ethanolamine and serine glycerophospholipids cannot substitute for cholineglycerophospholipids in activating this enzyme. Other enzymes that require specific glycerophospholipids for their activity include Naþ, Kþ-ATPase, Ca2þ, Mg2þ-ATPase, Ca2þ-ATPase, and adenylate cyclase (Farooqui et al., 2000b). These enzymes are involved in maintaining normal ion homeostasis in neurons and glial cells. Alterations in glycerophospholipid composition during disease processes result in changes in membrane fluidity and ion permeability. This process produces an uncontrolled Ca2þ-influx that can induce oxidative stress and inflammatory reactions in brain tissue (Farooqui et al., 2000b).

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Conclusion

Choline and ethanolamine glycerophospholipids are amphipathic molecules found in neural membranes. They are asymmetrically distributed in the bilayer. They, along with serine- and inositol-containing glycerophospholipids, not only form the backbone of neural membranes, but also provide the neural membranes with a suitable environment, fluidity, and ion permeability. The degree of saturation and the length of glycerophospholipid-acyl chains are important determinants of neural membrane characteristics. Choline and ethanolamine glycerophospholipids are synthesized at the endoplasmic reticulum and are transported to other membranous structures by phospholipid exchange and transfer proteins. Once glycerophospholipids are laid down in a membrane, they undergo interconversion reactions (base exchange, methylation, and decarboxylation). These reactions and activities of phospholipases A2, C, and D may be responsible for the turnover, compositional maintenance, and rearrangements of glycerophospholipids in membranes. This process results in the modulation of membrane function. Collective evidence suggests that glycerophospholipids are multifunctional molecules. They are a storage depot for precursors of second messengers, and may be involved in membrane fusion, apoptosis, and regulation of the activities of membrane-bound enzymes and ion channels. It is important to realize that the earlier discussion on metabolism, incorporation, and roles of glycerophospholipids does not circumscribe the entire dynamics of glycerophospholipid metabolism and its regulation by specific genes in brain tissue, but rather provides initial insight into the molecular complexity that is present in neural membranes. It is hoped that this discussion will initiate further studies not only on the regulation of biosynthesis and catabolism of various classes of glycerophospholipids by specific genes, but also on the generation of individual molecular species, their transport, sorting, trafficking, and the role of their second messengers in the central nervous system.

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Brain Phosphatidylserine: Metabolism and Functions

R. Mozzi . S. Buratta

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Phosphatidylserine in Cell Signaling: General Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40

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Phosphatidylserine in the Nervous Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41

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Metabolism of Phosphatidylserine in the Nervous Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42 Studies with Metabolic Labeling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42 In vitro Assays of PtdSer Synthesizing Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44 Regulation of PtdSer Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47 PtdSer Decarboxylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48 Degradation of PtdSer by Phospholipases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49

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Phosphatidylserine in Brain Damage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_3, # Springer ScienceþBusiness Media, LLC 2009

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Brain phosphatidylserine: metabolism and functions

Abstract: Phosphatidylserine (PtdSer) is involved in cell signaling and apoptosis. In the brain, PtdSer is enriched in polyunsaturated fatty acids, particularly docosahexaenoic acid (DHA). Numerous studies have indicated that the abundance of DHA in the brain is essential for optimal neuronal function. PtdSer concentration in the nervous tissue membranes varies with age, brain areas, cell type and subcellular components. PtdSer is synthesized by base exchange between free serine and the nitrogen base present in phosphatidylethanolamine or phosphatidylcholine. The capability to synthesize PtdSer by base exchange varies with cell types, subcellular fractions and developmental stage. At least two isoforms of PtdSer synthesizing enzymes are present in brain. PtdSer cellular levels also depend on its decarboxylation to phosphatidylethanolamine or conversion to lysoPtdSer by phospholipases. The mechanisms regulating PtdSer synthesis and degradation are still not defined. Thus, the role of PtdSer in cell signaling and apoptosis cannot be clearly established at molecular level. Several reports indicate that alteration in PtdSer synthesis might participate to development of brain damage. List of Abbreviations: BEE, base exchange enzyme; CGC, cerebellar granule cells; CHO, Chinese hamster ovary; DHA, docosahexaenoic acid; lysoPtdSer, lysophosphatidylserine; PLA1, phospholipase A1; PLA2, phospholipase A2; PtdCho, phosphatidylcholine; PtdEtn, phosphatidylethanolamine; PtdIns(4,5)P2, phosphatidylinositol-4,5-bisphosphate; PtdIns(3,4,5)P3, phosphatidylinositol-3,4,5-trisphosphate; PtdSer, phosphatidylserine; PSD, PtdSer decarboxylase; PSS, PtdSer synthase; [S]SBEE, serine base exchange enzyme specific for serine; [SE]SBEE, serine/ethanolamine base exchange enzyme

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Phosphatidylserine in Cell Signaling: General Aspects

Phosphatidylserine (PtdSer) is a membrane phospholipid that is receiving great attention for the role played in important cellular processes, such as signaling and apoptosis. Its role in signal transduction became evident after the demonstration that, owing to its structural properties, PtdSer acts as a binding site for the translocation of PKC to plasma membrane (Nishizuka, 1995). Interestingly, all PKC isoforms, independently of their calcium requirements, are strictly dependent on PtdSer for their activities. In a membrane model, the concentration of PtdSer influences the specificity of brain PKC (Newton and Koshland, 1990). Thus, it is possible that changes in PtdSer concentrations in restricted areas of natural membranes, at the PKC interaction site, may have a similar effect. This possibility has not been verified so far. PtdSer also regulates the activity of other enzymes involved in cell signaling, i.e., diglyceride kinase (Sakane et al., 1991), c-Raf-1 protein kinase (Ghosh et al., 1996) and, nitric oxide synthase (Calderon et al., 1994). Recently, the differential role of PtdSer, phosphatidylinositol-4,5-bisphosphate (PtdIns(4,5)P2) and phosphatidylinositol-3,4,5-trisphosphate (PtdIns (3,4,5)P3) in membrane recruitment of PKCa-C2 has been investigated performing biophysical, computational and cell studies. Collectively, these studies show that PtdIns(4,5)P2 and PtdIns (3,4,5)P3 augment the Ca2+- and PtdSer-dependent membrane binding of PKCa-C2 by elongating the membrane residence of the domain but cannot drive the plasma membrane recruitment of PKCa-C2 (Manna et al., 2008). PtdSer modulates also the binding properties of some receptors for their agonists (Foster et al., 1982; Levi et al., 1989; Gagne et al., 1996). More recently, it has been shown that PtdSer promotes neuronal survival (Kim et al., 2000; Salem et al., 2001; Akbar and Kim, 2002), facilitates Akt signaling (Akbar et al., 2005) and serves as signaling switch for the GTPase substrate preference of a GTPase-activating protein (Ligeti et al., 2004). The detection of N-acyl-PtdSer in mammalian brain and other cell types has indicated another possible function of this phospholipid (Guan et al., 2007) because some of the N-acyl PtdSer molecular species might be hydrolyzed to produce the corresponding N-acyl serines. An hypothetical pathway for the production of these compounds has been proposed together with the possibility of generating N-arachidonoyl-L-serine, a novel lipid mediator isolated from bovine brain in trace amounts. In plasma membrane of normal cells, PtdSer is localized in the inner leaflet (Devaux and Zachowski, 1994). This asymmetric distribution is maintained by ATP-dependent mechanisms (Daleke, 2003) but PtdSer is exposed to cell surface in senescent (Bratosin et al., 1998; Pereira et al., 1999) and apoptotic cells

Brain phosphatidylserine: metabolism and functions

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(Zwaal et al., 2005; Tyurin et al., 2008). Recently, Calderon and Kim (Calderon and Kim, 2008) studied intracellular localization of PtdSer in living cells. Annexin V, fused to green fluorescent protein, was expressed in Neuro2A and hippocampal neurons and subsequently monitored for calcium-dependent translocation by fluorescent microscopy. Upon stimulation with ionomycin, cytoplasmic GFP-annexin V translocated to the plasma membrane while nuclear GFP-annexin V moved to the nuclear membrane. Minimal fluorescence was detected in mitochondria, endoplasmic reticulum, Golgi complex and lysosomes, strongly suggesting that PtdSer distribution in the cytoplasmic face of these organelles is negligible. A juxtanuclear structure that was strongly labeled by GFP-annexin V has been identified as the recycling endosome. Exposure of PtdSer on cell surface represents an early event of apoptosis (Martin et al., 1995) and a recognition signal for phagocytes (Wu et al., 2006). This feature has led to the development of methodologies for the identification and quantitative determination of cells undergoing toward apoptotic cell death (Brumatti et al., 2008; Tait, 2008). In addition, more attention has been devoted to the mechanisms involved in the maintenance or loss of membrane asymmetry involving scramblases and flippases (Sahu et al., 2007; Poulsen et al., 2008; Smrz et al., 2008). However, other mechanisms might be also involved because it has been suggested a correlation between PtdSer exposure and PtdSer synthesis (Pelassy et al., 2000). This is in agreement with the stimulation of PtdSer synthesis, observed in thymocytes induced to apoptosis by dexametasone (Buratta et al., 2000), both by metabolic labeling and by measuring PtdSersynthesizing enzyme. Apoptosis represents an important mechanism in developing brain (Oppenheim, 1991) and neuronal apoptosis is associated with various neurodegenerative disorders and cerebrovascular stroke (Li et al., 1995; Nicotera et al., 1999). Microglia plays a critical role in the recognition and removal of apoptotic neurons, representing the tissue macrophages of the CNS. The mechanism of recognition appears complex. Using cocultures of primary microglial cells and cerebellar granule cells (CGCs) and inducing CGC neurons to apoptosis by exposure to the nitric oxide donor, it has been demonstrated that engulfment of apoptotic neurons by microglia is dependent on a carbohydrate–lectin interaction, a vibronectin-mediated mechanism and an interaction between PtdSer and an unidentified receptor for this phospholipid in microglia (Witting et al., 2000). A critical role in apoptosis is played by intracellular Ca2+ mainly stored in mitochondria (Szalai et al., 1999) and endoplasmic reticulum (Scorrano et al., 2003; Bassik et al., 2004). Recently, evidences have been reported that lysosomes respond to the apoptotic stimuli by releasing their luminal Ca2+ and this release is critical for apoptosis-dependent PtdSer exposure to the outer cell membrane surface (Mirnikjoo et al., 2009). All these observations, regarding the role of PtdSer in cellular signaling, suggest that its metabolism has to be tightly controlled at the level of the enzymes for its synthesis and degradation but also at the level of the reactions that convert PtdSer into other phosphoglycerides. This could be important also for another role attributed to PtdSer establishing the proper environment for a number of membrane-bound enzymes such as Na+/K+ ATPase (Palatini et al., 1977) and Ca2+-ATPase in dog brain synaptosomal membranes (Tsakiris and Deliconstantinos, 1985). Despite the great effort devoted to elucidate PtdSer metabolism in mammalian cells, many aspects are still unclear. The detailed description of PtdSer metabolism is particularly difficult for the nervous tissue because of the marked functional differences of brain regions, their cellular heterogeneity and developmental modifications.

2

Phosphatidylserine in the Nervous Tissue

Early observation pointed out that PtdSer content in the nervous tissue varies depending on animal species, age, brain areas, cell types or subcellular compartment (Norton et al., 1975; Porcellati and Goracci, 1976; Witter and Debuch, 1982; Sun and Foudin, 1985). More recent data report the abundance of this phospholipid, with respect to total phospholipids, in myelin (17.4%) and in synaptosomes (12.6%) (Zabelinskii et al., 1999), as well as in specialized membrane microdomains represented by photoreceptor rod outer segment membranes (Martin et al., 2005). On the other hand, PtdSer content can be extremely

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low, as in the primary cultures of astrocytes (2.4%) (Eichberg et al., 1976) or in neuroblastoma cells (2.1%) (Murphy and Horrocks, 1993). Brain is enriched in PtdSer compared to other tissues such as liver, particularly in molecular species that contain the polyunsaturated fatty acid docosahexaenoic acid (DHA, 22:6n-3) (Garcia et al., 1998; Kim et al., 2004). Interestingly, n-3 fatty acids can modulate PtdSer levels in brain cells. In fact, depletion of 22:6n-3 fatty acids causes a decrease of PtdSer synthesis and a reduction of PtdSer level in brain microsomes (Garcia et al., 1998). On the contrary, enrichment of membrane phospholipids with 22:6n-3 significantly promotes the incorporation of this fatty acid into PtdSer and the synthesis of PtdSer leading to an increase of its levels (Kim and Hamilton, 2000). Furthermore, an increase of rat brain 22:6n-3, as well as of the PtdSer content has been observed after intra-amniotic injection of ethyl docosahexaenoate (Green and Yavin, 1995). The correlation between 22:6n-3 content and PtdSer levels has been supported by Hamilton et al. (2000), who measured 22:6n-3 and PtdSer molecular species in rat brain cortex, brain mitochondria, and olfactory bulb, in comparison to liver and adrenal. In fact, brain cortex, brain mitochondria and olfactory bulb, where 22:6n-3 is highly concentrated, contain significantly higher levels of PtdSer in comparison to liver and adrenal where 22:6n-3 is a rather minor component. This study also demonstrates that, in brain cortex, brain mitochondria and olfactory bulb, 45–60% of PtdSer is represented by the molecular specie 18:0, 22:6n-3-PtdSer, whereas in liver and adrenal the most abundant specie is represented by 18:0, 20:4n6-PtdSer. In agreement with previous report (Garcia et al., 1998), dietary depletion of n-3 fatty acids during prenatal and postnatal period, decreased brain 22:6 n-3 and, in this condition, total PtdSer was reduced in rat brain cortex, brain mitochondria and olfactory bulb, but not in liver and adrenal (Hamilton et al., 2000).

3

Metabolism of Phosphatidylserine in the Nervous Tissue

In higher eukaryotes, PtdSer is produced by a energy-independent and calcium-dependent reaction, called base exchange reaction, in which the polar head group of an existing phospholipid (i.e., the choline moiety of phosphatidylcholine (PtdCho) or the ethanolamine moiety of phosphatidylethanolamine (PtdEtn)) is replaced by L-serine (Borkenhagen and Kennedy, 1958). Thus, the mechanism is different from the wellknown mechanism operating in prokaryotes and yeast in which CDP-diacylglycerol reacts with serine. The metabolic fate of PtdSer, synthesized by base exchange, is represented in > Figure 3-1. Apart from the possibility to be subjected to degradation by phospholipases, similarly to other glycerophospholipids, the metabolic correlation with PtdEtn and PtdCho represents a peculiarity that has to be taken into account when approaching the study of the role of PtdSer in cellular processes. In fact, it is well known that PtdSer can be decarboxylated to PtdEtn by the mitochondrial decarboxylase (Dennis and Kennedy, 1972) and that PtdEtn can be methylated also in brain to produce PtdCho (Blusztajn et al., 1979; Mozzi and Porcellati, 1979). The existence of these metabolic interconversion makes particularly difficult the study of the role of PtdSer in cell function especially in brain. In fact, the quantitative relevance of the various steps of the conversion of PtdSer to PtdCho as well as their capability to be modified in particular cell conditions could depend on the cell type. Indeed, studies using metabolic labeling should be accompanied by in vitro studies of the various enzyme activities.

3.1

Studies with Metabolic Labeling

Demonstration of base exchange reaction in vivo was obtained incubating brain cells from 16-day-old rats with radioactive serine (Yavin and Zeigler, 1977). On the basis of the effect of olygomicin and of the ionophore A23182, these authors suggested the existence of two separate mechanisms for base exchange reaction in these cells, one of which might be energy-dependent. They also reported the presence of radioactivity into PtdEtn, produced by PtdSer decarboxylation. More recently, Xu and colleagues (Xu et al., 1993) suggested that PtdSer synthesis in glioma cells may involve more than the headgroup exchange between serine and PtdEtn or PtdCho. The possibility that some of the alternative pathways for PtdSer

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3

. Figure 3-1 Schematic view of phosphatidylserine metabolism

synthesis (Pullarkat et al., 1981; Infante, 1984; Baranska, 1988) could become active in particular cell conditions cannot be excluded. As shown in > Figure 3-1, PtdSer can be decarboxylated to PtdEtn in mitochondria, which can be then methylated to PtdCho. The analysis of radioactivity into PtdEtn and PtdCho, produced from radioactive PtdSer, has been performed in various studies. Reports on the capability of neural cells to convert PtdSer into PtdCho, via PtdSer decarboxylation, are contrasting because of the relative lag period for detecting radioactivity into PtdCho and the necessity to verify that the labeling is in the base and not in the fatty acid moiety. The latter possibility has to be taken into account when considering that in cultured brain cells both PtdCho and PtdIns contained 2% of serine radioactivity incorporated into lipids (Yavin and Zeigler, 1977). Radioactive serine, injected intravenously into mice, labeled PtdCho significantly (Woronczak et al., 1995). Appreciable radioactivity into PtdCho has been also reported by Rhodes and colleagues (Rhodes et al., 1993), after intracerebral injection of radioactive serine in 1-day-old rat pups. Much higher synthesis of PtdCho from 3-3H serine injected intravenously into lateral ventricles of rat brain has been observed, with respect to in vitro studies (Gatti et al., 1989). Almost no radioactivity into PtdCho was observed incubating cerebrocortical slices from 30–60-day-old rats with radioactive serine (Mozzi et al., 1993). The conversion of PtdSer into PtdCho has been also reported in NG 108-15 cells (Rodriguez et al., 1996). The possibility exists that, as suggested by Woronczak et al. (1995), successive metabolic conversion of PtdEtn, produced from PtdSer, to PtdCho requires undisturbed structures of the brain. The use of metabolic labeling provided some interesting information such as the dependence of the capability to synthesize PtdSer in different brain areas and at different ages of the animals (Mozzi et al., 1993; Rhodes et al., 1993), which have been confirmed by in vitro studies (see below). Using C6-glioma cells incubated with radioactive serine, it has been pointed out another important aspect that is the rapid synthesis of PtdSer at the level of plasma membranes (Xu et al., 1994), which is in agreement with the presence of the enzyme in plasma membranes (see below). The study of base exchange reaction in vivo has been approached administering serine or ethanolamine by microdyalisis to rabbit hippocampus and continuously sampling extracellular fluid (Buratta et al., 1998). Local administration of serine or ethanolamine induced a significant increase of ethanolamine or serine, respectively, in the extracellular compartment of the rabbit hippocampus and the characteristics of this effect were conceivable with a base exchange between free serine or ethanolamine and the base present into PtdEtn or PtdSer. This study also

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demonstrated that D-serine can be incorporated in vivo by base exchange, releasing ethanolamine similarly to L-serine. This is in agreement with results in vitro (Kanfer, 1972) and the resulting modulation of the levels of extracellular D-serine may have significant physiological effects, in view of the role of D-serine as a modulator of NMDA receptors (Hashimoto et al., 1995). In this context, it is worthy to note that phosphatidyl-D-serine, which could be produced by this base exchange reaction, has no effects on PKC activity (Newton, 1995). Studies with metabolic labeling also gave useful information on the regulation of PtdSer synthesis that will be discussed below together with the results of in vitro assay of the enzymes.

3.2

In vitro Assays of PtdSer Synthesizing Enzymes

Early reports on the properties of the enzyme that synthesizes PtdSer in brain derive from biochemical studies, which were mainly devoted to establish functions and roles of base exchange reactions, able to convert a phospholipid into another according to the scheme reported in > Figure 3-2. Apart from the importance for PtdSer synthesis, it was hypothesized that base exchange reactions could produce restricted pools of PtdEtn and PtdCho, which could play particular roles with respect to the PtdEtn and PtdCho synthesized by the de novo pathway. The enzymes have been generally indicated as base exchange enzymes (BEE) and, in the case of the synthesis of PtdSer, serine base exchange enzyme (SBEE). The same enzymes were referred as PtdSer synthases (PSS) in studies of PtdSer synthesis in Chinese hamster ovary (CHO) cells and liver (Vance and Steenbergen, 2005). The properties of brain SBEE and of PSS in non-neural tissues have been studied with different approaches. Two main aspects have been investigated in both cases: the specificity for the phospholipid substrate and for the free exchanging base. On this basis, the existence of various enzyme isoforms has been demonstrated. Here, we will mainly report studies related to brain enzymes. From the early 70s, several studies, mainly carried at Porcellati’s and Kanfer’s laboratories, have tried to establish whether the observed incorporation of ethanolamine, serine or choline into the corresponding phospholipids by base exchange was because of the same or different enzymes. This aspect was first approached using a rat brain 35,000 g particulate, prepared by centrifugation of 10,000 g supernatant (Kanfer, 1972). In these experiments, the pH and Ca2+ dependences for the incorporation of choline, ethanolamine and serine in the corresponding phospholipids were determined. In addition, the effect of the pretreatments with phospholipases A2, D and C and, finally, the capability of the various unlabeled bases to displace the radioactive base present into PtdCho, PtdSer and PtdEtn were also studied. The results pointed out some similarities but also some differences between the exchanges with the three bases. One of the first information derived from the studies of base exchange reactions in brain membranes was the existence of one enzyme capable to use both ethanolamine and serine, producing PtdEtn and PtdSer, respectively (Porcellati et al., 1971). Further studies supported this hypothesis but the variability of results from different laboratories, regarding optimal assay conditions and kinetic properties, represented a difficulty in assessing number and

. Figure 3-2 Synthesis of glycerophospholipids by base exchange

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specificity of base exchange enzymes (Kanfer, 1972; Gaiti et al., 1974). This aspect was pointed out by Buchanan and Kanfer (1980) who demonstrated how the procedure for membrane preparation and assay conditions could affect the data on enzyme activity. Developmental studies of the base exchange for ethanolamine, serine and choline were carried out using 35,000 g particulate fraction or homogenates from rats at various ages (4 days to 3 months). In both cases, the developmental patterns for the three substrates were nearly identical with a major peak at approximately 25 days, followed by a gradual decline until another peak of activity was found around 2 months (Saito et al., 1975). Developmental changes in BEE, using serine, ethanolamine or choline as exchanging bases and whole brain homogenate and subcellular fractions of rats from 17 days embryos up to 4 months have been also reported (Kobayashi et al., 1988). Thus, the developmental stage of animals is another aspect that has to be taken into account when comparing data from various laboratories. With respect to the phospholipid substrate, the identification of PtdEtn as the substrate for serine base exchange in brain membranes was first reported by Porcellati and colleagues (Porcellati et al., 1971). The main difficulty in the study of base exchange enzymes is represented by the coexistence, in the same membrane, of both the enzyme and the phospholipid substrate. Thus, the rate of synthesis of PtdSer in various experimental conditions might also depend on the variations at the level of phospholipid annulus. The properties of base exchange for serine, ethanolamine and choline in a 35,000 g brain particulate from 27-days-aged rats were compared with the properties of a solubilized enzyme from the same source (Saito et al., 1975). The particulate enzyme had a pH optimum of 9 for all the three substrates; it was inhibited by p-chlorophenylsulfonate and the inhibition exerted by this sulphydryl-binding reagent was prevented by the presence of dithiothreitol during the incubation. The heat stability was also investigated with both the enzyme sources and a greater loss of choline incorporation was observed in comparison to that of serine or ethanolamine at all temperatures above 30 C. The incubation at 42.5 C for 30 min almost completely destroyed choline exchange activity but only 50% for the other two substrates. The solubilized enzyme had an optimum pH of 7.25 for the incorporation of all the three radioactive substrates and the Km (mol/liter) for ethanolamine, serine and choline were 1.33  10 5, 4.33  10 5 and 6.75  10 4, respectively. The same laboratory succeeded to remove from the solubilized material a protein having almost all the choline base exchange activity and to isolate a serine exchange enzyme that did not have detectable ethanolamine exchange activity (Miura et al., 1981). The optimum pH of this enzyme specific for serine ([S]SBEE) was near 8.0 and the calculated Km value for L-serine was 0.4 mM. Ethanolamine phospholipids were the most effective acceptors for L-serine incorporation. The Km values for the phospholipid substrates were 0.25 mM for ethanolamine plasmalogens (PlsEtn), 0.25 mM for pig liver PtdEtn and 0.66 mM for egg yolk PtdEtn. Neither ethanolamine nor choline inhibited the L-serine exchange activity. Using another protocol, the activity of an ethanolamine and serine base exchange enzyme ([SE]SBEE) of rat brain microsomes was copurified to near homogeneity (Suzuki and Kanfer, 1985). The parallel increase of the ethanolamine and serine exchange enzyme during the purification steps and the presence of a single protein band on SDS-PAGE (100 kDa apparent molecular mass) suggested that a single enzyme catalyzes the incorporation of both substrates. The competitive inhibition exerted by serine on ethanolamine incorporation and vice versa suggested that the enzyme catalyzes the incorporation of both ethanolamine and serine into their corresponding phospholipids. The Km for ethanolamine (0.02 mM) was similar to the Ki of the inhibition of serine incorporation by ethanolamine (0.025 mM). The Km for serine incorporation (0.11 mM) was also quite similar to the Ki for the inhibition of ethanolamine incorporation by serine (0.12 mM). These results suggest that the enzyme should have an identical binding site for ethanolamine and serine with a greater affinity for ethanolamine because the Km for serine is four times higher. Phosphatidylethanolamine and asolectins were the most effective phospholipid acceptors for ethanolamine and serine incorporation. The pH optimum was 7.0 with both substrates. In conclusion, at least two different SBEE isoforms are present in brain. The main difference between the two isoforms, which could be important to define their physiological roles, is the different affinities toward L-serine. In fact, [S]SBEE has a greater Km with respect to [SE]SBEE. Thus, the contribution to PtdSer synthesis of the two isoforms could depend on the intracellular L-serine concentration.

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As mentioned above, the great impulse in the knowledge of properties of the enzymes that synthesize PtdSer by base exchange derives from genetic studies using CHO cells as experimental model. These cells were utilized by Kuge and coworkers, who developed several protocols for the isolation of mutants defective in phospholipid biosynthesis, for studying the synthesis of PtdSer in vivo. In CHO cells, serine exchange is catalyzed by two enzymes at least, named PtdSer synthase I (PSS I) and PtdSer synthase II (PSS II) (Kuge and Nishijima, 1997; Kuge and Nishijima, 2003). PSS I utilizes serine, ethanolamine and choline as free exchanging bases and PtdCho as phospholipid substrate, whereas PSS II utilizes serine and ethanolamine as free exchanging bases and PtdEtn as phospholipid substrate. Recently, Kuge and coworkers (Kuge et al., 2003) purified Chinese hamster PSS II tagged with Flag and HA peptides (FH-PSS II) by successive affinity chromatography. The purified FH-PSS II catalyses serine and ethanolamine, but not choline base exchange. The apparent Km for L-serine was 89 mM and the apparent Vmax was 0.29 nmol/h/mg protein. In addition, the purified enzyme was shown to use PtdEtn but not PtdCho as the phospholipid substrate. Thus, the purified rat brain [SE]SBEE and the purified FH-PSSII have the same substrate specificity, with similar Km values for L-serine. Despite these similarities, molecular weights reported for the two enzymes are different. In fact, the apparent molecular mass (100 kDa) of the rat brain purified [SE] SBEE, determined by SDS-PAGE, is about twofold larger than the calculated molecular mass of Chinese hamster PSS II. Thus, it is uncertain whether or not the purified enzyme corresponds to rat [SE]SBEE. More recently, Tomohiro and colleagues (Tomohiro et al., 2009) purified epitope-tagged forms of human PSSI and PSSII from HeLa cells (FH-hPSSI and FH-hPSSII). The purified PSSII catalyzes the conversion of PtdEtn, but not PtdCho, to PtdSer, this being consistent with the substrate specificity in intact cells. The purified PSSI catalyzed the conversion of both PtdCho and PtdEtn into PtdSer, although in intact cells the enzyme does not contribute to the conversion of PtdEtn to PtdSer to a significant extent. The purified FH-hPSSI catalyzes serine, choline and ethanolamine base exchange for the production of corresponding phospholipids, whereas the purified FH-h PSSII utilizes serine and ethanolamine, but not choline. Authors report Km for L-serine in the presence of exogenous PtdCho or PtdEtn. The apparent Km for L-serine of the purified FH-hPSSI was 67 mM in the presence of 2 mM PtdCho and 24 mM in the presence of 1 mM PtdEtn. The Km for L-serine of the purified FH-hPSSII was 120 mM in the presence of 1 mM PtdEtn. This indicates that the different Km values for L-serine of the enzymes purified in different laboratories could be, in part, due to the use of different phospholipids in the assay mixture. The optimum pH for the purified FH-h-PSSI was between pH 7 and 7.5, both using PtdCho and PtdEtn as phospholipid substrates, whereas the optimum pH of the purified FH-hPSSII was in the vicinity of pH 7.5. Finally, the molecular weights determined by SDS-PAGE were around 40 kDa for FH-hPSSI and around 50 KDa for FH-hPSSII. Mouse PSS I and II have been also cloned. Both predicted proteins, which exhibited only 30% identity, contain 473 residues with a calculated molecular mass 50 KDa (Stone and Vance, 1999). Comparison of PSS I sequences of mouse liver, CHO-K1 and human myeloblast cells revealed a very high degree of conservation across species. At the level of their cDNAs, the three mammalian clones are 80% identical, whereas their amino acid sequences are > 90% identical (Stone et al., 1998). Recently, expression of mRNAs for PSSI and PSSII have been studied in brain. mRNAs encoding for the two PSSs are expressed in murine brain (Bergo et al., 2002). In particular, PSS I is ubiquitously expressed in the various murine tissues, whereas PSS II mRNA is highly expressed in Sertoli cells, Purkinje neurons and in pyramidal neurons of the hippocampus (Bergo et al., 2002). mRNAs for PSSI and PSSII have been also detected in rat cerebral cortex and this analysis was accompanied by the assay of SBEE activity in total membrane fraction (Buratta et al., 2005). The expression of mRNA for PSSI and PSSII was similar in 30-day- and 60-day-old animals, whereas SBEE activity decreased with age. The latter result was in agreement with previous findings on the dependence of PtdSer synthesis by the age of the animal in this brain area (Mozzi et al., 1993). The lowering of brain PtdSer synthesis by base exchange, which is not accompanied by modification of PSSs mRNA expression, could be due or to a regulatory mechanism that is established during the development or to the presence of an unknown isoform of SBEE in young rats. DHA enrichment in Neuro 2A cells increases PtdSer levels but not PSSI and PSSII mRNA expression (Guo et al., 2007).

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3

Another relevant aspect of base exchange enzyme is represented by the localization of the enzyme(s) that synthesize PtdSer by base exchange. The enzyme is generally referred as a microsomal enzyme and, in liver, PSSI and PSSII appears to be associated with mitochondria-associated membranes (Stone and Vance, 2000). Base exchange enzyme activities were reported in plasma membranes from neuronal cells prepared from rabbit brain cortex (Goracci et al., 1973) and in somal plasma membrane enriched fractions from rat brain cortex (Sun and Sun, 1983; Rhodes et al., 1993; Mozzi et al., 1997). It is conceivable that the enzyme located in endoplasmic reticulum should have functions different from that of the enzyme located in plasma membrane, which ought to be more involved in signal transduction. In this context, it is worthy to mention that SBEE activity has been found in Triton-insoluble membranes prepared from cerebellum in which the enzyme could modulate PtdSer level in the binding area for PKC, transforming PtdEtn into PtdSer and vice versa (Buratta et al., 2007). The presence of the enzyme has been confirmed in the Triton-insoluble fractions prepared from cerebral cortex and from cerebrocortical plasma membranes (Buratta et al., unpublished), supporting the role of the enzyme in signal transduction as a modulator of PKC activity. The presence of this enzyme in plasma membranes, in addition to that located in the luminal face of the endoplasmic reticulum (Czarny et al., 1992b), is also relevant because the mobilization of Ca2+ from the endoplasmic reticulum to the cytosol might reduce the activity of the ER-enzyme, and increase that of the plasma membrane enzyme. Base exchange enzymes are also normal constituents of synaptosomal plasma membranes as resulted from the incorporation of radioactive serine, ethanolamine and choline in the corresponding phospholipids in the presence of 1.25 mM Ca2+ (Holbrook and Wurtman, 1988). In this study, base exchange activities were also measured in synaptosomes and the apparent Km were 240 mM for choline, 65 mM for ethanolamine and 67 mM for serine. Base exchange activity with radioactive serine, ethanolamine and choline has been found in neuronal and glial cells (Raghavan et al., 1972) prepared from rat brain of 13–20-day-old animals, according to the procedure of Norton and Poduslo (1970). The incorporation occurred at pH 7.5 with all the three substrates although a second peak of activity was observed at more alkaline pH. The reaction did not require energy and there were no significant differences between the two cell types except for choline incorporation that appeared more effectively in glial cells than in neurons. Using neuronal-enriched and glial-enriched fractions from adult rabbit brain, prepared by a different procedure (Blomstrand and Hamberger, 1969), Goracci and colleagues (Goracci et al., 1973) suggested that base exchange could be used as a neuronal marker on the basis of data indicating that neurons possessed higher serine and ethanolamine base exchange activity than glia.

3.3

Regulation of PtdSer Synthesis

Several studies investigated on the mechanism(s) regulating mammalian PtdSer synthesis but no conclusive information has been obtained so far. Singh and colleagues (Singh et al., 1992a) observed that TPA stimulates the incorporation of serine into PtdSer in LA-N-2 cells. Thus, the effect appeared opposite to that reported for leukemic HL60 cells (Kiss et al., 1987). The discrepancy was not due to differences between neural and non-neural cells, since TPA decreased PtdSer formation in neuroblastoma NB2a cells (Czarny et al., 1992a) and C6-glioma cells (Czarny et al., 1995). The decreased labeling into PtdSer in TPA-treated C6 glioma cells was not due to a greater conversion to PtdEtn, neither to a greater PtdSer degradation. Authors suggested that PtdSer synthesis in these cells might be regulated by the PKC activity but they speculated that the decrease of PtdSer synthesis might play a protective role against persistent and supra-physiological PKC activation caused by TPA. Another attempt to verify whether or not PtdSer synthesis by base exchange was regulated by PKC was carried out by studying the effect of sphingosine and oleylamine on the incorporation of radioactive serine into PtdSer in LA-N-2 cultures (Singh et al., 1992b). Sphingosine and oleylamine stimulated the incorporation of radioactive serine into PtdSer but inhibited that of ethanolamine and choline into the corresponding phospholipids. The stimulatory effect occurred by a PKC-independent process. Authors suggested that some of the non-PKC-mediated effects of sphingosine might be due to the stimulation of PtdSer synthesis.

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On the other hand, they also considered the possibility that some of the PKC-mediated effects of sphingosine were partially ascribable to the enhanced production of PtdSer, the preferred PKC activator. Agonists of group I mGluRs, which include mGluR1s inhibit the incorporation of radioactive serine into PtdSer and the inhibition of PtdSer synthesis, mediated by mGluR1, may participate in the generation of mGluR-EPSP evoked by parallel fiber stimulation (Buratta et al., 2004). By measuring the incorporation of radioactive serine into phospholipids of glioma C6 cells, Czarny and colleagues (Czarny et al., 1992b) reported the inhibitory effects on PtdSer synthesis of glutamate, acetylcholine, thapsigargine and A23187, suggesting that the inhibition was caused by Ca2+-depletion in the endoplasmic reticulum. In LA-N-1 cells, 3H serine incorporation into PtdSer is stimulated by oxotremorine-M (Oxo-M), a muscarinic agonist (Mikhaevitch et al., 1994). Acetylcholine slightly decreases the serine incorporation into lipids and other muscarinic agonists, including pilocarpine and carbachol, have no effect. Stimulation of PtdSer synthesis by Oxo-M is prevented by G-protein activators and by G-protein inhibitors. Also, the protein kinase C inhibitor (H7) and overnight exposure to PMA are able to prevent the Oxo-M stimulation of serine incorporation. Interestingly, treatment with Oxo-M, which stimulates PtdSer synthesis, causes a release of Ca2+ from intracellular stores, as previously reported for Jurkat cells (Kaneda et al., 1993), and this appears in contrast with the results reported for C6 glioma cells (Czarny et al., 1992b). The stimulatory effect of chlorpromazine on PtdSer synthesis has been demonstrated measuring the incorporation of radioactive serine into brain slice PtdSer and SBEE activity in brain microsomes (Rhodes et al., 1993). SBEE activity of a membrane fraction from rat brain is stimulated by amphiphilic cations and inhibited by amphiphilic anions (Kanfer and McCartney, 1993). Studies on the regulation of base exchange activities in brain membranes demonstrated that base exchange enzymes are regulated by phosphorylation-dephosphorylation (Kanfer et al., 1988). On the other hand, the two isoforms of SBEE purified from brain membrane have different activators. In fact, the isoform specific for serine is stimulated by sphingosine, whereas the isoform capable to utilize both serine and ethanolamine is stimulated by arachidonate at concentrations that correspond to those reached in brain ischemia (Kanfer and McCartney, 1991). The regulation of PtdSer synthesis has been studied measuring the incorporation of radioactive serine into PtdSer in the homogenate of rat brain cortex and assaying SBEE activity in plasma membranes prepared from the same brain area. The results suggest that PtdSer synthesis is subjected to various regulatory mechanisms, involving ATP and G proteins, possibly acting on different enzyme isoforms (Mozzi et al., 1997). Another possibility is that PtdSer synthesis in brain membranes could be regulated by a feed-back mechanism, similar to that identified in CHO cells, in which the activity of PSSs are regulated by the end product PtdSer (Kuge et al., 1998, 1999). In fact, in brain microsomes (Rossi et al., 1990) and in plasma membranes from cerebral cortex (Mozzi, unpublished), SBEE activity is inhibited by the addition of exogenous PtdSer. The relationship between DHA and PtdSer contents has been mentioned above and it is interesting that this polyunsaturated fatty acid increases PtdSer content only in neuronal cells (Guo et al., 2007). To explain the difference between neuronal and non-neuronal cells, authors suggest the existence of a third isoform of PSS, different from PSSI and PSSII that may be responsible for PtdSer accumulation specifically in neuronal cells. The possibility that this unknown isoform of the enzyme could correspond to the isoform specific for serine, purified by Kanfer and coworkers (Miura et al., 1981), is suggestive. All together, the above reported results confirm that the major difficulty in assessing a cellular role to the enzyme(s) that synthesize PtdSer by base exchange is the lack of stated information on number and properties of the enzyme isoforms present in brain, even because the information obtained in other cell types cannot be directly extrapolated to brain and, in particular, to the various specialized cells present in this tissue.

3.4

PtdSer Decarboxylation

PtdSer decarboxylation occurs in mitochondria (Dennis and Kennedy, 1972) and is mediated by the enzyme PtdSer decarboxylase (PSD) which is present in the mitochondrial inner membrane (van Golde et al., 1974) with its active site facing the intermembrane space (Zborowski et al., 1983). The relative

Brain phosphatidylserine: metabolism and functions

3

contribution of the biosynthetic pathways to cellular PtdEtn content has not been firmly established, but it appears to depend on cell type. In rat liver and hamster heart, the CDP-ethanolamine pathway has been reported to produce the majority of PtdEtn (Zelinski and Choy, 1982; Tijburg et al., 1989). In contrast, in many types of cultured cells (Voelker, 1984; Kuge et al., 1986), the decarboxylation of PtdSer produces more than 80% of PtdEtn even when the culture medium is supplemented with ethanolamine, an obligatory substrate of CDP-ethanolamine pathway. In brain, PtdSer decarboxylation might be particularly important because the blood–brain barrier restricts the entry of most small polar compounds. A first indication that a considerable portion of PtdEtn is formed via a direct decarboxylation of PtdSer has been observed in differentiating cells from rat brain cerebral hemispheres in culture (Yavin and Zeigler, 1977). These authors also provided evidences of the possible decarboxylation of 1-alkyl-2-acyl-sn-glycero-3-phosphoserine to 1-alkyl-2-acyl-sn-glycero-3phosphoethanolamine in the same cells. In vitro studies demonstrated that brain mitochondria possess PSD activity (Butler and Morell, 1983), which is localized on the mitochondrial inner membrane (Percy et al., 1983). The localization of PSD in the inner mitochondrial membrane arise the question about the translocation of PtdSer from its synthesis site to decarboxylation site. In brain, PtdSer translocation is enhanced by Ca2+, but is not influenced by cytosolic factors. Furthermore, labeled PtdSer is transferred better to the mitochondrial membrane from microsomes than from artificial membranes such as liposomes containing microsomal phospholipids (Corazzi et al., 1993). Recently, Kuge and colleagues (Kuge et al., 2001) found that the transport-dependent decarboxylation of PtdSer in permeabilized CHO.K1 cells is remarkably enhanced by cytosolic factors from bovine brain. In the brain, the PtdEtn produced by PtdSer decarboxylation seems to be mainly utilized in the assembly of the inner mitochondrial membrane (Carlini et al., 1993; Camici and Corazzi, 1995). Recently, Wen and Kim (2007), using deuterium-labeled PtdSer, demonstrated that 18:0, 22:6 PtdSer is the best substrate for brain mitochondria PSD. Since, 22:6 n3 containing phospholipids are the preferred substrate for PtdSer synthesis (Kim et al., 2004), Authors suggest that the enzymes involved in maintaining the Ptdser status in brain favor the 22:6 containing species.

3.5

Degradation of PtdSer by Phospholipases

Early studies, using specifically labeled PtdSer in vitro, demonstrated that this phospholipid can be hydrolyzed by phospholipases A1 (PLA1) and A2 (PLA2) present in the nervous tissue (Woelk and Porcellati, 1973; Woelk et al., 1973, 1974). More recent reports established the presence of various isoforms of PLA2 in the nervous tissue, as well as in other tissues. These enzymes constitute a superfamily on the basis of their structural and catalytical properties which comprises 15 groups, grouped and numbered on the basis of the catalytic mechanism as well as functional and structural features (Burke and Dennis, 2009). All the groups can be divided into five principal kinds of enzymes, the sPLA2s, the cPLA2s, the Ca2+-independent PLA2s (iPLA2s), the PAF acetylhydrolases (PAF-AH), and the lysosomal PLA2s. There is a large body of evidence for the occurrence of enzymes, belonging to various groups in mammalian nervous tissue or neural cells even though none of them has been isolated (Farooqui and Horrocks, 2004). Indeed, the expression of genes coding for sPLA2, cPLA2 and iPLA2 isoforms has been demonstrated in neural cells in culture and in different regions of brain (Molloy et al., 1998; Zanassi et al., 1998). An extensive survey on brain PLA2 and their involvement in neurological disorders has been recently published (Farooqui and Horrocks, 2007). PtdSer seems to be a poor substrate for cPLA2 and iPLA2 which hydrolyze preferentially PtdCho or PtdEtn. The secreted PLA2, particularly sPLA2-IIA, is able to hydrolyze PtdSer as well as other phosphoglycerides producing 1-acyl-2-lyso- sn-glycero-3-phosphorylserine (2-lysoPtdSer). This enzyme is expressed in astrocytes and can be induced by TNF-a and IL-1b (Li et al., 1999; Tong et al., 1999; Lin et al., 2004). Thus, owing to the asymmetric distribution of membrane phospholipids, this enzyme may hydrolyze PtdSer when the phospholipid is exposed to the outer surface of damaged cells because the enzyme is strongly inhibited by proteoglycans (Murakami et al., 2000). sPLA2-type IIA is also present in intracellular compartments of neural cells. Particularly, the mitochondrial enzyme is released in the

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Brain phosphatidylserine: metabolism and functions

cytoplasm under reduced membrane potential in energy-deficient cells (Macchioni et al., 2004). Thus, in such conditions, PtdSer and other phospholipids of the inner leaflet might also be a potential substrate, causing membrane alterations and, very likely, cell damage. However, intracellular sPLA2-IIA could also have a protective effect removing oxidized fatty acids because of its low specificity for fatty acids at sn-2 position of glycerophospholipids. This possibility might be particularly relevant for preventing PtdSer exposition under mild oxidative stress because oxidized PtdSer translocates more easily to cell surface (Tyurina et al., 2000; Kagan et al., 2004). Meanwhile, great attention has been devoted to this class of hydrolytic enzymes; however, towing to the correlation with the production of lipid mediators (i.e., eicosanoids, platelet-activating factor, 2-lysoglycerophospholipids), studies on brain PLA1 are limited. In mammals, nine PLA1 molecules have been identified and six of them are extracellular enzymes (Aoki et al., 2007). One of these enzymes shows a marked specificity for PtdSer (PtdSer-PLA1) (Sato et al., 1997; Aoki et al., 2002) and produces 1-lyso-2acyl-sn-glycero-3-phosphorylserine (1-lysoPtdSer). cDNA encoding the human enzyme has been cloned and sequenced (Aoki et al., 2002). PtdSer-PLA1 is highly expressed in platelet, heart, and lung and to a minor extent in rat brain. In human tissues, it is expressed mainly in liver, testis, lung and kidney, whereas there is little or no expression in brain (Aoki et al., 2002). This enzyme is able to hydrolyze both PtdSer and 2-lysoPtdSer, the product of PLA2 hydrolysis of PtdSer (Sato et al., 1997). An alternative splicing form for PtdSer-PLA1 possesses only lysophospholipase activity specific for 2-lysoPtdSer (Nagai et al., 1999). PtdSer-PLA1 is a secreted enzyme and its action should be limited on the surface of cells where its substrate is almost absent in normal cells. Thus, it has been proposed that this enzyme may bind to surface proteoglycans and hydrolyze the exposed PtdSer in apoptotic cells (Hosono et al., 2001). The produced 1-lyso PtdSer could act as a lipid mediator similar to 2-lyso PtdSer produced by the action of PLA2. Evidence that lysoPtdSer may act as a lipid mediator has been reported (Martin and Lagunoff, 1979; Bruni et al., 1984), including the effect on the promotion of neurite outgrowth in PC12 cells (Lourenssen and Blennerhassett, 1998). In neural cells, it has been shown that lysoPtdSer induces rat astroglia stellation without altering the basal level of cAMP (Facci et al., 1987). A direct effect of lysoPtdSer on the cellular machinery underlying stellation has been proposed. An intracellular effect of lysoPtdSer has also been proposed because it partially inhibits purified PLD from rat brain (Ryu and Palta, 2000). Sometimes, studies on the effects of lysoPtdSer on biological system do not specify whether 1-lysoPdtSer, the product of PLA1, or 2-lysoPtdSer, the product of PLA2, were used. The structural and biochemical properties of the two products are rather different because 1-lysoPtdSer contains mainly polyunsaturated fatty acids whereas 2-lysoPtdSer is largely saturated. At the best of our knowledge, it is not known whether or not neural cells store and secrete PtdSer-PLA1. However, the presence of PLA1 and PLA2 able to hydrolyze membrane PtdSer of neural cells (Woelk et al., 1973) allows the remodeling of molecular species of this phospholipid and, together with lysophospholipases, the complete breakdown of PtdSer.

4

Phosphatidylserine in Brain Damage

Several reports demonstrated that ethanol exposure modifies PtdSer metabolism and this aspect may be relevant since ethanol influences signal transduction and alters various functions of cell brain membranes (Chandler et al., 1998; Gerstin et al., 1998). The acidic phospholipids have been suggested as possible targets for ethanol action (Sun and Sun, 1985). In adult rats, chronic ethanol administration causes an increase in the acidic phospholipids (PtdSer, PtdIns and phosphatidic acid) in rat brain membranes (Sun et al., 1984) and an increase in PtdSer in guinea pig synaptic plasma membranes (Sun and Sun, 1983). Those modification have been attributed to the development of an adaptative mechanism. In a neural-derived hybrid NG108-15 cell line, ethanol stimulated both serine incorporation into PtdSer and, after 2 days of treatment, PtdSer content (Rodriguez et al., 1996). However, ethanol exposure in utero, reduces brain PtdSer synthesis of newborn rats both in vitro and in vivo, as demonstrated by the assay of the serine base exchange activity in brain microsomes and by measuring PtdSer radioactivity after incubation of slices with radioactive serine or intracerebral injection of the precursor (Hu et al., 1992).

Brain phosphatidylserine: metabolism and functions

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The inhibitory effect of ethanol on PtdSer synthesis in utero has been confirmed incubating cortical slices from 5-day-old rat pups with radioactive serine or injecting cerebrally the radioactive precursor into newborn pups from the ethanol-treated dams, with respect to control. Interestingly, the effect could be reversed by chloropromazine administration (Rhodes et al., 1993). Reduction of PtdSer synthesis by ethanol exposure has also been reported in C6-glioma cells by Wojcik and colleagues (Wojcik et al., 2000), who utilized in their experiments the same protocol for alcohol exposure that caused increase of PtdSer synthesis in NG108-15 cells (Rodriguez et al., 1996). In evaluating the difference between the two cell types, it has to be considered that C6 cells are transformed non-excitable glial cells (Baranska et al., 1999), which do not contain voltage-gated and ligand–gated Ca2+ channels, whereas NG108-15 are excitable cells (Putney, 1993). Recently, Wen and Kim (2007) measured the effect of maternal exposure to ethanol, using an in vitro assay for PtdSer synthesis by incubating microsomes with deuterium-labeled phospholipids. Ethanol inhibited significantly microsomal PtdSer synthesis by base exchange with PtdCho liposomes. This inhibition particularly affected the utilization of 18:0, 22:6 molecular species which is the preferred substrate. A similar effect was also observed when assaying the enzyme with PtdEtn liposomes. The reduction of in vitro PtdSer synthesis using brain microsomes was consistent with the reduction of PtdSer in the cortex, similar to that already observed in hippocampus as a consequence of ethanol treatment (Wen and Kim, 2004). However, Authors did not observe variation in mRNA levels for PSSI and PSSII, nor in the expression of PSSI, for which a suitable antibody was available, and this led to suggest that PSSI and PSSII are modified by metabolites of ethanol or ethanol could have altered the microenvironment of PSS enzymes. Modification of PtdSer levels and/or synthesis has been reported in brain pathologies. For example, PtdSer level is increased in synaptosomal plasma membranes from cerebral cortex of Alzheimer disease (Farooqui et al., 1997) and the activity of PtdSer synthase is elevated in substantia nigra of patients with Parkinson’s disease (Ross et al., 2001). In rat cerebrocortical slices, N2 treatment stimulates the incorporation of radioactive serine into PtdSer and the effect is greater in adult than in young animals, which is known to be more resistant to hypoxia (Mozzi et al., 1993). On the other hand, administration of L-serine by microdialysis to the hippocampus of healthy rabbits causes a rapid and transient increase of extracellular levels of ethanolamine, produced by the base exchange reaction with membrane PtdEtn; this was followed in time by an increase in extracellular levels of phosphoethanolamine, which appeared because of a PKC-dependent phospholipase C activation (Buratta et al., 1998). Since extracellular levels of ethanolamine and phosphoethanolamine increase in brain ischemia (Hagberg et al., 1985), it is possible to hypothesize that stimulation of PtdSer synthesis represents one of the early event involved in brain hypoxia/ischemia. However, the presence of various mechanisms for regulating PtdSer synthesis may cause a different response to the same treatment in different brain areas and/or cell types. In fact, in cerebellar slices, N2 treatment inhibits the incorporation of radioactive serine into PtdSer; this is likely due to the activation of mGluR1 receptors by the released glutamate (Buratta et al., 2004). In fact, these receptors are highly expressed in cerebellum and poorly expressed in cerebral cortex (Catania et al., 1994). It is well known that brain aging is accompanied by changes in the overall composition of membrane lipids (Rouser et al., 1971; Sun and Samorajski, 1972; Horrocks et al., 1981). Several reports demonstrate that those changes include modification in PtdSer synthesis (Gatti et al., 1989; Ilincheta de Boschero et al., 2000; Giusto et al., 2002). Several compounds, including PtdSer, have been proposed as memory enhancer with some benefits for age-related memory decline; the experimental evaluation has been revised by McDaniel and colleagues (McDaniel et al., 2003).

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in cell signaling and apoptosis synthesis by base exchange and its modulation decarboxylation to phosphatidylethanolamine degradation by phospholipases in brain damage

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Metabolism and Enzymology of Cholesterol and Steroids

B. Stoffel-Wagner

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60

2 2.1 2.2 2.3 2.4 2.5 2.6

Synthesis and Metabolism of Steroids in the Human Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 Cytochrome P450 Cholesterol Side-Chain Cleavage (P450SCC) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 5a-Reductase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 3a-Hydroxysteroid Dehydrogenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62 Cytochrome P450 Aromatase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 17b-Hydroxysteroid Dehydrogenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 P450c17 (17a-Hydroxylase/C17-20-lyase), Steroid Sulfatase, Hydroxysteroid Sulfotransferase, Organic Anion Transporter Polypeptides, and 7a-Hydroxylase (CYP7B1) . . . . . . . . . . . . . . . . . . . . . . . . . 64 2.7 Other Steroidogenic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65 3

Clinical Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_4, # Springer ScienceþBusiness Media, LLC 2009

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Metabolism and enzymology of cholesterol and steroids

Abstract: This chapter summarizes the current knowledge on the metabolism of steroids in the human brain, the enzymes mediating these reactions, their localization, and the putative effects of steroids in the brain. The presence of the steroidogenic enzymes cytochrome P450SCC, aromatase, 5a-reductase, 3ahydroxysteroid dehydrogenase, 17b-hydroxysteroid dehydrogenase, and steroid sulfatase in human brain has now been firmly established by molecular biological and biochemical studies. Their presence in the cerebral cortex and in the subcortical white matter indicates that various cell types, either neurons or glial cells, are involved in the biosynthesis of neuroactive steroids in the brain. The following functions are attributed to specific neuroactive steroids: modulation of GABAA, N-methyl-D-aspartate (NMDA), nicotinic, muscarinic, serotonin (5-HT3), kainate, glycine and sigma receptors, neuroprotection, and induction of neurite outgrowth, dendritic spines, and synaptogenesis. The first clinical investigation in humans produced evidence for an involvement of neuroactive steroids in conditions such as depressive disorders, catamenial epilepsy, fatigue during pregnancy, premenstrual syndrome, and postpartum depression. Further and improved knowledge of the biochemical pathways of steroidogenesis and the actions of neuroactive steroids on the brain may enable new perspectives in the understanding of the physiology of the human brain as well as in the pharmacological treatment of its disturbances. List of Abbreviations: 3b-HSD, 3b-hydroxysteroid dehydrogenase; DHEA, dehydroepiandrosterone; DHEAS, dehydroepiandrosterone sulfate; GABAA, g-aminobutyric acid A; NMDA, N-methyl-D-aspartate; OATP-A, organic anion transporter polypeptide-A; OATP, organic anion transporter polypeptides; 17b-HSD, 17b-hydroxysteroid dehydrogenase; STS, steroid sulfatase; SULT2, hydroxysteroid sulfotransferase; 3a-HSD, 3a-hydroxysteroid dehydrogenase

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Introduction

Steroid hormones are mainly synthesized in the gonads, the adrenal glands, and the feto-placental unit. The brain is an important target organ of steroid hormones. Steroids can easily cross the blood–brain barrier because of their high lipid solubility. In the brain, an extensive steroid metabolism occurs. In addition, several brain regions are well equipped with enzymes necessary for steroid hormone biosynthesis (Martini and Melcangi, 1991; Lephart, 1993; Naftolin, 1994; Robel et al., 1999). Development, growth, maturation, and differentiation of the brain are strongly influenced by steroid hormones. As shown in animal studies, steroids synthesized de novo in the central nervous system (i.e., neurosteroids) can affect multiple brain functions (i.e., neuroendocrine and behavioral functions) via intracellular receptors that regulate transcriptionally directed changes in protein synthesis. These actions occur within hours or days. In addition to the classic genomic actions of steroids, neuroactive steroids are able to rapidly alter excitability of the central nervous system through binding to neurotransmitter-gated ion channels, thus modulating g-aminobutyric acid A (GABAA) and N-methyl-D-aspartate (NMDA) receptors (Majewska, 1992; Mellon, 1994). These actions occur within seconds or milliseconds via ligand- or voltage-gated ion channels. In the case of aromatase, the activity of steroidogenic enzymes was identified in human fetal brain tissue more than 30 years ago (Naftolin et al., 1975). However, the majority of biochemical, physiological, and behavioral studies on aromatase in brain tissue were carried out in rodents or other animal species. For a long time, studies in humans have been precluded due to the difficulty in obtaining fresh human brain tissue, coupled with a presumed low expression or activity of the respective enzymes. This also applies to other steroidogenic enzymes. Steroidogenesis requires numerous sequential enzymatic reactions to convert cholesterol to sex hormones, glucocorticoids, or mineralocorticoids. As the steroids produced within a tissue depend upon the enzymes present in this tissue, only systematic studies on the expression of all relevant steroidogenic enzymes would allow insight into the steroidogenic pathways and the capacity within the respective tissue, that is, the human brain. Reports on the expression and activity of the most important steroidogenic enzymes in the human brain have been published in recent years. This chapter reviews the current knowledge on metabolism and enzymology of cholesterol and steroids within the human brain and the evidence we have for its importance.

Metabolism and enzymology of cholesterol and steroids

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Synthesis and Metabolism of Steroids in the Human Brain

2.1 Cytochrome P450 Cholesterol Side-Chain Cleavage (P450SCC) The first and rate-limiting step in the synthesis of steroid hormones is the conversion of cholesterol to pregnenolone, catalyzed by the enzyme cytochrome P450 cholesterol side-chain cleavage (P450scc). Human P450SCC is encoded by a single gene on chromosome 15, the CYP11A1 gene (Chung et al., 1986). Not only is P450scc present in the adrenal glands and gonads, the major sources of steroid hormone production, it is also present in the placenta, primitive gut, and brain (Simpson and MacDonald, 1981; Mellon and Deschepper, 1993; Keeney et al., 1995). Once pregnenolone is produced from cholesterol, it may be converted to progesterone and other neuroactive steroids. However, the major role of P450SCC in the brain is probably the regulation of brain neurosteroid levels (Warner and Gustafsson, 1995). Recently, we investigated the expression of CYP11A1 mRNA in tissue specimens from temporal and frontal neocortex, subcortical white matter from the temporal lobe, and hippocampus from patients with medically intractable chronic temporal lobe epilepsy (Beyenburg et al., 1999; Watzka et al., 1999). In these brain areas, CYP11A1 mRNA was expressed in significant amounts in all tissue samples investigated, however, at a rate 200 times lower than that in adrenal tissue, which is known for highest CYP11A1 expression. Thus, CYP11A1 mRNA expression in the human brain is within the range previously estimated for rat brain in qualitative RT-PCR experiments (Mellon and Deschepper, 1993; Sanne and Krueger, 1995; Warner and Gustafsson, 1995). In the temporal and frontal neocortex as well as in the hippocampus of women, CYP11A1 mRNA concentrations were significantly higher than those found in men (Beyenburg et al., 1999; Watzka et al., 1999). During childhood, CYP11A1 mRNA concentrations in the temporal lobe increase markedly and reach adult levels at puberty (Watzka et al., 1999). These data showed for the first time that an age- and sex-dependent expression of CYP11A1 mRNA occurs in the human brain. Few data are available on the relative amount of CYP11A1 mRNA in the brain of male and female animals, but qualitative studies report no obvious sex differences in rats (Mellon and Deschepper, 1993; Kohchi et al., 1998). Due to the insensitivity of qualitative RT-PCR in detecting differences in mRNA expression at high cycle numbers, a careful quantitative reexamination of results obtained in rat brain with respect to sex differences of CYP11A1 mRNA expression seems to be necessary. In situ hybridization and cell culture experiments in rat brain demonstrated predominant CYP11A1 expression in the subcortical white matter (Hu et al., 1987; Sanne and Krueger, 1995). No such differences could be detected between neocortex and subcortical white matter tissue in the human brain (Watzka et al., 1999). Evidence that pregnenolone can be produced in the central nervous system is provided by the presence of CYP11A1 mRNA in human brain tissue.

2.2 5a-Reductase Numerous animal studies have shown that in the central nervous system, progesterone is rapidly metabolized to 5a-dihydroprogesterone (5a-DHP), which is then further reduced to the potent neurosteroid 3a, 5a-tetrahydroprogesterone (3a, 5aTHP) (Mellon, 1994). These conversions are catalyzed by 5a-reductase and 3a-hydroxysteroid dehydrogenase (3a-HSD). In humans, two isozymes of 5a-reductase, which differ in tissue distribution and biochemical characteristics as well as in their responsiveness to specific inhibitors of their enzymatic activity, have been identified (Andersson and Russell, 1990; Andersson et al., 1991). The majority of physiological and biochemical studies on the expression of 5a-reductase in the brain were carried out in rodents and other vertebrate species (Martini, 1982; Martini and Melcangi, 1991; Lephart, 1993; Li et al., 1997). However, some investigators documented 5-reductase activity in human fetal brain (Mickan, 1972; Schindler, 1976; Saitoh et al., 1982). Only in a few frontal lobe and temporal lobe tissue specimens was 5a-reductase activity demonstrated in the brain of adults (Jenkins and Hall, 1977; Celotti et al., 1986). Recently, we demonstrated the predominant expression of 5a-reductase type 1 mRNA in a large series of human temporal neocortex and subcortical white matter as well as hippocampal tissue specimens

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obtained from patients with chronic temporal lobe epilepsy (Stoffel-Wagner et al., 1998b; Stoffel-Wagner et al., 2000). The expression levels were about 100 times lower than in human liver tissue. 5a-Reductase type 2 mRNA was not expressed. Another study reported on 5a-reductase type 1 mRNA expression in a few human cerebellum, hypothalamus, and pons tissue specimens that were collected postmortem (Thigpen et al., 1993). In addition, a predominant expression of 5a-reductase type 1 mRNA was found in rat brain (Normington and Russell, 1992; Lephart, 1993). In rat brain, 5a-reductase type 1 mRNA is expressed at all stages of brain development and in adulthood, with a small increase around the time of birth. However, 5a-reductase type 2 mRNA is only transiently expressed during the late fetal and early postnatal life (Poletti et al., 1998). The expression patterns of this isoform overlapped the secretory profile of testosterone. It has been hypothesized that increased levels of circulating androgens occurring in male rats around the time of birth could modulate 5a-reductase type 2 expression. Hence, transient androgen-regulated expression of 5a-reductase type 2 may be important for sexual differentiation of the brain and for the formation of anxiolytic/anesthetic steroids originating from 3a-hydroxylation of 5a-reduced derivates of progesterone involved in stress responses associated with parturition. However, we still do not know whether 5a-reductase type 2 mRNA might be expressed transiently during fetal or early postnatal life within the human brain. We also measured 5a-reductase activity in human temporal neocortex and subcortical white matter tissue specimens (Stoffel-Wagner et al., 1998b; Steckelbroeck et al., 2001a). Although enzyme activity was present in all tissue specimens under investigation, the apparent Km values and the pH profile substantiated the predominant expression of the type 1 isoform. Moreover, we investigated the inhibitory effects of MK386, a specific inhibitor of the 5a-reductase type 1 isoform, and of finasteride, a specific inhibitor of the 5a-reductase type 2 isoform on 5a-reductase activity (Steckelbroeck et al., 2001a). MK386 turned out to be a strong inhibitor of human brain tissue 5a-reductase activity, with an IC50 value of 2.0 nmol/l, whereas finasteride was a poor inhibitor of the reaction, with an IC50 value of 142.8 nmol/l (Steckelbroeck et al., 2001a). Furthermore, we observed a potent inhibition of the pH-dependent reaction by MK386 but not by finasteride. An, at least predominant, activity of the 5a-reductase type 1 isozyme in the human brain is substantiated by these findings (Steckelbroeck et al., 2001a). There were no sex-specific differences in the expression levels of 5a-reductase type 1 mRNA in human brain tissue or in the activity of 5a-reductase (Stoffel-Wagner et al., 1998b, 2000; Steckelbroeck et al., 2001a). These findings are consistent with previous animal studies, where no significant sex-specific differences concerning 5a-reductase activity were found in rat brain (Massa et al., 1975; Selmanoff et al., 1977) and in neural tissue of rhesus macaques during fetal development (Resko et al., 1988).

2.3 3a-Hydroxysteroid Dehydrogenase Multiple cDNAs encode proteins related to 3a-HSD in humans (Qin et al., 1993). However, at least four 3a-HSD isozymes exist, which share at least 84% of its amino acid sequence identity (Khanna et al., 1995a, b; Penning, 1997; Penning et al., 2000). These are known as type 1 3a-HSD (AKR1C4), type 2 3a-HSD (AKR1C3), type 3 3a-HSD (AKR1C2), and 20a(3a)-HSD (ACR1C1). This isoform is predominantly a 20a-HSD, and this change in positional specificity implies that it may play an important role in regulating progesterone action (Penning et al., 2000). Penning and coworkers (2000) demonstrated that all human 3a-HSD isoforms and the human 20a-HSD act as 3-, 17-, and 20-ketosteroid reductases as well as 3-, 17-, and 20-hydroxysteroid oxidases. Recently, we could demonstrate the expression of the mRNA of type 2 and 3 isozyme of 3a-HSD as well as 20a-HSD in the hippocampus and the temporal lobe of patients with medically intractable temporal lobe epilepsy, whereas the mRNA of the type 1 isozyme of 3a-HSD was not expressed (Stoffel-Wagner et al., 2000; Steckelbroeck et al., 2001a). The expression levels of 3-HSD 2 were about one fifth of that in liver tissue, those of 3a-HSD 3 about one tenth of that in liver tissue, and those of 20a-HSD were about 2% (1/ 40) of that in liver tissue (own unpublished data). The expression levels of 3a-HSD 2 and 3 as well as 20a-HSD mRNAs in human hippocampus did not differ significantly between the sexes. This is in accordance with data on 3a-HSD activity in the rat brain (Stoffel-Wagner et al., 2000).

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All of these three isoforms, 3a-HSD 2, 3 and 20a-HSD are capable of producing the neuroactive tetrahydrosteroids that modulate the GABAA receptor (Penning et al., 2000). Consequently, the meaning of the differential expression of the single isoforms is less established than ever.

2.4 Cytochrome P450 Aromatase Cytochrome P450 aromatase, which catalyzes the conversion of androgens into estrogens in specific brain areas, is the product of the CYP19 gene, which has been cloned and sequenced (Corbin et al., 1988; Harada, 1988). Only in a few fetal brain specimens has aromatase activity itself been determined (Naftolin et al., 1971a, b; Doody and Carr, 1989). Previously published data demonstrated aromatase activity in human temporal and frontal brain areas (Wozniak et al., 1998). The authors studied biopsy materials removed at autopsy from normal adult control subjects and from patients with Alzheimer’s disease. Temporal aromatase activity was always significantly higher than frontal aromatase activity regardless of sex and/or disease state. This difference was consistent with our own studies on the expression of temporal and frontal CYP19 mRNA in fresh brain tissue specimens from adult patients with chronic epilepsy undergoing neurosurgery (StoffelWagner et al., 1999a). CYP19 mRNA was not only expressed in temporal and frontal neocortex, but also in subcortical white matter of the temporal lobe and in the human hippocampus (Stoffel-Wagner et al., 1998a, 1999a). Sex-specific differences in CYP19 mRNA expression could be observed in none of these brain areas. In our laboratory, we were able to characterize aromatase activity in the temporal lobe in brain tissue specimens of a similar cohort of patients with epilepsy (Steckelbroeck et al., 1999a). We demonstrated a specific, dose-responsive, and competitive inhibition of its activity by atamestane, which is a known specific and competitive inhibitor of placental aromatase activity (Henderson et al., 1986). Compared with its high activity in the placenta, aromatase activity in the human brain was low. However, rates of aromatase activity in the brain were in the same order of magnitude as in human adipose and testicular tissue (Ackerman et al., 1981; Rowlands et al., 1991). Aromatase activity was significantly higher in the cerebral neocortex than in the subcortical white matter (Steckelbroeck et al., 1999a). For CYP19 mRNA expression in the human temporal lobe, this difference could not be found (Stoffel-Wagner et al., 1998a). However, in the human temporal neocortex, CYP19 mRNA concentrations were significantly lower in children than in adults (Stoffel-Wagner et al., 1998a). This finding could not be confirmed by measurement of aromatase activity (Steckelbroeck et al., 1999a). These contradictory findings indicate that aromatase might be regulated on the posttranslational level.

2.5 17b-Hydroxysteroid Dehydrogenase Seven human isozymes of 17b-hydroxysteroid dehydrogenase (17b-HSD) have so far been cloned. They all play a major role in the regulation of the biological activity of sex hormones, and they are essential for the biosynthesis of the strong androgens and estrogens testosterone and estradiol from their weaker precursors androstenedione and estrone (Krazeisen et al., 1999; Peltoketo et al., 1999). These conversions are reversible and thus can lead to a deactivation of the respective sex hormones (Labrie et al., 1997). The different isozymes show an individual cell-specific expression and substrate specificity. The importance of the 17b-HSD activity in the maintenance of physiological levels of estradiol and testosterone is reflected by the ubiquitous distribution of 17b-HSD in peripheral tissues (Martel et al., 1994). 17b-HSD activity in the human brain has been reported about 35 years ago (Jaffe, 1969; Jenkins and Hall, 1977). However, studies on the expression of the enzyme in the human brain remain rare. Western immunoblot analysis revealed the presence of 17b-HSD 1 in human fetal brain (Milewich et al., 1990). Recently, we demonstrated the expression of 17b-HSD 1, 3, 4, and 5 mRNA in the human temporal lobe and hippocampus (Stoffel-Wagner et al., 1999b; Steckelbroeck et al., 2001b), whereas an in tandem pseudogene of 17b-HSD 1 and 17b-HSD 2 mRNA was not expressed (Stoffel-Wagner et al., 1999b; Steckelbroeck et al., 2001b). We also characterized androgenic and estrogenic 17b-HSD activity in the

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human temporal lobe and found the NAD-dependent oxidation of testosterone and estradiol as well as the NADPH-dependent reduction of androstenedione and estrone (Steckelbroeck et al., 1999b). Substrate specificity, pH optima, cofactor requirement patterns, and kinetic properties suggest the activity of at least two isozymes, namely the activating 17b-HSD 3 and the deactivating 17b-HSD 4, in the human brain. No sex differences in the expression or activity of 17b-HSDs were observed. However, the expression levels of 17b-HSD 3, 4, and 5 mRNAs as well as the conversion of androstenedione, testosterone, estrone, and estradiol were significantly higher in the subcortical white matter than in the cerebral neocortex (Steckelbroeck et al., 1999b, 2001b; Stoffel-Wagner et al., 1999b). The predominant expression of 17bHSD in the subcortical white matter suggests that glial cells could play a role in the biosynthesis and deactivation of sex steroids in the brain. Among a host of potential functions of glia, glial cells are involved in the formation of myelin. This suggests a possible correlation among sex steroids, these enzymatic activities, and the formation or functions of myelin. In a recent study on the human 17b-HSD 7 gene (HSD17B7), its promotor revealed binding sites for brain-specific transcription factors corresponding to expression domains in the developing brain as identified by in silico Northern Blot (Krazeisen et al., 1999). To date, 17b-HSD 8 expression has not been investigated in the human brain.

2.6 P450c17 (17a-Hydroxylase/C17-20-lyase), Steroid Sulfatase, Hydroxysteroid Sulfotransferase, Organic Anion Transporter Polypeptides, and 7a-Hydroxylase (CYP7B1) Within the brain, dehydroepiandrosterone (DHEA) and its sulfate (DHEAS) were found to be subject to a series of enzyme-mediated conversions. A large number of studies suggest importance of DHEA(S) or their cerebral metabolites for vitality, development, and functions of the brain (Majewska, 1995; Baulieu and Robel, 1996; Compagnone and Mellon, 1998; Herbert, 1998; Garcia-Estrada et al., 1999; Weill-Engerer et al., 2003). In steroidogenic glands, DHEA is synthesized by conversion of cholesterol to pregnenolone via P450scc followed by conversion of pregnenolone to DHEA via P450c17 (17a-hydroxylase/C17-20-lyase). Previous studies failed to demonstrate 17a-hydroxylase activity or P450c17 mRNA in the adult rat brain (Mellon and Deschepper, 1993; Baulieu and Robel, 1996). However, P450c17 mRNA as well as P450c17 protein were detected in the brain of rat embryos using ribonuclease protection assays and immunocytochemistry (Compagnone et al., 1995). In adults, conflicting data have been reported: Compagnone and coworkers (1995) reported expression of P450c17 mRNA only in the peripheral nervous system of rats and mice, while others demonstrated the presence of P450c17 mRNA in various brain regions of adult rodents (Stromstedt and Waterman, 1995). Using fresh human temporal lobe biopsies, neither activity nor mRNA were detected of the enzymes essential for the formation of DHEA from pregnenolone (i.e., P450 c17) and DHEAS from DHEA (i.e., hydroxysteroid sulfotransferase SULT2) (Steckelbroeck et al., 2004). These data demonstrate that within the human temporal lobe, DHEA(S) are not synthesized de novo. The conversion of DHEAS into DHEA is catalyzed by microsomal steroid sulfatase (STS). The hydrolysis of the sulfate ester bond of other 3b-hydroxysteroid sulfates, such as cholesterol sulfate, pregnenolone sulfate, and estrone sulfate, is also catalyzed by this enzyme (Shapiro, 1985). It is encoded by a single X-chromosomal gene (Yen et al., 1987; Stein et al., 1989). Recently, strong activity and mRNA expression of DHEAS desulphating was found in temporal lobe biopsies, twice as high in cerebral cortex than in subcortical white matter (Steckelbroeck et al., 2004). Immunohistochemistry revealed STS in adult cortical neurons as well as in fetal and adult Cajal–Retzius cells (Steckelbroeck et al., 2004). The question arises whether DHEA(S) is produced de novo within the human central nervous system or whether high levels of circulating DHEAS contribute to cerebral DHEA(S) levels. Cellular uptake of organic anions, such as steroid sulfates, from the blood across the blood–brain barrier and through the plasma membrane requires specific membrane transporters (Hagenbuch et al., 2002). Among these, organic anion transporter polypeptide-A (OATP-A) is expressed throughout the human brain and was present at the blood–brain barrier (Kullak-Ublick et al., 1998; Gao et al., 2000). Previously, cerebral expression of OATP-B, -D, and -E

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has been demonstrated (Tamai et al., 2000). In a recent study, OTAP-A, -B, -D, and -E showed high mRNA expression levels with distinct patterns in human cerebral cortex and subcortical white matter (Steckelbroeck et al., 2004). These membrane transporters might be involved in the transport of steroid sulfates from the blood into the CNS. In the CNS, DHEAS and/or other 3b-hydroxysteroids can be converted via neuronal STS activity. Steroid hormones are known to act via binding to specific transactivating receptor proteins. As no such DHEA(S) receptor has been found, it is assumed that DHEA(S) do not act like other steroid hormones that directly regulate gene expression. It has been suggested that intracrine metabolism of DHEA(S) is an important factor in mediating their effects (Labrie et al., 1998, 2000). Moreover, nongenomic effects of DHEA(S) are obviously also responsible for their actions in the CNS. In the rodent brain, DHEA is primarily catalyzed via 7a-hydroxylase and 17b-HSD activity (Baulieu and Robel, 1996). Both reactions require reduced pyridine nucleotides as coenzymes and lead to the formation of 7a-hydroxy-DHEA (androst-5-ene-3b, 7a-diol-17-one) and D5-androstenediol (androst-5-ene-3b,17b-diol), respectively. Recently, membrane-associated DHEA activity and expression of high levels of CYP7B1 mRNA expression were demonstrated in the human temporal lobe (Steckelbroeck et al., 2002). 7a-Hydroxylase activity was significantly higher in the cerebral cortex than in the subcortical white matter. The high levels of CYP7B1 mRNA in the brain as well as in a variety of other tissues and the ubiquitous presence of 7a-hydroxylase activity in the human brain allowed the assumption of a neuroprotective function of the enzyme, such as counteracting deleterious effects of neurotoxic glucocorticoids or regulation of the immune response rather than a distinct brain-specific function such as neurostimulation or neuromodulation.

2.7 Other Steroidogenic Enzymes Other important steroidogenic enzymes are 3b-hydroxysteroid dehydrogenase (3b-HSD), 21-hydroxylase (cytochrome P450c21), 11b-hydroxylase (cytochrome P45011b), and cytochrome P450 aldosterone synthetase (P-450aldo). 3b-HSD catalyzes the conversion of D5-3b-hydroxysteroids into D4-3-ketostreroids (i.e., the conversion of pregnenolone into progesterone). 21-Hydroxylase converts progesterone to 11-deoxycorticosterone and 17-hydroxyprogesterone to 11-deoxycortisol, the substrates required for the production of the main adrenal steroids, corticosterone, aldosterone, and cortisol. 11b-Hydroxylase (cytochrome P45011b) catalyzes the formation of glucocorticoids (cortisol and corticosterone). Cytochrome P450 aldosterone synthetase (P-450aldo) exerts three enzyme activities (11b-hydroxylation, 18-hydroxylation, and 18-oxidoreduction) and catalyzes the formation of mineralocorticoids (aldosterone). Only a small number of studies on the expression of 21-hydroxylase in the brain exist to date. In rodents, 21-hydroxylase was detected in the brain stem using the reverse transcription polymerase chain reaction assay and immunohistochemical methods (Iwahashi et al., 1993; Stromstedt and Waterman, 1995), whereas other investigators could not find 21-hydroxylase mRNA in any extra-adrenal tissue (Mellon and Miller, 1989). As this may be due to the limited sensitivity of the mRNA quantification assay, we investigated the expression of 21-hydroxylase mRNA in the human hippocampus using a highly sensitive nested RT-PCR assay (Beyenburg et al., 2001). We demonstrated for the first time the expression of 21-hydroxylase mRNA in the human hippocampus. In the hippocampus, the expression levels are approximately 10,000 times lower than that the adrenal gland, which is known for high 21-hydroxylase expression (Beyenburg et al., 2001). However, we could not measure the enzyme activity of 21-hydroxylase since only small amounts of tissue specimens were available. Although our results clearly demonstrate that 21-hydroxylase mRNA is expressed in small amounts in the human hippocampus, it remains debatable whether hippocampal tissue contains sufficient 21-hydroxylase to produce neuroactive steroid concentrations of physiological or pathophysiological relevance. The mRNAs of 3b-HSD 1 and 2 as well as cytochrome P45011b and cytochrome P450 aldosterone synthetase were expressed neither in the human temporal lobe nor in hippocampus (own unpublished data). For these investigations a sensitive, nested competitive RT-PCR assay was used. However, several

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studies demonstrated the expression of 3b-HSD mRNA (Dupont et al., 1994; Guennoun et al., 1995; Sanne and Krueger, 1995) and 3b-HSD protein (Guennoun et al., 1995) in the rat brain. Data concerning the expression of cytochrome P45011b in rodent brain are conflicting: while some authors report the expression throughout the rat brain (Stromstedt and Waterman, 1995; Gomez-Sanchez et al., 1996), others found only low expression levels in rat brain (Mellon and Deschepper, 1993; Erdmann et al., 1996) or no expression in mouse brain (Stromstedt and Waterman, 1995). Cytochrome P450 aldosterone synthetase expression and activity have been demonstrated in various regions of rat brain including hypothalamus, hippocampus, amygdala, and cerebellum (Gomez-Sanchez et al., 1996; Gomez-Sanchez et al., 1997).

3

Clinical Implications

The presence of the already mentioned steroidogenic enzymes cytochrome P450SCC, aromatase, 5a-reductase, 3a-HSD, 17b-HSD, and STS in human brain has now been firmly established by molecular biological and biochemical studies. These findings provide evidence that neuroactive steroids can be produced within the human brain. However, the (patho) physiological significance of these findings remains to be elucidated. > Figure 4-1 presents a summary of current knowledge and open questions on biochemical pathways of steroid metabolism in the human brain.

. Figure 4-1 Current knowledge and open questions concerning the biochemical pathways of metabolism of cholesterol and steroids in the human brain. Solid arrows indicate that the activity of the respective enzyme as well as the expression of its mRNA has been documented with the exception of P450SCC and 21-hydroxylase (marked by an asterisk) as here only the expression of its mRNA has been shown. Dashed arrows indicate that the occurrence of the enzyme has not yet been found in the nervous system. DOC, deoxicorticosterone; DHT, dihydrotestosterone; 5a-DHP, 5a-dihydroprogesterone; 3a,5a-THP, 3a,5a-tetrahydroprogesterone (allopregnanolone); 5aR, 5a-reductase; 3a-HSD, 3a-hydroxisteroid dehydrogenase; 3b-HSD, 3b-hydroxisteroid dehydrogenase; 17b-HSD, 17b-hydroxisteroid dehydrogenase; 21-H, 21-hydroxylase; CYP7B1, oxysterol 7a-hydroxylase

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Steroid hormone effects on the brain have typically been associated with gene regulation via intracellular steroid receptors. These reproductive and neuroendocrine actions of steroids via intracellular receptors, which regulate transcriptionally directed changes in protein synthesis, generally occur within hours or days. In addition to the classic sites of steroid synthesis, neurosteroids can rapidly alter the excitability of the central nervous system by modulating neurotransmitter-gated ion channels such as GABAA and NMDA receptors (Baulieu and Robel, 1990; Majewska, 1992, Mellon, 1994). GABA, a major inhibitory neurotransmitter, mediates fast synaptic inhibition by activating ligandgated chloride channels. Binding of 3a-reduced neurosteroids to GABAA receptors results in either inhibition or potentiation of the inhibitory effects of GABA (> Figure 4-2).

. Figure 4-2 Effects of neurosteroids on GABAA receptor function

Hence, anticonvulsive, anesthetic, and anxiolytic effects of neuroactive steroids are mediated by their capacity to positively modulate GABAA receptor function, that is, these substances act to increase GABAergic effects by increasing of frequency and duration of chloride channel openings (Majewska, 1992; Mellon, 1994). On the other hand, inhibition of GABAA receptor function, which is mostly documented for the neuroactive steroids pregenenolone sulfate and DHEAS, produces effects ranging from anxiety and excitability to seizure susceptibility (Paul and Purdy, 1992; Baulieu, 1997, 1998). In addition, other actions of neuroactive steroids have been described in the brain including the inhibition of NMDA receptor function as well as the modulation of other receptors, such as serotonin (5-HT3), nicotinic, muscarinic, glycine, kainate, and sigma receptors (Wu et al., 1991; Prince and Simmonds, 1992; Lambert et al., 1995; Monnet et al., 1995; Mensah-Nyagan et al., 1999; Rupprecht and Holsboer, 1999). Moreover, it has been postulated that neuroactive steroids act on nerve cells through membrane receptors coupled to G proteins (Orchinik et al., 1992) and may also interact with various neuropeptide receptors (Grazzini et al., 1998). In summary, neuroactive steroids exert both genomic and nongenomic effects, and regulate neuronal function via their concurrent influence on gene expression and transmitter-gated ion channels. These actions suggest that neuroactive steroids play a crucial role in mediating many brain functions. Moreover, the systemic effects of neuroactive steroids may be beneficial for a variety of neuropsychiatric disorders. So far, the majority of physiological and behavioral studies have been carried out in rodents or other vertebrate species. In recent years, evidence for an intensive formation of neuroactive steroids within the human brain has emerged and now the first clinical investigations exist to support the results obtained in preclinical animal studies. As early as 1941, Seyle had suggested the potential anesthetic properties of neuroactive steroids, resulting in the development of steroid anesthetics, for example, alphaxalone (Richards and Hesketh, 1975). However,

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side effects have hindered the development of steroid anesthetics for routine clinical use (Paul and Purdy, 1992). The observation that epileptic seizures in cycling women are less frequent in the luteal phase, when circulating levels of progesterone are high, appears to be associated with cyclical variations in the metabolism of progesterone to allopregnanolone in the brain (Backstrom, 1975, 1976; Mellon, 1994; Reddy, 2004). Progesterone and 3a-reduced neuroactive steroids have potent anticonvulsant effects (Belelli et al., 1990; Kokate et al., 1994). Synthetic derivates of neuroactive steroids are under investigation for treatment of epilepsy disorders. Already, some preliminary investigations in healthy volunteers and in patients with medically intractable epilepsies have been undertaken. Ganaxolone, for example, showed a promising pharmaco-kinetic profile and was well tolerated in a trial with healthy volunteers (Monaghan et al., 1997a, b). It was also effective in clinical studies with patients with epilepsy (Shields et al., 1997; Kerrigan et al., 2000). Although promising, potential side effects call for caution. For example, progesterone and 3a, 5a-THP have benzodiazepine-like effects (Belelli et al., 1990; Kokate et al., 1994), and progesterone withdrawal may lead to an increase in seizure susceptibility. The development of sensitive assays to measure cerebral fluid or blood neurosteroid concentrations enabled researchers to document alterations in neurosteroidogenesis in human diseases. Recently, Strohle and coworkers (1999) demonstrated decreased 3a,5a-tetrahydroprogesterone plasma concentrations in patients with major depression compared with healthy control subjects. In addition, clinically effective antidepressant treatment was accompanied by an increase of 3a,5a-tetrahydroprogesterone in the plasma of these patients. Neuroactive steroids may also be involved in physiological conditions where fluctuations of the hormonal balance occur. For example, increased fatigue during pregnancy may be the result of higher concentrations of progesterone and GABA agonistic 3a-reduced neuroactive steroids such as 3a,5a-THP (Biedermann and Schoch, 1995), whereas a rapid decline in these substances may lead to the premenstrual syndrome or postpartum depression (Wang et al., 1996; Rupprecht, 1997). Moreover, fluctuations in neuroactive steroid concentrations may in part contribute to the increased risk of developing psychiatric diseases in women at the perimenstrual phase, during pregnancy and the postpartum period, and around menopause. In alcoholic patients, reduced plasma concentrations of GABA agonistic 3a-reduced neuroactive steroids have been found during ethanol withdrawal (Romeo et al., 1996). This decline in 3a-reduced neuroactive steroid concentrations may contribute to the increased seizure liability during ethanol withdrawal. DHEA and DHEAS are the most abundant circulating steroid hormones in humans. Their concentrations decrease with age and under stress (Orentreich et al., 1992; Goodyer et al., 1996). It was hypothesized that DHEA and DHEAS may be neuroprotective agents as both age and stress are associated with neuronal vulnerability to degeneration. Indeed, neuroprotection by DHEA and DHEAS was observed in vivo in hippocampal structures (Kimonides et al., 1998). The mechanisms by which DHEA and DHEAS act are still unknown. In patients with Alzheimer’s disease and multi-infarct dementia, decreased DHEAS concentrations have also been reported (Nasman et al., 1991; Hillen et al., 2000; Magri et al., 2000). To date, trials in which DHEA was administered for a short period of 2 weeks have failed to demonstrate any benefit of DHEA therapy in cognitive performance (Wolf et al., 1997, 1998; Huppert et al., 2000). Hence, high-quality trials are required with the duration of DHEA treatment in excess of a few weeks and with a sufficiently large number of participants to detect possible effects with the outcome measures including objective tests of cognitive function.

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S. Petrosino . V. Di Marzo

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76

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Biosynthesis and Inactivation of Acylethanolamides in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76 Biosynthesis of Acylethanolamides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76 Brain Distribution of N-Acyl-Phosphatidyl-Ethanolamine-Selective Phospholipase D . . . . . . . . . . . . 79 Degradation of Acylethanolamides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 Transport of Acylethanolamides across the Plasma Membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80

3 3.1 3.2 3.3 3.4 3.5 3.6 3.7

Established and Potential Molecular Targets of Acylethanolamides in the Nervous System . . . . 80 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80 Cannabinoid Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 82 ‘‘Orphan’’ G-Protein-Coupled Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Ion Channels: TRPV1 and TRPM8 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84 Other Ion Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Peroxisome-Proliferator Activating Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 Unidentified Binding Sites and GPCRs for Other Mediators . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85

4 4.1 4.2 4.3 4.4

Role of Anandamide and Other Acylethanolamides in the Nervous System . . . . . . . . . . . . . . . . . . . . 86 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 Control of Food-Intake, Satiety, and Gastrointestinal Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 Role in Neuroprotection and Neuroinflammation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88 Control of Nociception . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89

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Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 90

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_5, # Springer ScienceþBusiness Media, LLC 2009

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Abstract: Fatty acid ethanolamides, also known as N-acylethanolamines or acylethanolamides (AEs), have been known as naturally occurring lipids in animals and plants since the 1950s. Interest in their biological function and pharmacology in the central nervous system was revived after the identification of one of them, arachidonoylethanolamide (anandamide, AEA), as the first endogenous ligand of cannabinoid CB1 receptors, the most abundant G-protein-coupled receptors in the mammalian brain. Next came the discoveries that some AEs can also activate peroxisome-proliferator-activating receptors as well as transient receptor potential vanilloid type channels. The regulation and major known biological functions of AEA and other AEs in the nervous system are reviewed in this chapter.

1

Introduction

Since the discovery of arachidonoylethanolamide (known as anandamide, AEA, from the Sanskrit word ananda for bliss) as the first endogenous ligand of the receptors for marijuana’s active principle D9-tetrahydrocannabinol (THC) (Devane et al., 1992), the fatty acid ethanolamides (or acylethanolamides, AEs), endogenous lipids known since the 1950s (> Figure 5-1) (Long and Martin, 1956; Bachur et al., 1965), have received ever increasing attention. Emerging evidence indicates that AEA, which apart from cannabinoid receptors interacts also with several ion channels (Oz, 2006), is not the only member of this family of fatty acid amides to be endowed with important pharmacological activities and neural functions. Palmitoylethanolamide (PEA), perhaps the first pharmacologically active AE to have been discovered, is emerging as an important neuroprotective and anti-inflammatory mediator acting at several molecular targets in both central and sensory nervous systems as well as immune cells (Lo Verme et al., 2005; Re et al., 2007). Stearoylethanolamide (SEA), a compound with proapoptotic (Maccarrone et al., 2002a) and anorexic (Terrazzino et al., 2004) properties, has been suggested to recognize specific and as yet uncharacterized binding sites in the brain (Maccarrone et al., 2002b). Finally, oleoylethanolamide (OEA), regulates food-intake and lipogenesis (Rodriguez de Fonseca et al., 2001; Fu et al., 2003; Oveisi et al., 2004), by activating peroxisomeproliferator-activating receptor a (PPAR-a), thus exerting pharmacological actions on energy homeostasis that are opposite to those that AEA exhibits via cannabinoid CB1 receptors (Di Marzo and Matias, 2005; Matias et al., 2007). Other long-chain acylethanolamides, such as myristoyl-, linoleoyl-, linolenoyl, and docosahexaenoyl-ethanolamides (> Figure 5-1), have also been investigated (Aloe et al., 1993; Maurelli et al., 1995; Bisogno et al., 1999; Movahed et al., 2005), although less thoroughly than AEA, PEA, and OEA. Therefore, much less is known about their pharmacology. The same is true for some polyunsaturated members of this family, such as 5Z,8Z,11Z-eicosatrienoyl-ethanolamide (Priller et al., 1995), di-homo-g-linolenoyl-ethanolamide, and 7Z,10Z,13Z,16Z-docosatetraenyl-ethanolamide (> Figure .5-1), also known as ‘‘anandamides’’ because, like AEA, they potently activate cannabinoid receptors (Hanus et al., 1993; Pertwee et al., 1995). In this chapter, we shall review the biochemistry and pharmacology of AEA and other AEs, with particular emphasis on their proposed functions in the nervous system.

2

Biosynthesis and Inactivation of Acylethanolamides in the Brain

2.1 Biosynthesis of Acylethanolamides All AEs appear to share, although possibly only to a certain extent, common biosynthetic and degradative pathways. The enzyme most likely responsible for the biosynthesis of AEs from their direct biosynthetic precursors, the corresponding N-acyl-phosphatidyl-ethanolamines (NAPEs), is known as NAPE-selective phospholipase D (NAPE-PLD) and has been cloned (Okamoto et al., 2004; Wang et al., 2006). N-arachidonoyl-phosphatidyl-ethanolamine (NArPE), N-oleoyl-PE, and N-palmitoyl-PE are respectively converted into AEA, OEA, and PEA by NAPE-PLD. However, other possible pathways exist for the conversion of NAPEs into the corresponding AEs (Di Marzo and Petrosino, 2007; Liu et al., 2008) (> Figure 5-2). This is suggested by the fact that NAPE-PLD-null mice do not contain lower levels of AEA, OEA, or PEA in all tissues compared with wild-type mice (Leung et al., 2006). In particular, only the levels

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. Figure 5-1 Chemical structures of the most studied endocannabinoids and acylethanolamides

of saturated AEs were decreased in the brain of NAPE-PLD( / ), and this reduction was most dramatic for AEs bearing very long acyl chains (>or = C20). It is emerging that: (1) NAPEs can be hydrolyzed also by a secretory phospholipase 2 (sPLA2) into N-acyl-lyso-phosphatidylethanolamines (lyso-NAPE) before being further hydrolyzed to AEs by a lysophospholipase D (Sun et al., 2004); (2) NAPEs are also substrates for a-/ b-hydrolase 4 (Abh4) acting as a lysophospholipase/phospholipase B for the formation of glycerolphospho-AEs in the mouse brain (Simon and Cravatt, 2006); and finally (3) a PLC-dependent pathway for N-arachidonoyl-PE (NArPE) conversion to phospho-AEA, followed by formation of AEA via the protein tyrosine phosphatase N22 (PTPN22), might also exist (Liu et al., 2006, 2008) (> Figure 5-2).

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. Figure 5-2 Biosynthesis and degradation of acylethanolamides (AEs). Abh4, a,b-hydrolase 4; AEA, arachidonoylethanolamide (anandamide); FAAH, fatty acid amide hydrolase; FAAH-2, fatty acid amide hydrolase-2; NAAA, N-acylethanolamine acid amidase; NAPE, N-acylphosphatidyl-ethanolamine; lyso-PLD, lyso-phospholipase D; NAPE-PLD, NAPE-specific phospholipase D; OEA, oleoylethanolamide; PEA, palmitoylethanolamide; PLC, phospholipase C; PTPN22, protein tyrosine phosphatase N22; sPLA2, soluble phospholipase A2. AEA, OEA, and PEA are produced from the direct or indirect processing of the corresponding NAPEs. Also other AEs can be formed and degraded through these pathways

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2.2 Brain Distribution of N-Acyl-Phosphatidyl-Ethanolamine-Selective Phospholipase D Although other enzymes might be responsible for AE biosynthesis in the mammalian brain, so far only the distribution of NAPE-PLD has been investigated in the mouse brain by means of in situ hybridization and immunohistochemical techniques. Egertova´ and colleagues (2008) recently showed that the enzyme is most expressed in the dentate gyrus of the hippocampus, particularly in the axons of granule cells (mossy fibers). Intense NAPE-PLD immunoreactivity was detected in the axons of the vomeronasal nerve that project to the accessory olfactory bulb. NAPE-PLD expression was also detected in other brain regions (e.g., cortex, thalamus, hypothalamus, cerebellum), but with significantly lower intensity. The authors suggested that NAPE-PLD is expressed by specific populations of neurons in the brain and targeted to axonal processes, and therefore that AEs generated by this enzyme in axons may act as anterograde synaptic signaling molecules that regulate the activity of postsynaptic neurons. Similar results were also reported by Nyilas et al. (2008), who also used high-resolution quantitative immunogold labeling to demonstrate that this Ca2+-sensitive enzyme is localized predominantly on the intracellular membrane cisternae of axonal Ca2+ stores. Not completely overlapping results were instead obtained by Cristino et al. (2008), who did confirm the presence of NAPE-PLD in the dentate gyrus, although at both the levels of somata and fibers, and, by using two different polyclonal antibodies, identified the enzyme also in the CA3 or CA1 regions of the hippocampus, in somata of pyramidal cells or presynaptic to pyramidal cells, respectively. A possible explanation for these discrepant results is that the antibodies and staining conditions used by these latter authors also labeled NAPE-PLD at its site of production in the endoplasmic reticulum of the somas. In fact, by using ins situ hybridization, the NAPE-PLD mRNA was identified in the somas of pyramidal cells in the CA3 region (Egertova´ et al., 2008; Nyilas et al., 2008). Cristino et al. (2008) also found NAPE-PLD immunoreactivity in the somata of some Purkinje’s cells of the cerebellar cortex and also presynaptically to these cells, in the molecular layer, possibly at the level of GABAergic basket cells. The authors suggested that AEs generated by this enzyme, apart from acting as anterograde signals (e.g., in the hilus area of the dentate gyrus and in the CA1 region of the hippocampus), might also work as intracellular messengers in postsynaptic neurons (such as CA3 pyramidal neurons and Purkinje’s cells) by activating transient receptor potential vanilloid type-1 (TRPV1) channels. TRPV1 is indeed one of the molecular targets for AEA, OEA, and linoleoylethanolamide, which activate this channel by acting at an intracellular binding site (Movahed et al., 2005; Starowicz et al., 2007a). Furthermore, TRPV1 is coexpressed with NAPE-PLD as well as with the AE-hydrolyzing enzyme fatty acid amide hydrolase (FAAH) in the somata of CA3 pyramidal neurons and Purkinje’s cells (Cristino et al., 2008). Despite these recent advances, little is known about the distribution and function of NAPE-PLD outside the central nervous system (CNS), for example in the autonomic and sensory peripheral nervous systems.

2.3 Degradation of Acylethanolamides As anticipated earlier, also the proteins involved in the degradation of AEs, which occurs almost uniquely via their enzymatic hydrolysis (> Figure 5-2), have been identified and cloned. FAAH, an intracellular integral membrane protein of 597 amino acids belonging to the amidase family of enzymes and characterized by the optimal pH value of 8.5–10, catalyzes the hydrolysis of AEs (Cravatt et al., 1996; Giang and Cravatt, 1997). It is widely distributed in the brain (Tsou et al., 1998), for example, in somata and dendrites of pyramidal cells of the hippocampus (Gulyas et al., 2004). Furthermore, FAAH is present in the somatodendritic compartment of principal cells, but not in interneurons (Tsou et al., 1998; Egertova´ et al., 2003; Cristino et al., 2008), and appears to be located mostly on the membrane surface of intracellular organelles known to store Ca2+ (e.g., mitochondria, smooth endoplasmic reticulum) as demonstrated by means of ultrastructural analysis (Gulyas et al., 2004). In the cerebellum, Purkinje cells and their dendrites are intensively FAAH-immunoreactive, and so is a sparse axon plexus at the border of the Purkinje cells/granule layers (Cristino et al., 2008). To date, evidence indicates that FAAH is primarily a postsynaptic enzyme, very often coexpressed with TRPV1 receptors (Cristino et al., 2008) and complementary to axon terminals that express cannabinoid CB1 receptors

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(Egertova´ et al., 2003). This evidence suggests a role for FAAH in the regulation of either presynaptic AEA acting at CB1 receptors, or postsynaptic AEA and AEs acting at TRPV1 channels. Another FAAH enzyme, named FAAH-2, was recently cloned (Wei et al., 2006). The gene that encodes this enzyme was found in several primates, marsupials, and more distantly related vertebrates, but not in a number of lower placental mammals, including mouse and rat. FAAH-2 shares 20% sequence identity with FAAH, hydrolyzes primary fatty acid amide substrates (e.g., oleamide) at equivalent rates as FAAH, but exhibits much lower affinity for AEA and PEA and similar affinity for OEA. Both enzymes were sensitive to the principal classes of FAAH inhibitors synthesized to date, including O-aryl carbamates and a-keto heterocycles. Interestingly, the C-terminal catalytic domain of FAAH-2 appears to be located in the luminal compartments of the cell, as opposed to that of FAAH, which is located in the cytosol. However, the virtual absence of FAAH-2 expression in the human brain suggests that this enzyme does not play a major role in CNS lipid signaling (Wei et al., 2006). Another enzyme not related to FAAH, with instead some structure homology to ceramidase and belonging to the family of choloylglycine hydrolases, was recently cloned and found to hydrolyze preferentially PEA and only to a low extent OEA and AEA (Ueda et al., 2001). The enzyme was named N-acylethanolamine-hydrolyzing acid amidase (NAAA), and is highly expressed in macrophages and the lungs, as well as in various rat tissues including the brain (Tsuboi et al., 2005, 2007). It is characterized by an optimal pH of 5, is activated by self-catalyzed proteolysis and is stabilized by N-glycosylation of Asn-37, Asn-107, Asn-309, and Asn-333 (Zhao et al., 2007). Its distribution in the brain has not been studied yet.

2.4 Transport of Acylethanolamides across the Plasma Membrane In order to interact with some of its targets (for example the cannabinoid receptors), AEA needs to be released from cells (> Figure 5-2). AEA transport across the plasma membrane occurs as an immediate consequence of its de novo biosynthesis from NArPE and its increased intracellular versus extracellular concentrations. Likewise, since AEA-hydrolyzing enzymes are intracellular, this AE needs to be transported into the cell in order to be inactivated, and again this transport occurs as a consequence of its higher extracellular versus intracellular concentration. Strong, although still controversial, indirect evidence, based on biochemical, pharmacological, and immunohistochemical techniques, suggests that AEA transport across the plasma membrane does not occur via simple passive diffusion, but is instead facilitated by specific membrane and/ or intracellular proteins. Thus, while AEA cellular reuptake is driven by its intracellular hydrolysis, but still requires proteins different from FAAH, AEA release seems to occur via a mechanism that is sensitive to the same inhibitors that block cellular reuptake, and is driven by AEA de novo biosynthesis (Di Marzo et al., 1994; Hillard et al., 1997; Deutsch et al., 2001; Ligresti et al., 2004; Hillard et al., 2007). Unfortunately, little progress has been made toward the molecular identification of the proteins that facilitate AEA transport across membranes, although it seems clear that AEA release from postsynaptic neurons can be regulated by elevated electrical activity (Adermark and Lovinger, 2007), and that nitric oxide as well as the presence of lipid rafts instead regulate AEA cellular uptake (Maccarrone et al., 2000; McFarland et al., 2004). It has also been established that the mechanism(s) through which AEA and PEA are taken up by cells are pharmacologically distinguishable, whereas OEA and AEA seem to use mechanisms with very similar pharmacology (Bisogno et al., 1997; Hillard et al., 1997; Hillard et al., 1997, 2007; Jacobsson and Fowler, 2001).

3

Established and Potential Molecular Targets of Acylethanolamides in the Nervous System

3.1 Introduction AEA and other AEs have been described to date to interact directly, and often in a very promiscuous way, with members of at least three of the four major classes of receptor proteins, i.e., with G-protein-coupled receptors (GPCRs), ion channels, and nuclear receptors (> Table 5-1). No example exists to date for their

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direct interaction with receptor tyrosine kinases, although some examples have been reported for direct interaction of AEs with various enzymes, including: (1) AEA capability to modulate protein kinase C – with stimulation of phosphatidylserine (PS)-induced PKC activation (EC50 = 40 mM), and inhibition of dioleylglycerol-induced potentiation of both Ca2+- and Ca2+/PS-induced PKC activation (IC50 = 8 and 30 mM, respectively (De Petrocellis et al., 1995); and (2) AEA stimulation of phospholipase D at physiological concentrations (0.1–1.0 mM) (Kiss, 1999). The physiological relevance of these two effects has never been

. Table 5-1 Proposed molecular targets for acylethanolamides in the mammalian brain Acylethanolamide Anandamide

GPCRs  CB1 agonist Ki = 89 nM Devane et al. (1992)  CB2 agonist Ki = 371 nM Devane et al. (1992)  GPR55 agonist EC50 = 18 nM Ryberg et al. (2007)

Oleoylethanolamide

 GPR119 agonist EC50 = 3.2 mM Overton et al. (2006)  GPR55 agonist EC50 = 4 nM Ryberg et al. (2007)

Palmitoylethanolamide

Linoleoylethanolamide

Linolenoylethanolamide

Ion Channels  L-type Ca2+, inhibitor IC50 = 4–40 mM Shimasue et al. (1996)  T-type Ca2+, inhibitor IC50 = 1 mM Chemin et al. (2001)  Lead K+ TASK, inhibitor IC50 = 0.7 mM Maingret et al. (2001)  Delayed rectifier K+ Kv3.1/4.3 inhibitor IC50 = 80 nM Oliver et al. (2004)  Shaker-related K+, inhibitor IC50 = 2.7 mM Poling et al. (1996)  TRPV1, agonist pEC50 = 6.4} Movahed et al. (2005)  TRPM8, antagonist No effect De Petrocellis et al. (2007)

Ligand-gated ion Channels  TRPV1 agonist pEC50 = 5.94 Smart et al. (2000)  TRPM8 antagonist IC50 = 0.15 mMa IC50 = 3.1 mMb De Petrocellis et al. (2007)  5-HT3 inhibitor IC50 = 94 nM Fan (1995)  a7nACh inhibitor IC50 = 230 nM Oz et al. (2003)  GlyR inhibitor IC50 = 200–300 nM Lozovaya et al. (2005)  GlyR agonist EC50 = 78–86 nM Hejazi et al. (2006)

Nuclear Receptors  PPAR-a agonist? Artmann et al. (2008)  PPAR-g agonist EC50 = 8 mM Bouaboula et al. (2005)

 PPAR-a agonist EC50 = 120 nM Fu et al. (2003)  PPAR-a agonist EC50 = 3.1 mM Lo Verme et al. (2005)

 TRPV1 agonist pEC50 = 6.2} Movahed et al. (2005)  TRPV1 agonist pEC50 = 6.3} Movahed et al. (2005)

continued

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. Table 5-1 (continued) Acylethanolamide 5Z,8Z,11Z-eicosatrienoylethanolamide

7Z,10Z,13Z,16Zdocosatetraenoylethanolamide

Di-homo-g-linolenoylethanolamide

GPCRs  CB1 agonist Priller et al. (1995)  CB2 agonist Priller et al. (1995)  CB1 agonist Hanus et al. (1993)  CB2 agonist Hanus et al. (1993)  CB1 agonist Hanus et al. (1993)  CB2 agonist Hanus et al. (1993)

Ion Channels

Ligand-gated ion Channels

Nuclear Receptors

 TRPV1 agonist pEC50 = 6.8 Movahed et al. (2005)

Abbreviations: a7nACh, a-7 nicotinic acetylcholine receptor; CB1, cannabinoid receptor type-1; CB2, cannabinoid receptor type-2; GlyR, glycine receptor; GPCR, G-protein-coupled receptor; GPR55, orphan G-protein-coupled receptor 55; GPR119, orphan G-protein-coupled receptor 119; PPAR-a, peroxisome proliferator-activated receptor-a; PPAR-g, peroxisome proliferator-activated receptor-g; TRPM8, transient receptor potential of melastatin type-8 channel; TRPV1, transient receptor potential of vanilloid type-1 channel a Icilin-induced TRPM8 gating of Ca2+ influx b Menthol-induced TRPM8 gating of Ca2+ influx

investigated, although AEA stimulation of PKC was suggested to underlie part of its sensitizing effect on TRPV1 receptors (Premkumar and Ahern, 2000).

3.2 Cannabinoid Receptors As mentioned earlier, the renewed interest from the scientific community in the family of the acylethanolamides originated from the discovery of AEA, which in turn was the consequence of the identification and cloning, first in the brain (Devane et al., 1988; Matsuda et al., 1990), and then in peripheral (immune) organs (Munro et al., 1993), of specific binding sites for THC, the cannabinoid CB1 and CB2 receptors. These are GPCRs, whose activation is transduced into cellular responses via a variety of intracellular signals (Howlett, 2005), including: (1) inhibition or, less often, stimulation of adenylate cyclase via Gi/o or Gs proteins, with subsequent inhibition or stimulation of protein kinase A pathways; (2) activation of mitogen-activated protein kinases via Gi/o proteins; (3) activation, mostly in the case of CB1 receptors, of phosphoinositide-3-kinase and its downstream pathways, via unidentified G-proteins, and of phospholipase C-b, via either Gq/11 or bg subunits of Gi/o proteins; and, selectively for CB1 receptors (4) direct inhibition, of various types of voltage-activated ion channels and stimulation of G-protein-activated inwardly rectifying K+ channels, as well as stimulation of type A K+ channels via inhibition of AMP levels. These pathways, the localization of CB1 and CB2 receptors, the paracrine or autocrine nature of the action

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of AEA and the other ‘‘endocannabinoid,’’ 2-arachidonoylglycerol (2-AG), and their biosynthetic mechanisms confer to this ‘‘endocannabinoid’’ signaling the general property of being activated ‘‘on demand’’ (also 2-AG, like AEA, is not prestored in cells but instead is released from its biosynthetic precursor immediately prior to its release from cells), only ‘‘when and where’’ needed, generally following transient or chronic perturbation of the homeostasis of other mediators, in the attempt to restore their steady-state levels (Di Marzo and Petrosino, 2007). Thus, CB1 receptors regulate neurotransmitter release, thereby controlling also neuropeptide release from neurons (Matias and Di Marzo, 2007), whereas both CB1 and CB2 regulate cytokine and inflammatory mediator expression and release from immune cells (Klein, 2005) and hormone expression and release from hypothalamic and endocrine cells (Pagotto et al., 2006). Of all AEs, only AEA, and perhaps other ‘‘anandamide-like’’ AEs, such as 5Z,8Z,11Z-eicosatrienoylethanolamide, di-homo-g-linolenoyl-ethanolamide and 7Z,10Z,13Z,16Z-docosatetraeonyl-ethanolamide (see earlier), but not PEA and OEA, are capable of directly activating CB1 and CB2 cannabinoid receptors. Nevertheless, it has been suggested that ‘‘nonendocannabinoid’’ AEs might, under certain circumstances, enhance the activity of coreleased cannabinoid receptor-active AEs via ‘‘entourage effects’’ (Mechoulam et al., 1998), i.e., for example, by inhibiting their inactivation by FAAH, either by substrate competition, as in the case of OEA and lynoleoylethanolamide (Maurelli et al., 1995), or by suppressing FAAH expression, as in the case of PEA (Di Marzo et al., 2001a). Also SEA was shown to enhance AEA-induced and CB1mediated inhibition of adenylate cyclase in mouse cortical slices via an unknown mechanism (Maccarrone et al., 2002a, 2002b). CB1 receptors are perhaps the most abundant GPCRs in the mammalian brain, and appear to be mostly located presynaptically and to inhibit both excitatory and inhibitory neurotransmitter release within the framework of both short- and long-term synaptic plasticity, once they are activated by postsynaptically released endocannabinoid acting in a retrograde manner (Kreitzer and Regehr, 2001; Ohno-Shosaku et al., 2001; Wilson and Nicoll, 2001; Chevaleyre et al., 2006). Yet, between 2-AG and AEA, the former compound has been suggested to be playing this function in most cases, at least in the adult brain (Szabo et al., 2006). However, recent evidence suggests that AEA might act as the retrograde mediator in the emergence of striatal long-term depression in the postnatal brain (Ade and Lovinger, 2007).

3.3 ‘‘Orphan’’ G-Protein-Coupled Receptors Recently, it has been suggested that some ‘‘orphan’’ GPCRs might represent specific molecular targets for some AEs. In particular, GPR55 was found to be activated by submicromolar concentrations of cannabinoid-like synthetic and natural molecules (Johns et al., 2007) as well as AEA and PEA (but much less so OEA) (Ryberg et al., 2007). This receptor seems to be heterogeneously distributed in the human brain, with very high levels in the caudate nucleus and putamen, but its physiological role, which is likely to be mediated by Gq/11 activation and mobilization of intracellular Ca2+, has not been clarified yet. However, there seems to be no general consensus regarding the pharmacology of GPR55, with three different groups having published so far often qualitatively different results. In particular, Oka et al. (2007) could not confirm that GPR55 is activated by AEA and PEA, and proposed instead lysophosphatidylinositol as the true ligand for this receptor. Another orphan GPCR, GPR119 was shown to be activated with micromolar potency by OEA, whereas other AEs were significantly less potent and efficacious (PEA > SEA > > AEA) (Overton et al., 2006). Also this receptor, which is most abundant in the pancreas, is expressed in the brain, although again with functions that have not been fully investigated, and which definitively include inhibition of appetite. Unlike CB1 and CB2, GPR119 seems to be mostly coupled to stimulation of adenylate cyclase via the Gs protein. No studies have been performed so far conclusively demonstrating, for example, by using GPR55- or GPR119-null mice, that some of the pharmacological actions of AEs are indeed mediated by these two receptors.

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3.4 Ion Channels: TRPV1 and TRPM8 A plethora of studies (recently reviewed by Ross, 2003; van der Stelt and Di Marzo, 2004; Starowicz et al., 2007b) have substantiated the original hypothesis (Zygmunt et al., 1999; Smart et al., 2000; see Di Marzo et al., 2001b for a commentary) that one of the preferential targets for AEA is the TRPV1 receptor, which is sensitized and then desensitized by this AE as well as by other mono and polyunsaturated AEs with 18 carbon atoms (Movahed et al., 2005). Like other TRP channels, TRPV1 is a ligand-gated nonselective cation channel, whose major function in the sensory nervous system is to transduce thermal and inflammatory pain in response to several different types of proalgesic stimuli, including acid, high temperature, and stimulation with pronociceptive mediators (Caterina et al., 1997; Tominaga and Tominaga, 2005). The detailed pharmacology of TRPV1 and ‘‘endovanilloids’’ in the CNS has been recently reviewed (Starowicz et al., 2007b; Starowicz et al., 2008a), and will not be discussed here. AEA binds to TRPV1 to an intracellular site (De Petrocellis et al., 2001), very probably the same site for the natural sensitizer of this receptor, the pungent hot chili peppers constituent, capsaicin (Jordt and Julius, 2002). Like with other TRPV1 agonists, AEA binding to TRPV1 lowers the threshold of its activation by temperature, so that physiological temperatures (as opposed to those >42 C normally required to gate the channel) can open the channel and let Ca2+ ions in, thus be ‘‘interpreted’’ as heat by the animal. When TRPV1 is expressed on the plasma membrane of neuronal cells, this results in their depolarization and the subsequent release of neurotransmitters, such as substance P and calcitonin gene-related peptide in sensory neurons, and glutamate in central neurons (Starowicz et al., 2007a, 2007b). Therefore, this TRPV1-mediated excitatory effect of AEA might oppose its CB1-mediated inhibitory one. However, when CB1 and TRPV1 receptors are expressed in the same cells, at least two types of cross talk can occur between the two receptors, especially when AEA activates CB1 first and TRPV1 later (Hermann et al., 2003). Thus, previous activation of CB1 can either inhibit or enhance activation of TRPV1, depending on whether or not the cAMP-dependent cascade has been concomitantly activated, respectively. Furthermore, cannabinoid-mediated modulation of TRPV1 receptor activation is switched from inhibition to stimulation also after exposure to high nerve growth factor (Evans et al., 2007). Finally, a recent study highlighted how, when TRPV1 receptors are instead expressed postsynaptically to presynaptic CB1 receptors, their activation by AEA can indirectly inhibit the activity of CB1 by counteracting the biosynthesis of 2-AG otherwise acting as a retrograde signal (Maccarrone et al., 2008). Independently from CB1 receptors, AEA was also shown to act as an intracellular signal amplifying Ca2+ influx via TRPV1 following depletion of intracellular Ca2+ stores in sensory neurons of the dorsal root ganglia, thus favoring intracellular store replenishment (van der Stelt et al., 2005). In vivo, several effects of AEA have been associated with its capability of activating neuronal TRPV1 channels and span from the regulation of pain at the peripheral, spinal, and supra-spinal level, body temperature, movement, anxiety, cardiovascular tone, respiration, and emesis (see Starowicz et al., 2007b, 2008a for reviews). However, paradoxically, AEA might exert similar actions also by activating CB1 receptors, and only few studies have been carried out to date with AEA in TRPV1( / ) mice to conclusively demonstrate that any pharmacological effect of this compound is exclusively mediated by either receptor. This applies also to OEA, for which, however, it has been shown that its acute anorexic effects are absent in TRPV1( / ) mice (Wang et al., 2005). A recent report suggested that AEA could also interact with another TRP channel expressed in sensory neurons, the transient receptor potential of melastatin-type 8 (TRPM8), which is activated by temperatures Figure 5-1) and their metabolic enzymatic machinery, which is shared in part with both OEA and PEA, and the cannabinoid CB1 receptors, have been detected in all central and peripheral tissues involved in the control of energy intake, processing and storage, including the hypothalamus (Di Marzo et al., 2001c), the nucleus accumbens (Berrendero et al., 1998), the vagus nerve and the nodose ganglion (Burdyga et al., 2004), and the myenteric neurons, and epithelial cells of the large intestine (Coutts and Izzo, 2004). Strong evidence for the presence of this system also in nonneuronal cells and tissues, such as the liver and hepatocytes (Osei-Hyiaman et al., 2005a), the white adipose tissue (Engeli et al., 2005), the adipocytes (Bensaid et al., 2003; Cota et al., 2003; Matias et al., 2006; Roche et al., 2006), the skeletal muscle (Cavuoto et al., 2007), and the pancreas (Starowicz et al., 2008b) has also been published, thus suggesting that endocannabinoids also control energy storage and consumption and not just intake and assimilation. Indeed, CB1 receptors and endocannabinoids control energy homeostasis via both central and peripheral mechanisms, as they stimulate lipogenesis and fat accumulation in adipocytes (Cota et al., 2003; Matias et al., 2006), whereas in the liver they enhance fatty acid synthesis by inhibiting the AMP kinase (Kola et al., 2005) and upregulating the

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expression of the transcription factor sterol-responsive element binding protein 1c (SREBP-1c) and subsequently of its target genes acetylCoA-carboxylase and fatty acid synthase (Osei-Hyiaman et al., 2005a). Some of these peripheral effects are also exerted in the hypothalamus, where THC and endocannabinoids stimulate fatty acid synthase, an enzyme that plays a pro-orexigenic role (Osei-Hyiaman et al., 2005b). However, unlike the liver, endocannabinoids and THC stimulate the AMP kinase in the hypothalamus, again with a subsequent pro-orexigenic effect (Kola et al., 2005). As reviewed in detail recently by Matias and Di Marzo (2007), most of the central orexigenic effects of endocannabinoids are exerted, however, by exploiting the capability of CB1 receptors of either tonically inhibiting the expression of anorectic neuropeptides (Cota et al., 2003; Osei-Hyiaman et al., 2005a) or inhibiting excitatory and inhibitory neurotransmission release onto anorexic or orexigenic neurons, respectively (Di et al., 2003; Jo et al., 2005; Tasker, 2006). Unlike the other major endocannabinoid 2-AG, AEA levels are not increased in the hypothalamus following food deprivation, and are not decreased following food consumption (Kirkham et al., 2002). However, following food-deprivation, AEA levels do increase in the duodenum (Gomez et al., 2002) and limbic forebrain (Kirkham et al., 2002), and this phenomenon was suggested to participate in CB1-mediated inhibition of satiety via capsaicin-sensitive sensory fibers terminating in the brainstem and in CB1-mediated stimulation of the hedonic aspects of food intake, respectively. Furthermore, AEA, unlike 2-AG, does not seem to participate in those types of retrograde signaling at CB1 receptors that are involved also in the control of orexigenic and anorexic mediator release in the hypothalamus. Thus, it is possible that rather than contributing to the hypothalamic control of appetite, AEA is involved in inhibition of satiety and stimulation of the motivation to consume palatable foods (Matias and Di Marzo, 2007). However, this hypothesis still awaits further investigations in order to be corroborated or discarded. Unlike AEA, OEA exerts anorexic actions (Rodriguez de Fonseca et al., 2001; Fu et al., 2003), which appear to be mediated at least in part by PPARa (Fu et al., 2003). In fact, chronic OEA fails to cause satiety and to reduce body weight in PPAR-a knockout mice (Fu et al., 2003). Again, unlike AEA, OEA seems to act mostly at the peripheral level (Rodriguez de Fonseca et al., 2001). In fact, the effect of intestinal OEA results in central actions that could be erased after destruction of capsaicin-sensitive peripheral fibers, thus suggesting that this compound acts as a peripheral inhibitor of centrally controlled food ingestion behavior. However, this latter finding is also in agreement with the proposal that another molecular target participates in OEA anorexic effects, i.e., the TRPV1 channel, which is activated and desensitized by capsaicin, and which OEA activates with moderate potency and good efficacy (Ahern, 2003; Movahed et al., 2005; Wang et al., 2005). Indeed, as suggested by studies with knockout mice, it is possible that while PPAR-a mediates the chronic anorectic effects of OEA, TRPV1 is responsible for its acute effects (Wang et al., 2005). Another target of OEA, GPR119, also participates in food-intake, as shown by the fact that synthetic agonists of this receptor, like OEA, exhibit anorectic properties (Overton et al., 2006). However, no studies in GPR119( / ) mice have been carried out with OEA. In a comprehensive study, Proulx et al. (2005) provided evidence that OEA suppresses feeding in rats without causing visceral illness, conditioned taste aversion and sodium appetite, and that neither ghrelin, peptide YY, glucagon-like peptide 1, apolipoprotein A-IV, nor CCK play a role in its effects. The authors concluded that despite the fact that OEA-induced anorexia is unlikely to be due to impaired motor activity, caution should be used when interpreting how specific the behavioral and metabolic effects of OEA are. In a previous study, however, OEA was found to decrease plasma ghrelin in both fasted and fed rats (Cani et al., 2004). Interestingly, despite their opposite effects on food intake, AEA and OEA share the capability of delaying both gastric emptying and intestinal motility (Capasso et al., 2005; Aviello et al., 2008; Di Marzo et al., 2008). While in the case of OEA delayed gastric emptying, which however was not affected by CB1, TRPV1, or PPAR-a antagonists, might contribute to its anorectic actions (Aviello et al., 2008), in the case of AEA, delayed intestinal motility (which, like its effect on gastric emptying, is accounted for by activation of prejunctional CB1 receptors on mesenteric fibers) might participate in increased nutrient assimilation. Surprisingly, PEA shares with OEA the capability of delaying intestinal motility but not gastric emptying (Capasso et al., 2005; Aviello et al., 2008). Like with AEA, these gastrointestinal effects of OEA and PEA are probably exerted tonically, since pharmacological inhibition of their degradation by FAAH also

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delays gastric emptying and retards intestinal motility in a way only partly antagonized by CB1 receptor antagonism (Capasso et al., 2005; Aviello et al., 2008). However, it is not known whether also OEA and PEA, like AEA, exert these effects by acting at mesenteric neurons, and through which receptors. OEA and AEA levels are differently regulated in the small intestine, where a sevenfold increase of AEA levels (Gomez et al., 2002) and a marked decrease of OEA (Rodriguez de Fonseca et al., 2001; Fu et al., 2003) are observed following food deprivation in comparison to ad lib-fed control rats. By contrast, no changes in brain, liver and heart OEA levels are found following food-deprivation (Fu et al., 2007). Recently, a 300-fold increase of OEA levels in the small intestine of fed compared with fasted Burmese pythons was also reported (Astarita et al., 2006). Two studies investigating the biochemical mechanisms underlying this phenomenon have been published, although with discrepant results. In one study, feeding was found to increase the levels of OEA and other unsaturated AEs in the duodenum and jejenum of rats without affecting those of saturated AEs, including PEA (which exerts no effect on food intake), nor the levels of the NAPE precursors for OEA and PEA, thus underscoring the selectivity of OEA in food-intake (Fu et al., 2007). Increased OEA levels during feeding was accompanied by an increase of the expression and the activity of the NAPE-PLD and by a decrease of the expression and the activity of FAAH (Fu et al., 2007). Opposing results were obtained in a previous study by Petersen et al. (2006), who showed that the activity of biosynthetic and degrading enzymes did not change during food deprivation and refeeding, and that differential changes in AEA, OEA, and PEA small intestine levels were likely due to differential changes in the levels of their NAPE precursors. The reasons why these studies produced discrepant outcomes are to date still unknown. Nevertheless, these data still emphasize the fact that the levels of different AEs can be regulated in different and even opposing ways during food-intake and deprivation. The resulting activation/inactivation of CB1 and PPAR-a receptors on sensory fibers connecting the gut with the brainstem might result in opposing effects on satiety. Importantly, perhaps in agreement with the opposing regulation of AEA and OEA following food deprivation/consumption, combined chronic treatment of rats with OEA and an antagonist of AEA and endocannabinoid actions at CB1 receptors was recently found to result in synergic inhibitory effects on food intake (Serrano et al., 2008). However, in human beings, AEA, OEA, and PEA levels in the blood decrease after food consumption and are permanently elevated in the blood of type 2 diabetes patients (Matias et al., 2007), thus possibly suggesting that the peripheral metabolism of these three AEs is under the tonic negative control of insulin. Also SEA was shown to cause food-intake inhibitory effects. The anorectic effect of this compound, which exhibits little if any activity at PPAR-a, TRPV1, and CB1 receptors, was accompanied downregulation of liver stearoyl-CoA desaturase-1 (SCD-1) mRNA expression (Terrazzino et al., 2004). The exact molecular mechanism through which SEA exerts anorectic effects has not yet been investigated.

4.3 Role in Neuroprotection and Neuroinflammation The potential neuroprotective actions of AEs have been known for a long time (see Hansen et al., 2002; Fowler, 2003 for reviews) and are also strongly suggested by the repeated finding of their accumulation in the brain following ischemic conditions (Berger et al., 2004; Degn et al., 2007). Recently, the potential for a neuroprotective and anti-inflammatory role of AEA and AEs has been revisited in view of their actions at their proposed molecular targets, i.e., cannabinoid, TRPV1, and PPAR receptors. The important function of CB1 and CB2 receptors and their endogenous ligands, including AEA, in neuroprotection has been recently reviewed (Bisogno and Di Marzo, 2007; Centonze et al., 2007) and will not be discussed here in detail. In particular, it has been pointed out that: (1) the endocannabinoid system is activated ‘‘on demand’’ as an early adaptive response to excitotoxic (i.e., excessive glutamate release), neurotoxic and neuroinflammatory stimuli generated during hypoxia/ischemia and traumatic brain injury, or unbalanced neuronal activity and neural lesions typical of acute neurological conditions and neurodegenerative disorders; (2) this protective response is initially tightly ‘‘time- and space-dependent,’’ although it might loose specificity with the continuation and chronicization of the pathological condition – when this happens, especially in some neurodegenerative disorders, the endocannabinoids and CB1 and CB2 receptors might start contributing to their late symptoms; and (3) sometimes AEA might play a role different from that of

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2-AG during these neurological disorders in view of its capability of activating and eventually desensitizing the TRPV1 receptor, which is expressed in the brain and seems to be involved in the control of both ischemia and excitotoxicity (see van der Stelt and Di Marzo, 2005, and Kim et al., 2007, for reviews), as well as of motor activity in animal models of Parkinson’s (Lee et al., 2006; Morgese et al., 2007) and Huntington’s (Lastres-Becker et al., 2003) disease and of multiple sclerosis (Cabranes et al., 2005). Both agonists and antagonists of TRPV1 receptors counteract the formation of edema in an in vivo animal model of excitotoxicity (Veldhuis et al., 2003), thus suggesting that TRPV1, possibly by participating in glutamate release and in neuroinflammatory processes, contributes to the damage induced by excitotoxic drugs, and that the reason why agonists are beneficial is because they are capable of pharmacologically desensitizing these channels. Thus, one can hypothesize for de novo biosynthesized AEA acting at TRPV1 receptors during excitoxicity both a protective and counterprotective role, depending on whether or not it immediately desensitizes these receptors. Since the formation of AEA during excitotoxic conditions is often accompanied by that of OEA and PEA, which either directly or indirectly can influence the activity of TRPV1 receptors, it is likely that also these and other AEs participate in these processes. Indeed: (1) PEA was described to reduce seizures in an animal model of epilepsy (Lambert et al., 2001; Sheerin et al., 2004), and to exert neuroprotective effects against neuronal oxidative stress (Lombardi et al., 2007), and in some in vitro models of excitotoxicity (Skaper et al., 1996) but not in others (Andersson et al., 2000; Lombardi et al., 2007), via as yet undefined molecular targets; and (2) OEA was recently shown to reduce infarct volume after middle cerebral artery occlusion in mice, although this effect was shown to be mediated by PPAR-a receptors as it was not observed in PPAR-a( / ) mice (Sun et al., 2007). Indeed, PPAR-a has been recently implicated in several types of brain protective responses against neurological conditions including stroke and neurodegenerative disorders (see Bordet et al., 2006, for a recent review). Furthermore, activation of this receptor inhibits several types of inflammatory responses as well as autoimmune responses such as those contributing to multiple sclerosis (see later, and Rizzo and Fiorucci, 2006; Racke et al., 2006 for reviews). Fenofibrates, the prototypical activators of PPAR-a, inhibit microglia-mediated neuroinflammatory responses (Xu et al., 2005), exert protective and neurological recovery-promoting actions in traumatic brain injury (Besson et al., 2005; Chen et al., 2007), protect against oxidative stress and inflammatory response evoked by transient cerebral ischemia/ reperfusion (Collino et al., 2006) and prevent the MPTP-induced dopaminergic cell loss in the substantia nigra pars compacta, and whilst attenuating the loss of tyrosine hydroxylase immunoreactivity in the striatum of mice (Kreisler et al., 2007). Therefore, one should expect that both OEA and PEA (which despite its lower potency at PPAR-a possesses stronger anti-inflammatory effects than OEA, see later) exert neuroprotective actions in a number of neurological conditions during which their endogenous levels are elevated. Specific studies addressing this possibility need to be performed in parallel with strengthening the hypothesis that PPAR-a can be a good target for treating acute and chronic CNS pathological states. The same applies to PPAR-g, which is also implicated in neuroprotective mechanisms (Kapadia et al., 2008), although this receptor is activated only by high micromolar concentration of AEA (Bouaboula et al., 2005).

4.4 Control of Nociception Of the three most studied AEs, i.e., AEA, PEA, and OEA, the former two are the ones that have been tested in almost all models of acute, inflammatory, visceral and neuropathic pain, with emphasis on both peripheral and spinal–supraspinal mechanisms of action (Mazzari et al., 1996; Lambert et al., 2002; Rice et al., 2002; Darmani et al., 2005; Walker et al., 2005; Hohmann and Suplita, 2006; Re et al., 2007; Rea et al., 2007). Furthermore, the tissue levels of these compounds in tissues involved in nociception (skin, spinal cord, periaqueductal grey, rostral ventromedial medulla, etc.) have been shown to undergo changes following the administration of acute and nociceptive stimuli in both laboratory animals and humans (Walker et al., 1999; Darmani et al., 2005; Jhaveri et al., 2006; Agarwal et al., 2007; Degenhardt et al., 2007; Petrosino et al., 2007). We shall limit our discussion here to the role of these compounds in neurogenic inflammation and in spinal and central mechanisms of nociception.

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AEA exerts most of its analgesic actions by stimulating CB1 receptors, whose activation, via inhibition of substance P and CGRP release from sensory fibers, and of glutamate from spinal cord neurons, is widely accepted as one of the major strategies to inhibit nociception (see Maione et al., 2006a, for review). By contrast, AEA can contribute to pain transmission by activating TRPV1 receptors, particularly under conditions of inflammatory pain, when the sensitivity of these channels to AEA is sensibly increased (see Starowicz et al., 2007b, for review). Recent data, however, suggest that by activating TRPV1 receptors on excitatory antinociceptive output neurons of the ventrolateral periaqueductal grey, AEA can also inhibit nociception by stimulating the descending antinociceptive pathway and activating OFF neurons in the rostral ventromedial medulla (Maione et al., 2006b; Starowicz et al., 2007a). By converse, AEA can cause hyperalgesia by activating CB1 receptors on glutamatergic neurons innervating the excitatory antinociceptive output neurons, and counteract nociception by activating CB1 receptors on GABAergic interneurons innervating these same output neurons (Maione et al., 2006b). Finally, intrathecal AEA can also cause analgesia by activating/desensitising TRPV1 receptors potentially located on glutamatergic pronociceptive neurons of the spinal cord (Di Marzo et al., 2000b; Horvath et al., 2008). In conclusion, AEA can variedly affect pain by interacting with both CB1 and TRPV1 receptors. It has been shown that also OEA can cause visceral nociception by activating TRPV1 (Wang et al., 2005). The anti-inflammatory (Lo Verme et al., 2005) and part of the antinociceptive (Lo Verme et al., 2006) effects of PEA seem instead to be mostly mediated by its activity at PPAR-a. Interestingly, PEA activity at this nuclear receptor is efficacious enough to induce these actions but not to induce anorexia, whereas OEA, which is more potent as a PPAR-a activator, exerts weaker antinociceptive effects than PEA against visceral and inflammatory pain (Suardı´az et al., 2007) and carrageenan-induced paw edema (Wise et al., 2008). However, the antinociceptive effects of OEA are not mediated by PPAR-a or TRPV1 receptors, and appear to involve the participation of glutamatergic transmission (Suardı´az et al., 2007). As to the analgesic actions of PEA, in the animal model of acute/inflammatory pain induced by formalin, they seem to be mediated, downstream to PPAR-a, also by calcium-operated K(Ca)3.1 and K(Ca)1.1 potassium channels and reduced firing rate of DRG neurons, which were shown to express this nuclear receptor (Lo Verme et al., 2006). However, it has not been clarified yet how the usually long-term effects that follow the activation of nuclear receptors underlie these acute effects of PEA. It is possible that not all analgesic effects of PEA are due to activation of PPAR-a, as recently suggested by Wallace et al., 2007, who found that the antihyperalgesic actions of a metabolically stable PEA analog, palmitoylallylamide in various rat models of neuropathic pain were not always blocked by a PPAR-a antagonist. Other possible mechanisms for the analgesic actions of this compound might involve entourage effects at CB1 and TRPV1 receptors (Re et al., 2007), activation of CB2-like receptors (Farquhar-Smith et al., 2002) and, in the case of inflammatory pain and neurogenic inflammation, where local vasodilation plays an important role, GPR55 (Ryberg et al., 2007). Finally, while PEA effects at the level of neurogenic inflammation and peripheral pain control are likely to be caused, to a large extent, by interference with mast cell hyperactivity (Mazzari et al., 1996; Re et al., 2007), the compound was recently shown to inhibit the peripheral inflammation caused by carrageenan also when administered intracerebroventricularly, via acute activation of central PPAR-a and subsequent inhibition of nuclear factor-kappa B activation in the spinal cord (D’Agostino et al., 2007). In summary, AEs have several ways to influence pain perception, possibly through several mechanisms of action, some of which have not been identified yet. For this reason, FAAH inhibitors (which, however, might act also via other fatty acid amides) have proved to be very efficacious in several experimental models of pain, and are being developed as new analgesic and anti-hyperalgesic drugs (Jhaveri et al., 2007).

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Conclusions

As reviewed in this chapter, AEs are emerging as a very important, albeit certainly not novel, class of brain lipid mediators, with key roles in the control of energy intake and processing, in nociception and in the protection from neuronal damage and inflammation. While considerable progress has been made to understand the molecular mechanisms underlying the regulation of their levels, and the neuropathological conditions in which such regulation occurs, information on the mode of action of most of these

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compounds is still incomplete. The identification of the possibly more than one receptor for each of the most abundant AEs will certainly lead to new important discoveries in molecular neurobiology and open new avenues in the development of novel therapeutic strategies against metabolic and neurological disorders and disabilitating pain.

Acknowledgments The authors are grateful to Epitech group S.r.l. for continued support.

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Wang J, Okamoto Y, Morishita J, Tsuboi K, Miyatake A, et al. 2006. Functional analysis of the purified anandamidegenerating phospholipase D as a member of the metallobeta-lactamase family. J Biol Chem 281: 12325-12335. Wang X, Miyares RL, Ahern GP. 2005. Oleoylethanolamide excites vagal sensory neurones, induces visceral pain and reduces short-term food intake in mice via capsaicin receptor TRPV1. J Physiol 564: 541-547. Wei BQ, Mikkelsen TS, McKinney MK, Lander ES, Cravatt BF. 2006. A second fatty acid amide hydrolase with variable distribution among placental mammals. J Biol Chem 281: 36569-36578. Wilson RI, Nicoll RA. 2001. Endogenous cannabinoids mediate retrograde signalling at hippocampal synapses. Nature 410: 588-592. Wise LE, Cannavacciulo R, Cravatt BF, Martin BF, Lichtman AH. 2008. Evaluation of fatty acid amides in the carrageenan-induced paw edema model. Neuropharmacology 54: 181-188. Xu J, Storer PD, Chavis JA, Racke MK, Drew PD. 2005. Agonists for the peroxisome proliferator-activated receptor-alpha and the retinoid X receptor inhibit inflammatory responses of microglia. J Neurosci Res 81: 403-411. Zhao LY, Tsuboi K, Okamoto Y, Nagahata S, Ueda N. 2007. Proteolytic activation and glycosylation of N-acylethanolamine-hydrolyzing acid amidase, a lysosomal enzyme involved in the endocannabinoid metabolism. Biochim Biophys Acta 1771: 1397-1405. Zygmunt PM, Petersson J, Andersson DA, Chuang H, Sørga˚rd M, et al. 1999. Vanilloid receptors on sensory nerves mediate the vasodilator action of anandamide. Nature 400: 452-457.

6

Chemistry, Tissue and Cellular Distribution, and Developmental Profiles of Neural Sphingolipids*

G. Tettamanti . L. Anastasia

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100

2 2.1 2.2 2.2.1 2.2.2 2.2.3 2.2.4 2.3 2.3.1 2.3.2 2.3.3

Chemical Structure of Sphingolipids of the Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100 Simple Sphingolipids: Sphingoid Bases and Ceramide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100 Complex Sphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 Gangliosides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 Sulfatides and Other Sulfo-Glycosphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 110 Neutral Glycosphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 110 Cationic Glycosphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Synthetic Sphingolipids, Derivatives of Sphingolipids and Neo-Sphingolipids . . . . . . . . . . . . . . . . . 111 Chemical Synthesis of Sphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Chemical Synthesis of Sphingolipid Derivatives Useful for Biological Investigation . . . . . . . . . . . 113 Chemical Synthesis of Sphingolipid Analogs, Unnatural Sphingolipids and Neo-Sphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118

3

Compositional and Developmental Profiles of Sphingolipids in the Nervous System of Different Animals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122 Analytical Approaches for the Detection, Structural Characterization and Quantification of Sphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123 Analytical Biochemistry of Sphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123 Immunochemical Methods for ‘‘In Situ’’ Detection of Sphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . 126 Regional Cellular and Subcellular Localization of Sphingolipids in the Nervous System of Different Animals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 128 Sphingolipid Composition and Distribution in the Central and Peripheral Nervous Tissue of Vertebrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 128 Regional, Cellular and Subcellular Localization of Sphingolipids in the Nervous Tissue of Vertebrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 Development Profiles of Sphingolipids in the Nervous System of Different Animals . . . . . . . . . . 142 Developmental Profiles of the Main Sphingolipid Components of Brain . . . . . . . . . . . . . . . . . . . . . . 142 Developmental Profiles of Individual Gangliosides and Other Glycosphingolipids . . . . . . . . . . . . 144 Developmental Changes of the Fatty Acid and Long Chain Base Composition of Individual Sphingolipids, Particularly Gangliosides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152

3.1 3.1.1 3.1.2 3.2 3.2.1 3.2.2 3.3 3.3.1 3.3.2 3.3.3

4

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 154

* This review is dedicated to the memory of Prof. Lars Svennerholm

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_6, # Springer ScienceþBusiness Media, LLC 2009

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Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

Abstract: Sphingolipids constitute a class of lipids characterized by the presence of a long chain aminoalcohol (sphingoid base) and are particularly abundant in the nervous system. They include simple molecular species(sphingosine - or sphinganine -, sphingosine-1-phosphate, ceramide, ceramide-1-phosphate), and the ceramide containing complex sphingolipids (sphingomyelin, cerebrosides, sulfatides, neutral glycosphingolipids, acidic glycosphingolipids-gangliosides-, etc.). First, the chemical details are reported of both the naturally occurring sphingolipids, mostly present at the level of cellular membranes, and the synthetic sphingolipids, derivatives of sphingolipids, and mimetics of sphingolipids, that are extremely useful for biological investigations. Owing to the compositional complexity of sphingolipids, the analytical approaches employed for their detection, structural characterization, quantification, and “in situ” detection, are also briefly reviewed, in order to provide a basic and rationale background to investigators interested in the field. Then, the compositional profiles of sphingolipids in the nervous system of different animals, with particular emphasis to humans, are described, illustrating the analogies and differences, with regard to regional, cellular and subcellular localization of the individual sphingolipid species, with special attention to gangliosides, that display the wider array of composition. The differences in the long chain base and fatty acid composition, together with those in the saccharide composition in glycosphingolipids are also outlined, as a necessary chemical premise to understand the intricacy of the related metabolic pathways and to acknowledge the specifically distinct features of their functional implications. Finally, the developmental profiles of sphingolipids in the course of neural development and ageing in the different animals are described, illustrating common trends and peculiar differences among animals.

1

Introduction

Sphingolipids, phosphosphingolipids and glycosphingolipids, constitute a family of molecules particularly abundant in the nervous system. The interest in sphingolipids is increasing at a rapid rate, as documented by the thousands of published papers per year in the last 4–5 years. There are solid reasons for this interest. First of all, sphingolipids offer an extremely diversified molecular array, owing to the wide variety of both the saccharide structures present in the hydrophilic head group of glycosphingolipids, and the fatty acids and long chain bases (sphingosines and sphinganines), components of ceramide, the hydrophobic tail group of all sphingolipids. As sphingolipids are almost exclusively membrane components, particularly of the plasma membranes, they are suitable to interact with external ligands or other cells though their hydrophilic groups, and with protein and lipid partners of the membrane through their hydrophobic groups. In both cases they can affect the pathways of trans-membrane signaling. A second reason relies on the evidence that under proper stimulations some sphingolipids, namely phosphosphingolipids, can be induced to release ceramide with formation of potent bioregulators, i.e., ceramide itself, ceramide-1-phosphate, sphingosine and sphingosine-1-phosphate. A further reason is that the absence of genes for some of the enzymes involved in sphingolipid degradation gives rise to very severe syndromes, the sphingolipidoses. The aim of this review is not only to provide an update on some aspects of sphingolipids neurobiology, but also to recall the main historical findings that still are the fundamental basis for further studies, and to stimulate reflections and directions for further enterprises. This aim includes the intent to supply a platform of fundamental knowledge to ‘‘newcomers’’ in the field, who have to be properly introduced and oriented. The review dedicates particular attention to the intricate chemistry of sphingolipids, the analytical aspects of the investigations on sphingolipids, the basics of sphingolipid distribution in neural cells and tissues, and behavior during neural differentiation, maturation and ageing.

2

Chemical Structure of Sphingolipids of the Nervous System

2.1 Simple Sphingolipids: Sphingoid Bases and Ceramide The sphingoid long chain bases most frequently occurring in the nervous system are C-18 (2-amino-1,3dihydroxy-octadec-4ene) and C-20-sphingosine (2-amino-1,3-dihydroxy-eicos-4ene), carrying a double

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6

bond in the C4–C5 position, followed, generally in small proportions, by C-18 and C-20 sphinganine, lacking the C4–C5 double bond (Karlsson, 1970). All of them are in the trans-D-erythro form, and assume the 2S,3R steric configuration (Carter and Fujino, 1956). The sphingoid long chain bases can be linked by an amide bond to a fatty acid, producing ceramide that constitutes the hydrophobic portion, or tail, of complex sphingolipids. The fatty acids have a long, even a very long, carbon chain, the predominant species being palmitic acid (C16:0) and stearic acid (C18:0); hydroxylated forms of fatty acids are also frequently found (Karlsson, 1970). Exceptional is the occurrence of the N-acetylated form of long chain bases (C2-ceramide) (Lee et al., 1996). Besides being part of complex sphingolipids, both long chain bases and ceramides occur in the cells in free forms and in the phosphorylated forms at the primary alcoholic group (C1), namely sphingosine-1-phosphate (Sph-1-P) (Lee et al., 1998) and ceramide-1-phosphate (Cer1-P) (Dresslers and Kolesnick, 1990). Sphingosine is also present as the monomethyl or dimethyl derivative of the amino group (monomethyl-sphingosine, dimethyl-sphingosine) (Igarashi and Hakomori, 1989), or glycosidically linked at C1 to galactose (galactosyl-sphingosine, Gal-Sph) (Igisu and Suzuki, 1984), glucose (glucosyl-sphingosine, Glc-Sph) (Barranger and Ginns, 1989), or choline-phosphate (sphingosylphosphocholine, Sph-P-Chol) (Desai and Spiegel, 1991). O-acylated forms of N-acetylsphingosine were also detected (Abe et al., 1996). The chemical features of the most represented simple sphingolipids are reported in > Figure 6-1. A more detailed description of the chemical characteristics of simple sphingolipids is easily available (Heller and Kronke, 1984; Kolesnick, 1992; Hakomori and Igarashi, 1993; Hannun and Bell, 1993; Spiegel and Milstien, 1995; Pyne and Pyne, 2000).

2.2 Complex Sphingolipids Ceramide is attached through its primary alcoholic function to: (1) phosphocholine, giving origin to the phospholipids of the sphingomyelin family, (2) a saccharide structure, producing two distinct families of glycosphingolipids: (a) the neutral glycosphingolipids, that carry a single monosaccharide, glucose or galactose (it is the case of cerebrosides, gluco-cerebroside or galacto-cerebroside) or a neutral oligosaccharide chain, and (b) the acidic glycosphingolipids that carry an oligosaccharide containing sialic acid (gangliosides), sulfate (sulfatides and other sulfoglycolipids), or glucuronic acid. The phosphocholine or saccharide portion of complex sphingolipids constitutes the hydrophilic head of the molecule. The terms sphingomyelin, cerebrosides and gangliosides reflect the abundance of these compounds in brain structures (white matter, gray matter, ‘‘ganglion’’ for neuronal cells) from which the same compounds were first isolated and chemically defined (Thudicum, 1884; Klenk, 1942). The oligosaccharide portion of both neutral and acidic glycosphingolipids may contain also more than ten monosaccharide units: glucose, galactose, N-acetylglucosamine, N-acetylgalactosamine, different species of sialic acid, fucose and glucuronic acid are the most frequently present. In all glycosphingolipids the saccharide unit that is linked to ceramide is glucose (in few cases galactose) and the linkage is b-glucosidic (or b-galactosidic). Glycosphingolipids are commonly classified on the basis of the core oligosaccharide structures present in their molecules, in series:ganglio-series, globo-series, lacto-series, etc. (> Table 6-1). It should be remembered that each of the complex sphingolipids with a homogeneous hydrophilic portion is heterogeneous in both the sphingosine and fatty acid components, thus constituting a mixture of compounds with potentially different physicochemical and biological properties.

2.2.1 Gangliosides The most varied class of glycosphingolipids (in terms of different chemical entities) present in neural tissues is that of gangliosides. Sialic acid, that characterizes the class, is mainly N-acetyl neuraminic acid (5-amino3,5-dideoxy-D-glycero-D-galacto-non-2-ulopyranosonic acid, 5-N-acetylneuraminic acid, Neu 5Ac), followed by 5-N-acetyl,-9-O-acetylneuraminic acid (Neu 5,9 Ac2) and 5-N-glycolylneuraminic acid, Neu5Gc (this form is absent in human cells and tissues, including the nervous system). Sialic acid is a-glycosidically (more precisely a-chetosidically) linked to galactose (a2! 3 linkage), or N-acetylgalactosamine

101

102

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Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

. Figure 6-1 Chemical structure of the most represented simple sphingolipids present in the vertebrate nervous system

(a2! 6 linkage), or to another residue of sialic acid (a1! 8 linkage) (Yu and Ledeen, 1972; Sandhoff and Christomanou, 1979; Svennerholm, 1980; Hakomori, 1981; Ando, 1983; Wiegandt, 1985; Ledeen, 1989; Yu and Saito, 1989; Suzuki and Yamakawa, 1991). Gangliosides were also described in brain carrying an ester linkage between the carboxylic group of a terminal sialic acid and the C-9 hydroxyl group of a contiguous sialic acid residue (Riboni et al., 1986) or C-4 of galactose (Nores et al., 1987) (ganglioside-lactones). Conventionally,

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6

. Table 6-1 Core oligosaccharide structures of glycosphingolipids: some examples. To these structures can be attached residues of sialic acid, fucose and poly-LacNAc[bGal(1–4)bGlcNAc] units in a linear or branched order Series Ganglio

Code name

Abbreviated structure

Lact-Cer Gg3-Cer Gg4-Cer Gg5-Cer

bGal(1–4)bGlc(1–1)Cer bGalNAc(1–4)bGal(1–4)bGlc(1–1)Cer bGal(1–3)bGalNAc(1–4)bGal(1–4)bGlc(1–1)Cer bGalNAc(1–4)bGal(1–3)bGalNAc(1–4)bGal(1–4)bGlc(1–1)Cer

Gb3-Cer Gb4-Cer Gb5-Cer

aGal(1–4)bGal(1–4)bGlc(1–1)Cer bGalNAc(1–3)aGal(1–4)bGal(1–4)bGlc(1–1)Cer bGal(1–3)bGalNAc(1–3)aGal(1–4)bGal(1–4)bGlc(1–1)Cer

Globo

Isoglobo iGb3-Cer iGb4-Cer Lacto, type 1 (para-globo) Lc4 Lacto, type 2 (neo-lacto) nLc4-Cer nLc6-Cer nLc8-Cer

aGal(1–3)bGal(1–4)bGlc(1–1)Cer bGalNAc(1–4)aGal(1–3)bGal(1–4)bGlc(1–1)Cer bGal(1–3)bGlcNAc(1–3)bGal(1–4)bGlc(1–1)Cer bGal(1–4)bGlcNAc(1–3)bGal(1–4)bGlc(1–1)Cer [bGal(1–4)bGlcNAc(1–3)]2bGal(1–4)bGlc(1–1)Cer [bGal(1–4)bGlcNAc(1–3)]3bGal(1–4)bGlc(1–1)Cer

Nomenclature is according to Svennerholm (1980), the IUPAC-IUB recommendations (1998), and some more recent indications (Fahy et al., 2005)

gangliosides are divided into groups depending on the neutral saccharide core to which sialic acid residues are linked. The most common saccharide cores present in neural gangliosides are: lactose [bGal(1!4)Glc], gangliotriose[bGalNAc(1!4)bGal(1!4)Glc] and ganglio-tetraose [bGal(1!3)bGalNAc(1!4)bGal (1!4)Glc] (gangliosides of the ganglio-series). Less common are the ganglio-pentaoses GalNAc-ganglio-tetraose [bGalNAc(1!4) bGal (1!3)bGalNAc(1!4)bGal(1!4)Glc] (Svennerholm et al., 1973), Gal-ganglio-tetraose [aGal(1!3) bGal(1!3)bGalNA(1!4)bGal(1!4)Glc] (Ariga and Yu, 1998) and lacto-neotetraose bGal (1!3)bGlcNAc(1!3)bGal(1!4)Glc (Molin et al., 1987) or lacto-tetraose bGal(1!4) bGlcNAc(1!3)bGal (1!4)bGlcNAc (Molin et al., 1987) (gangliosides of the lacto- and neolacto-series). Fucose, when present, is linked by a1!2 glycosidic linkage to a galactose moiety (Ariga et al., 1987a; Ghidoni et al., 1976). Depending on the position where the sialic acid residue(s) is (are) linked to the neutral core and the (specific) sialyltransferase (s) catalyzing the sialylation steps a further distinction in ‘‘series’’ has been adopted: O-, a-, b-, c- and a(or Chol1)-series (Ando et al., 1992; Kolter et al., 2002). The presence of sulfated gangliosides has been also reported (Galustian et al., 1997). All gangliosides with one exception (ganglioside GM4) are linked to ceramide through the glucose moiety by a b1!1 glucosidic linkage. The schematic formulas of the nervous tissue gangliosides of the different series are presented in > Table 6-2, where gangliosides of some neurotumoral cell lines that undergo a neural-type differentiation in culture under proper conditions are also included. Peculiar is ganglioside GM4 (present in the brain white matter) where sialic acid is linked to the galactose moiety (a2 !3 chetosidic linkage) of galactosylceramide (Yu and Iqbal, 1979). The complete chemical structure of some gangliosides occurring in the vertebrate nervous system, chosen in order to show the basic chemical features [the different length of the neutral oligosaccharide core; the different sialosyl linkages, including lactonization; the presence of fucose, O-acetylated sialic acid, N-glycolyl-neuraminic acid (Tettamanti et al., 1965) and sulfate], is presented in > Figure 6-2. In this figure the nomenclature follows the indications of Svennerholm (1980); and the recommendations of the IUPAC-IUB Joint Commission on Biochemical Nomenclature (1998).

103

Kuhn and Wiegandt (1963)Ledeen and Salsman (1965) Kuhn and Wiegandt (1963)

Ghidoni et al. (1976)

Ariga et al. (1987a)

Kuhn and Wiegandt (1963)

Ghidoni et al. (1976)

Ando and Yu (1977)

GM1

Fuc-GM1

Fuc-Gal-GM1

GD1a

GD1a(Neu5Gc)

GT1a

Ledeen et al. (1973)

Yu and Iqbal (1979) Yu and Ando (1980) Yu and Ando (1980)

GM2

Abbreviated structure

6

Code name o-series GM4 GM1b GD1c a-series GM3

. Table 6-2 Schematic formulas of the gangliosides occurring in neural cells and tissues, and in commonly used neurotumoral cell lines. The code names are according to the suggestions of Svennerholm (1980) and the IUPAC-IUB recommendations (1998)

104 Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

Kuhn and Wiegandt (1963)

Ariga et al. (1995)

Sonnino et al. (1978)

GD1b

9-O-Ac-GD1b

Fuc-GD1b

Ariga et al. (1987a)

Svennerholm et al. (1973)

Kuhn and Wiegandt (1963)

Fuc-Gal-GD1b

GalNAc-GD1b

GT1b

Riboni et al. (1986)

Kuhn and Wiegandt (1964)

GD2

Lactone-GD1b

Kuhn and Wiegandt (1964) Ariga et al. (1995)

b-series GD3 9-O-Ac-GD3

continued

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6 105

Ghidoni et al. (1980)

Kuhn and Wiegandt (1963)

Chigorno et al. (1982b)

Yu and Ando (1980)

Waki et al. (1993b)

Yu and Ando (1980)

Waki et al. (1993a)

Ishizuka and Wiegandt (1972)

GQ1b

9-O-Ac-GQ1b

c-series GT3

9-O-Ac-GT3

GT2

9-O-Ac-GT2

GT1c

Abbreviated structure

Code name 9-O-Ac-GT1b

. Table 6-2 (continued)

106

6 Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

Hirabayashi et al. (1990)

Ando et al. (1992)

Ando et al. (1992)

Molin et al. (1987)

Molin et al. (1987)

GD1a

GT1a

GQ1a

Others 3’isoLM1

6’LM1

continued

Ro¨sner et al. (1985)

GP1c

a-series

Ishizuka and Wiegandt (1972)

GQ1c

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6 107

Chou et al. (1996)

Fukushima et al. (1984)

Galustian et al. (1997)

Galustian et al. (1997)

9-O-Ac-3’LD1

SLex-gangliosidea

O3S-6’-SLexgangliosidea

O3S-6-SLexgangliosidea

a

SLex: sialylLewisx This compound was found to occur in endothelial cells, including brain microvessels b The presence of this compound was observed in PCl2 pheochromucytoma cells (neurotumoral cells)

 O3S-6’,6-bis-a SLexgangliosideb



Galustian et al. (1997)

Chou et al. (1996)

3’LD1

6



Chou et al. (1996)

Abbreviated structure

3’LM1

Code name

. Table 6-2 (continued)

108 Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids . Figure 6-2 Complete chemical structure of some gangliosides occurring in the vertebrate nervous system

6

109

110

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Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

2.2.2 Sulfatides and Other Sulfo-Glycosphingolipids The most abundant sulfate containing glycolipids in the nervous system (particularly in myelin) are sulfated galactosyl-ceramide [SO3-3bGal(1!1) Cer], followed by sulfated lactosyl-ceramide [SO3-3bGal(1!4)b Glc(1!1)Cer], both better known as sulfatides (Vos et al., 1994). In sulfatides sulfate esterification is mostly at C3 (rarely at C6) of the galactose moiety (Vos et al., 1994). Less abundant sulfo-glycosphingolipids characteristically contain sulfated glucuronic acid (GlcA) in C3 (Ariga et al., 1987b, 1990; Needham and Schnaar, 1993; Jungalwala, 1994; Vukelic et al., 2005). These are SO3-3bGlcA (1!3)bGal(1!4)bGlcNAc (1!3)bGal (1!4)bGlc(1!1)Cer (SGGL-1) and SO3-3bGlcA(1!3)bGal(1!4)bGlcNAc(1!3)bGal (1!4bGlcNAc(1!3)bGal (1!4)bGlc(1!1)Cer (SGGL-2), where the neutral oligosaccharide backbone is of the ‘‘lacto’’ type. As already mentioned, some gangliosides contain sulfate groups (see > Table 6-2).

2.2.3 Neutral Glycosphingolipids The most abundant neutral glycosphingolipids in neural cells and tissues are the already cited cerebrosides, Galb1!1,1Cer and Glcb1!1,1Cer, followed by lactosylceramide [bGal(1! 4) bGlc(1!1)Cer] (Barranger and Ginns, 1989). Less common neutral glycosphingolipids are the neutral backbone oligosaccharides present in gangliosides: ganglio-triose, ganglio-tetraose, Gal-ganglio-tetraose, GalNAc-ganglio-tetraose, and oligosaccharides of the lacto-type [bGlcNAc(1!3) bGal(1!4) bGlc(1!1)Cer] and the globo type [aGal(1!3)aGal(1!4)bGal(1!4)bGlc(1!1)Cer; aGal(1!3)aGal(1!3)aGal(1!4)bGal(1!4)bGlc (1!1)Cer; aGal(1!3)aGal(1!3)aGal(1! 3) aGal(1!4) bGal(1!4) bGlc(1!1)Cer; aFuc(1!2)aGal (1!3) aGal(1!4)bGal(1! 4) bGlc(1!1)Cer; bGalNAc(1!3) aGal(1!3) aGal(1!4)bGal(1!4)bGlc (1!1)Cer] (Vanier et al., 1980; Schwarting et al., 1986; Ariga et al., 1988; Chou et al., 1989; Pal et al., 1996; Ariga and Yu, 1998). The schematic formulas of the most common neutral glycosphingolipids present in neural tissues are given in > Table 6-3.

. Table 6-3 Schematic formulas of neutral glycosphingolipids present in neural cells and tissues and in commonly used neurotumoral cell lines. Code names according to the IUPAC-IUB recommendations (1998) Code name Glc-Cer (Gluco-cerebroside) Gal-Cer (Galacto-cerebroside) Lact-Cer GA2 (asialo-GM2) GA1 (asialo-GM1) Asialo Fuc-GM1 Gb3Cer Gal-Gb3-Cer Gal2-Gb3-Cer Gal3-Gb3-Cer Fuc,Gal-Gb3 GalNAc, Gal-Gb3-Cer Asialo 6’-LM1 Lexantigen (SSEA-1) X-antigen (Lex) (Fuca-nLc4 and Cer)

Abbreviated structure bGlc(1–1)-Ceramide bGal(1–1)-Ceramide bGal (1–4)bGlc(1–1)Ceramide bGalNAc(1–4)bGal (1–4)bGlc (1–1)Ceramide bGal(1–3)bGalNAc(1–4)bGal (1–4)bGlc (1–1)Ceramide aFuc(1–2)bGal (1–3)bGalNAc(1–4)bGal (1–4)bGlc (1–1)Ceramide aGal(1–4)bGal(1–4)bGlc(1–1)Ceramide aGal(1–3)aGal(1–4)bGal(1–4)bGlc (1–1)Ceramide aGal(1–3)aGal(1–3)aGal(1–4)bGal(1–4)bGlc (1–1)Ceramide aGal(1–3)aGal(1–3)aGal(1–3)aGal(1–4)bGal(1–4)bGlc (1–1) Ceramide aFuc(1–2)aGal(1–3)aGal(1–4)bGal(1–4)bGlc (1–1)Ceramide bGalNAc(1–3)aGal(1–3)aGal(1–4)bGal(1–4)bGlc (1–1)Ceramide bGal(1–4)bGlcNAc(1–3)bGal(1–4)bGlc (1–1)Ceramide bGal[aFuc(1–3)](1–4)bGlcNAc(1–3)bGal[aFuc(1–3)]bGlcNAc(1–3)-bGal(1–4)bGlc (1–1)Ceramide bGal[aFuc(1–3)]bGlcNAc(1–3)bGal(1–4)bGlc (1–1)Ceramide

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2.2.4 Cationic Glycosphingolipids Cationic sphingolipids that are extremely minor brain components contain an ionized free amino group. Chemically defined examples of these compounds are: (1) glucosyl-sphingosine, galactosyl-sphingosine, lactosyl-sphingosine and sphingosylphosphocholine, already mentioned; (2) galactosyl-sphingosine, where the hydroxyl groups in C3, C4 or C4, C6 of the galactose moiety are engaged in 3,4- or 4,6-cyclic-acetallinkage with palmitoylaldehyde (Hikita et al., 2002); (3) glyceroplasmalo-psychosine where the galactose hydroxyl group in C6 is linked, together with a primary alcoholic group of glycerol, to oleylaldehyde (Hikita et al., 2002); and (4) the de-N-acetyl-Lc3-Cer (where GlcNAc is deacylated) and de-N-acetyl-GM3 (Hanai et al., 1988; Manzi et al., 1990; Hikita et al., 2002) (where Neu5Ac is de-N-acetylated). The formation of lysoderivatives of more complex glycosphingolipids by the action of particular deacylases (endoceramidases) (Ito et al., 1995) cannot be excluded. However, the occurrence of this process under physiological conditions seems to be at least rare.

2.3 Synthetic Sphingolipids, Derivatives of Sphingolipids and NeoSphingolipids The large majority of natural sphingolipids occur in cells, tissues and body fluids in small amounts. Therefore, only very few of them (sphingomyelin, ceramide, some gangliosides) can be prepared from natural sources in amounts satisfying the needs of research plans. On the other hand, the increasing evidence of the exceptionally wide variety of natural sphingolipids, particularly glycosphingolipids, in both their chemical composition and physio-pathological implications, gave rise to a large interest for investigations in the field. This contributed to the urgency to develop methods for the chemical synthesis of sphingolipids starting from easily accessible compounds, and for preparing derivatives or mimics of sphingolipids, as fundamental research tools. A further and often fundamental advantage of synthetic approaches is to produce sphingolipids with a homogeneous hydrophobic portion in both the long chain base and fatty acid components.

2.3.1 Chemical Synthesis of Sphingolipids Because of the wide heterogeneity of their chemical composition, sphingolipids, particularly glycosphingolipids, constituted a real challenge to synthetic organic chemists. Since the mid-1950s, the introduction of new analytical and synthetic methods, and the availability of novel physicochemical methods for structural explorations, led to the development of procedures capable to synthesize virtually any sugar derivative from readily accessible saccharides, like D-glucose. Also sialic acid was chemically synthesized starting from N-acetyl-D-glucosamine (Baumberger and Vasella, 1986; Csuk et al., 1988). The same applies to sphingosine and ceramide (and sphingomyelin), the former being obtained from simple monosaccharides like galactose (Schmidt and Zimmermann, 1986; Curfman and Liotta, 2000), and the latter ones from sphingosine and a fatty acid (and phosphocholine) (Byun et al., 1994; Bushnev and Liotta, 2000). For glycosphingolipid synthesis the problems were to obtain regio-selective and stereo-selective linkages, among the saccharide units, and, in the case of gangliosides, to warrant a-glycosidic linkage of sialic acid to other sugar residues. Different strategies were devised (also in the perspective of large scale preparations) in order to have the glycosyl-donor properly activated at the anomeric C atom, the remainder hydroxyl groups protected and the glycosyl acceptor possessing all the hydroxyl groups protected besides the one to be engaged in the O-glycosidic bond. When dealing with more complex oligosaccharide structures, the generally adopted procedure was to synthesize first the entire oligosaccharide chain and then to link it to either the sphingosine (producing the lyso-glycosphingolipid) or the ceramide moiety. Lyso-glycosphingolipids were then N-acylated at the level of the sphingoid base. The oligosaccharide chain was built up starting from the unit to the linked to ceramide, having its anomeric C properly protected. In a few cases a small

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oligosaccharide (for example a trisaccharide) was directly linked to another small oligosaccharide (for example a disaccharide) before linking to ceramide (Paulsen and Bu˝nsch, 1980). Details on the evolution of the procedures used to synthesize sphingolipids with particular emphasis to glycosphingolipids are reported in valuable reviews (Shapiro, 1970; Gigg, 1980; Paulsen, 1982; Schmidt, 1986, 1989). Notably, attempts were also made to accomplish the synthesis of the glycosidic bonds present in glycosphyngolipids by the use of proper enzymes (glycosyltransferases, transglycosidases, multi-enzyme systems) (Koeller and Wong, 2000). The first synthesis of a glycosphingolipid was that of galactosyl-ceramide containing dihydrosphingosine (sphinganine) (Shapiro and Flowers, 1959), followed by that of glucosyl-ceramide (Shapiro and Flowers, 1961), and of 3’-sulpho-galactosyl-ceramide (Taketomi and Yamakawa, 1964; Flowers, 1966; Jatzkewitz and Nowoczek, 1967). Since then, tens of different glycosphingolipids were synthesized and relying on the presently available procedures, it is expected that any new glycosphingolipid detected in nature can be obtained by chemical synthesis in amounts fulfilling the research needs. Generally, chemically synthesized sphingolipids carry C18 sphingosine (or sphinganine) and stearic or palmitic acid. Of course, any other form of long chain bases and fatty acids can be introduced. A reference list of glycosphingolipids that occur in neural tissues and were obtained via chemical synthesis is given in > Table 6-4.

. Table 6-4 A list of neural glycosphingolipids that were prepared by chemical synthesis, indicating the most characteristic chemical features of the different compounds. The schematic formulas of the corresponding natural compounds are given in > Tables 6-1 and > 6-2. The code names are the same as those of the corresponding natural compounds also given in > Tables 6-1 and > 6-2 Code name Gangliosides GM4 GM3 GM2 GM1 GM1b GalNAc-GM1b -SO3-3-GM1b GD3 GT1a GQ1a SLex ganglioside Sulfo-glycosphingolipids 3-Sulfated Gal-ceramide (3’-sulfatide) 6- Sulfated Gal-ceramide (6’-sulfatide) Sulfated lactosyl-ceramide Neutral glyco-sphingolipids Gal-Cer (Galacto-cerebroside) Glc-Cer (Gluco-cerebroside) Lact-Cer (asialo-GM3) Asialo-GM2 Asialo-GM1 Gb3Cer Asialo 6’-LM1 Lex -antigen (Fuc2a- nLC8)Cer

References Murase et al. (1989a), Numata et al. (1987) Murase et al. (1989b), Numata et al. (1990) Shapiro et al. (1973), Sugimoto et al. (1986) Sugimoto et al. (1986) Prabhanjan et al. (1991) Sugimoto et al. (1990) Komori et al. (2002) Ito et al. (1989), Castro-Palombino et al. (2001) Ito et al. (1997) Hotta et al. (1995) Kameyama et al. (1991), Hasegawa and Kiso (1994) Flowers (1966), Jatzkewitz and Novoczek (1967) Flowers (1966), Jatzkewitz and Novoczek (1967) Schram et al. (1963) Shapiro and Flowers (1959) Koike et al. (1987a) Rapport and Graf (1964) Shapiro et al. (1973), Sugimoto et al. (1985) Sugimoto et al. (1985) Nicolau et al. (1988) Bommer and Schmidt (1989), Koike et al. (1987b) Sato et al. (1987, 1988)

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2.3.2 Chemical Synthesis of Sphingolipid Derivatives Useful for Biological Investigation The inspections at the cellular and subcellular locations of sphingolipids, the routes of their metabolism and intracellular traffic, and the molecular aspects of their functional implications, stimulated the design of procedures capable to obtain derivatives of sphingolipids suitable to approach the different problems. The rationale of these efforts was to introduce into the sphingolipid molecule a proper probe at the level of either the hydrophilic head or the hydrophobic tail. The availability of properly labeled sphingolipids to be used as substrates for the enzymes (extractive or recombinant) involved in their metabolism was also very useful. Notably, the accessibility of pure synthetic sphingolipids greatly facilitated the task of producing ‘‘ad hoc’’ derivatives. The wealth of strategies and successful results in this field is witnessed by the publication of volumes of the ‘‘Methods in Enzymology’’ with review articles dedicated to this topic (Merrill and Hannun, 2000a, b; Lee and Lee, 2003a, b). Radiolabeled sphingolipids. The first approach aimed at preparing radioactive sphingolipids were to inject animals (rats, rabbits) with [3H]- or [14C]-containing metabolic precursors like serine (Kanfer and Bates, 1970) (precursor of the long chain bases) or glucose (Suzuki and Korey, 1964; Tettamanti et al., 1974) and glucosamine (Maccioni et al., 1971) (precursors of the saccharidic components) and then extract, separate and purify the individual radioactive sphingolipids from the proper organs or tissues (mostly brain). Apart from the valuable information regarding the metabolic steps underwent by the precursor, the specific radioactivity of the isolated compounds was too small (order of magnitude 10–25 mCi/mM using [14C]) for a significant use in in vitro experiments. Moreover, the cost would be enormous. Therefore, chemical and enzyme-assisted procedures were developed to obtain a sufficiently high specific radioactivity under reasonable costs. Incorporation of [3H] into simple sphingolipids (sphingosine, ceramide, and their phosphorylated or methylated derivatives) can be achieved by catalytic reduction with tritium (tritium gas or NaB3H4) at the level of the 4,5-double bond of the sphingosine moiety (Gatt, 1966; Schwarzmann, 1978), the label being at C4 and C5. A second procedure is on the basis of a sequential oxidation-reduction reaction where the primary alcoholic group in C1 of sphingosine is first oxidized by 2,3-dicloro-5,6dicyanobenzoquinone (DDQ) with formation of an aldeyde that is then reduced back to the alcoholic function by NaB3H4, the label being at C1 (Toyokuni et al., 1991). A similar procedure consists in the oxidation of the secondary hydroxyl group on C3 with formation of a 3-cheto group that is reduced to the original alcoholic group by NaB3H4 (Gaver and Sweeley, 1966; Radin, 1974; Ghidoni et al., 1981; Bielawska et al., 2000). This reaction gives rise to the threo- and erythro-stereoisomers, necessitating their subsequent separation by HPLC unless special protection of the 3-cheto-sphingoid base is accomplished (Kostinen and Kostinen, 1993). It should be reminded that the tritiation at the 4,5-double bond produces sphinganine that, in terms of signaling potential, has a quite different behavior than that of sphingosine. In the case of ceramide an alternative radiolabeling procedure is on the basis of the acylation of the long chain base with a [3H] or [14C] labeled fatty acid (Kopaczyk and Radin, 1965; Bielawska et al., 1996). Radiolabeling of sphingosine-1-phosphate and sphinganine-1-phosphate is on the basis of two different approaches, both enzyme-assisted. In the first one, sphingosine-kinase is used to phosphorylate sphingosine, in the presence of ATP. Depending on the use of radioactive sphingosine or ATP (on the terminal phosphate group), the resulting sphingosine-1-P can be [3H], [14C] or [32P] labeled (Preiss et al., 1986; Van Veldhoven et al., 1995). This procedure can also be adopted to prepare radiolabeled ceramide-1-phosphate, as the kinase recognizes ceramide as well. According to the second procedure, [3H] labeled dihydro-lysosphingomyelin is treated with a bacterial phospholipase D, with the formation of labeled sphinganine-1phosphate (Van Veldhoven and Mannaerts, 1991). Sphingomyelin can be radiolabeled (Bielawska et al., 2000) on: (1) C-3 of the sphingosine moiety by the oxidation/reduction of the secondary hydroxyl group of the long chain base, as mentioned above; (2) the fatty acyl moiety, starting from sphingosyl-phosphorylcholine and N-acylation with a radiolabeled fatty acid; and (3) the choline moiety by quaternization of ceramide-1-phosphoryl-N,N-dimethylethanolamine(dimethylated sphingomyelin) with [14C] or [3H] methyliodide (Stoffel et al., 1971). Radiolabeling of complex glycosphingolipids, particularly gangliosides, can be obtained at the level of the ceramide tail or the oligosaccharide head, or at both portions (Sonnino et al., 2000). A [3H] radioactive label on C-1 or C-3 of the sphingosine moiety can be introduced by the DDQ/NaB 3H4 oxidation/reduction

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procedures reported above. With reference to the DDQ oxidation of the hydroxyl group of C-3 of the long base chain the required high specificity and efficiency of the reaction can be achieved by conducting the reaction in toluene in the presence of Triton X-100 yielding reversed mixed micelles with the hydrophobic tail oriented on the micelle surface (Ghidoni et al., 1981). The obtained erythro- and threo-stereoisomers are conveniently purified and separated by HPLC (Sonnino et al., 1984a). By this procedure [3H] labeled gangliosides GM3, GM2, GM1, Fuc-GM1, GD1a and asialo-GM1 were prepared but not disialosyl-chain containing gangliosides, like GD1b, GT1b and GQ1b, probably because of the strong repulsive forces between the highly charged oligosaccharide chains that prevent the formation of mixed micelles. After selective removal of the fatty acid residue in KOH-deoxygenated anhydrous propanol (Sonnino et al., 1992), and re-N-acylation with [3H] or [14C] stearic acid (or another fatty acid) ganglioside GM3, GM2 and GM1 labeled in the fatty acid moiety can be obtained. Two different procedures have been developed to introduce radioactivity into the saccharide portion of glycosphingolipids, particularly gangliosides. The first procedure is on the basis of the oxidation of the primary alcoholic group of galactose (or N-acetylgalactosamine) terminally located in the glycolipid molecule, by the action of galactose-oxidase. The produced aldehyde group is then reduced back to alcohol by treatment with NaB3H4. The process, poorly efficient on pure substrates (Hajra et al., 1966) becomes quite satisfactory when mixed micelles of glycosphingolipid and Triton X-100 are used (Ghidoni et al., 1977; Leskava et al., 1984). This procedure was successfully applied to prepare GM1, GD1b and asialo-GM1 radioactive on the terminal Gal, and GM2 on the terminal GalNAc. The second procedure, applied to gangliosides GM3, GM2 and GM1, is on the basis of the finding (Chigorno et al., 1985) that alkaline hydrolysis in butanol-aqueous tetramethyl-ammonium hydroxide yields a mixture of the derivative lacking the acetyl group of sialic acid and the derivative lacking both the acetyl group of sialic acid and the fatty acid chain. The derivative with de-acetylated sialic acid, separated by silica gel-100 column chromatography, is then re-acetylated with [3H] acetic anhydride, with formation of [3H] sialic acid containing gangliosides. Of course, this procedure provides the opportunity to prepare doubly-labelled gangliosides (Sonnino et al., 2000b), with first re-acylation with a [3H]- or [14C]- fatty acid under selective conditions and then acetylation with [3H] acetic anhydride. This procedure was applied to GM3, GM2 and GM1, and provided a proper approach to particular metabolic studies. The specific radioactivity obtained in gangliosides is higher with [3H], ranging from 1 to 5 Ci/mMol using the galactose-oxidase/DDQ oxidation/reduction and the Neu5Ac de-acylation/re-acylation methods, and lower with [14C] using the N-acylation of the lyso-form (0.05–0.06 Ci/mM). A further radio-labeling procedure potentially applicable to all sphingolipids consists of: (1) the selective de-N-acylation of the compound using sphingolipid ceramide-N-deacylase (SCDase) (Ito et al., 1995) under conditions promoting catalytic hydrolysis (acidic pH 5.0–6.0; 0.8% Triton X-100), followed by (2) acylation of the lyso-derivative in the presence of a [3H] or [14C]-labelled fatty acid (Ito et al., 2000a, b). This procedure proved to be efficient for the preparation of the lyso-derivative of Glc-Cer, O3S-Gal-Cer, Lac-cer, sphingomyelin, Gb4-Cer, Gb5-Cer, a number of gangliosides (from GM3 to GQ1b) (Ito et al., 2000a) and already successfully employed to prepare [14C] labeled Gal-Cer, Gb4-Cer, O3S-Gal-Cer, and GM1a. Notably, using the same enzyme, under conditions that favor the reverse reaction (N-acylation) (neutral/basic pH, 7.0–8.0; 0.1% Triton X-100), in the presence of the corresponding lyso-sphingolipid (possibly produced by chemical synthesis) and a radiolabeled fatty acid, radiolabeled sphingolipids can be obtained. Sphingolipids labeled with fluorescent, paramagnetic, photoreactive and other probes. The exploration of the subcellular location, membrane topology, intracellular traffic of sphingolipids, and of their interactions with other cell components posed the need to make sphingolipids recognizable by insertion of a proper probe into their molecule. Ideally, the probe should (1) be chemically inert (with the exception of photoreactive probes to be inert only in the dark); (2) not modify the overall physicochemical properties of the sphingolipids; (3) not interfere with the recognition properties of the sphingolipid; (4) undergo the same metabolic pathways run by the corresponding natural compound and (5) be easily introduced into the sphingolipid molecule. Of course, some ‘‘tolerance’’ in the adoption of these criteria had to be cautiously allowed. A very common scheme used to label sphingolipids (from ceramide to the most complex glycosphingolipids) with these probes is to prepare the corresponding lyso-derivative, by alkaline treatment

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(Sonnino et al., 1992), hydrazinolysis (Suzuki et al., 1984) (in the case of the simpler sphingolipids) or sphingolipid ceramide-N-deacylase hydrolysis (Ito et al., 1995), and then N-acylate them using a fatty acid carrying the probe generally at the CH3-end of the chain. The length of the chain depends on the dimension of the probe, having in mind the criterion to maintain the derivatized sphingolipid spacially close to the natural compound. Of course, all these derivatized sphingolipids carry the probe in their hydrophobic tail. With fluorescent sphingolipids, problems of intracellular location and traffic could be explored by fluorescence microscopy, and very sensitive methods be set up for the assay of many sphingolipid hydrolases (Gatt et al., 1981; Dinur et al., 1984). Anthracene (7-nitrobenz-2-oxa-1,3-diazo-4-yl), pyrene, carbazol (7-nitrobenz-2-oxa-1,3-diazo-4-yl, NBD), dansyl, lissamine rhodamine (LR), BODIPY (Boron dipyrromethene difluoride, BOD) linked via a 12-carbon spacer, or dimethyl BODIPY (DMB), linked via a 5-carbon spacer (Pagano and Sleight, 1985; Acquotti et al., 1986; Dagan et al., 2000), were employed as the fluorescent probes (Pagano and Sleight, 1985; Acquotti et al., 1986; Dagan et al., 2000). In particular the following sphingolipids, labelled with one of these probes, are commercially available: LR-12-ceramide, BOD-12-ceramide, DMB-5-ceramide, LR-12-Glc-Cer, BOD-12-Glc-Cer, DMB-5-Glc-Cer; LR-12-Gal-Cer, BOD-12-Gal-Cer, DMB-5-Gal-Cer, LR-12-SM, BOD-12-SM, DMB-5-SM; LR-12- O3S-Gal-Cer, BOD12-O3S-Gal-Cer, DMB-5-O3S-Gal-Cer, BOD-12-Lac-Cer, DMB-5-Lac-Cer, BOD-12-GalNAc-Gal-GlcCer, BOD-12-GM1, DMB-5-GM1, BOD-12-GM2, where the number -5 or12- indicates the length of the chain carrying the fluorescent probe. Very recently, brilliant red tetramethylrhodamine (TMR) was successfully used as a fluorescent probe for gangliosides GM1, GM2, GM3, asialo-GM1, asialo-GM2, lactosylceramide, glucosylceramide and ceramide (Larsson et al., 2007). By these derivatives the sensitivity of detection lies in the zepto mol (10–21 mol) range when assayed in capillary electrophoresis using laserinduced fluorescence (LIF) (Zhao et al., 1994). This extremely high sensitivity would allow to analyzing sphingolipid metabolites at the single cell level. Fluorescent sphingolipids are taken up by cells in culture, internalized by endocytosis, and submitted to metabolic processing. The first steps of this process concern the administered sphingolipid, the further ones pertain to its metabolites. Not always it is easy to distinguish the starting labelled sphingolipid from its metabolite(s), leading to possible misinterpretation of the results. In this respect, the parallel use of NBD-6-Cer (that undergoes metabolic processing) and the 1-O-methyl-NBD-6-Cer (that is not metabolized) enabled to show in cultured fibroblasts that not the exogenously administered NBD-6-Cer but its metabolites accumulated in the Golgi apparatus (Pu˝tz and Schwarzmann, 1995). Details on the preparation of the different fluorescent sphingolipids, selection of the fluorescent sphingolipid most proper for the specific applications (cell studies; enzyme assays; intracellular fate; intracellular traffic), operative conditions for in vivo and in vitro experiments, and cautions are given in authoritative reviews (Pagano and Sleight, 1985; Acquotti et al., 1986; Dagan et al., 2000; Pagano et al., 2000). Examples of sphingolipids carrying different fluorescent probes are given in > Figure 6-3. A peculiar approach to fluorescence tagging of gangliosides is on the basis of the strategy developed by Bertozzi and coworkers (Laughlin and Bertozzi, 2007) for in vivo labeling of sialoglycoconjugates. According to this strategy cells are fed with peracetylated N-a-azido acetylmannosamine, which, after intracellular deacetylation by esterases, is metabolized to N-a-azido-sialic acid and ultimately to azido-sialoglycoconjugates including azido-gangliosides Azido-gangliosides are then biotinylated with biotin-phosphine and labelled with sterptavidine-FITC (Bussink et al., 2007). This procedure can be used for both recognizing gangliosides at the cell level by fluorescence flow cytometry and establishing the pattern of the individual species after extraction and fractionation. Paramagnetic probes were introduced into the molecules of some gangliosides at the level of their oligosaccharide head (Lee et al., 1980) or ceramide tail (Acquotti et al., 1986). In the former case a residue of 4-amino-2,2,4,4-tetramethylpiperidine-1-oxyl was introduced, through the amino group, into the glycerol tail of ganglioside sialic acid. In the latter case GM1 was first submitted to de-N-acylation of the sphingosine moiety and de-N-acetylation of the sialic acid residue according to Sonnino et al. (1992), then re-N-acylated with one of the two doxyl (4,4-dimethyl-3-oxazolinyl oxy) derivatives of stearic acid, 5-doxyl- and 16-doxyl-stearic acids, where the paramagnetic probe was inserted in C6 or C12 of the fatty acid, respectively. The de-N-acetylated doxyl derivatives, after proper purification, were submitted to re-Nacetylation of the sialic acid residue (Sonnino et al., 1989), yielding 5-doxyl- and 12-doxyl-GM1. The two doxyl-carrying GM1 molecules resulted to have the same aggregation properties and ability to be

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incorporated into phosphatidylcholine liposomes and taken up by cells in culture, as well as to undergo intracellular metabolic processing as natural GM1 (Acquotti et al., 1986). The chemical structure of a sphingolipid carrying a paramagnetic probe namely doxyl-GM1, is given in > Figure 6-3. An improvement in the sensitivity and precision of the experimental approach aimed at defining the plasma membrane localization and the mechanism of uptake and endocytosis of gangliosides exogenously administered to cells in culture (dermal fibroblasts, neuroblastoma cells of different origin) was achieved by preparing a derivative of GM1 carrying a biotinyl residue. This residue is linked to the amino group of GM1 sialic acid, through a C6-spacer (amino caproic acid) (Albrecht et al., 1997). As the biotinyl group is linked to a terminal component of the saccharide portion of ganglioside it is suitable to interact specifically with a . Figure 6-3 Examples of the derivative of a glycosphingolipid, ganglioside GM1, carrying: (a) a fluorescent probe; (b) a paramagnetic probe; (c) a biotinyl probe; (d) a phenyldiazinyl probe

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. Figure 6-3 (continued)

goat anti-biotin antibody conjugated to ultra-small gold particles recognizable by electron microscopy (Albrecht et al., 1997). The synthesis of biotinyl-GM1 starts from the de-N-acylated, de-N-acetylated GM1 obtained as reported above. This compound is first submitted to N-acylation of the sphingoid base with a C8 or C18 fatty acid, and then to conjugation of the amino group of sialic acid with a biotinyl amino caproyl residue, yielding biotinyl-C8-GM1 and biotinyl-C18-GM1. The acyl portion of the compounds can be [3H] or [14C] radio-labeled obtaining doubly-labelled GM1 derivatives. Biotinyl-tagged-GM1 can be sequentially hydrolyzed by a b-galactosidase and a b-N-acetylhexosaminidase to biotinyl-GM2 and biotinyl-GM3, respectively (Albrecht et al., 1997). Instead, it is resistant to the action of sialidases, likely because of the steric hindrance caused by the bulky and spacer linked biotin moiety (Albrecht et al., 1997). Therefore, administered and endocytosed biotinyl-GM1, differently from other derivatives of GM1, is expected to undergo only partial degradation. This difference may help to better interpret the data obtained in cultured cells after administration of differently labeled gangliosides, for instance fluorescent and biotinylated gangliosides. The structure of biotinylated-GM1 is given in > Figure 6-3. Details of the preparation and use of biotinyl-GM1 are given in a review (Schwarzmann et al., 2000). The identification of receptors for sphingolipids, specific binding sites of enzymes affecting sphingolipids, proteins (or other cell components) that are partners of sphingolipids in membrane domains and carriers of sphingolipids, may take advantage of the use of photoaffinity labeling of sphingolipids (Bayley and Knowles, 1977; Peters and Richards, 1977; Brunner, 1993). The sphingolipid has to be modified by incorporation of a chemically inert but photo-chemically activable functional group. On irradiation, the photo-activable group is converted into a very reactive species that reacts with residues of the receptor or other sphingophilic compounds that are vicinal to the sphingolipids, with formation of a covalent bond. The concomitant presence of a radiolabel in the sphingolipid moiety allows to recognizing the bound partner and, in the case of the protein, the protein domain or individual amino acids engaged in the bond. The general strategy used to prepare sphingolipids carrying photoreactive probes is to link the probe (with a possible spacer) to the free amino group of the long chain base or sialic acid obtained by proper hydrolysis from the natural (or synthetic) sphingolipid, as previously reported. The general criteria for ideal photolabeling of sphingolipids are reported in the review by Knoll et al. (2000).

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Most of the photoaffinity reagents used to label sphingolipids are precursors of nitrenes, (azides, especially aryl azides) and carbenes (diazinines, especially trifluoromethyldiazinines) and can be divided into those that aim at recognizing the interactions involving the lipid portion of the molecule (all sphingolipids) and those regarding the interactions implicating the saccharide portion (glycosphingolipids). Examples of photoaffinity labels on the saccharide portion of glycolipids are the [125I] –nitrene derivative of GD1b (at the level of the terminal sialic acid residue), employed to identify the gangliophilic domain of Tetanus toxin (Shapiro et al., 1997), and the carbene-derivatives of GM3 (not carrying radioactivity) that acts as specific inhibitor of GM2-synthase and GD3-synthase. Examples of photoaffinity labels on the lipid moiety are the following: (1) the radioiodinated 3-trifluoromethyl-3-[4-(2-p-toluenesulfonyloxyethyl)-phenyl] diazinine-ceramide used to recognize the specific binding to and activation of protein kinase c-Raf by ceramide (Weber and Brunner, 1995); (2) the radioiodinated azido-derivatives of sphingomyelin and glucosyl-ceramide (Zegers et al., 1997), employed in an attempt to identify compartment specific proteins involved in sphingolipid sorting and metabolism; (3) the radioiodinated azido derivative of galactosyl-ceramide, used to examine the Gal-Cer-binding protein in human cerebrospinal fluid (Chiba et al., 1994); (4) the biotinylated-[14C] labeled phenyldiarizine derivative of Lac-Cer employed in the study of the glycosyl acceptor region of GM3-synthase (Hatanaka et al., 1995); (5) the derivative of GM1containing a nitrophenyl azide group at the terminus of the fatty acid moiety and a [3H] label at the acetyl group of sialic acid, used to investigate the protein involved in GM1 uptake, internalization and metabolic processing by human fibroblasts in culture (unlabeled and photoreactive GM1 provided the same metabolic pattern) (Sonnino et al., 1989), and the lyso-GM1 derivative containing a carbene precursor (N-diazinyl) linked to the amino group of the long chain base (tritiated at C4–C5) that, upon irradiation, was proved to link to phosphatidylcholine, phosphatidylserine and cholesterol in both liposomes and calf brain microsomes (Meier et al., 1990); and (6) the carbene precursor N-(2-diazo-3,3,3-trifluoropropionyl) derivative of GM2, with the photolabeling probe linked to the amino group of the sphingosine moiety, employed for exploring the glycolipid binding domain of the GM2 activator protein (Knoll et al., 2000). The chemical structure of a sphingolipid carrying a photo-affinity label, in particular phenyldiazinyl-GM1, is given in > Figure 6-3.

2.3.3 Chemical Synthesis of Sphingolipid Analogs, Unnatural Sphingolipids and Neo-Sphingolipids The availability of efficient methods for the chemical synthesis of sphingolipids together with the consolidated notion of their great biological potential stimulated the search of (1) analogs to be used for exploring the metabolic and functional implications of sphingolipids, (2) un-natural sphingolipids to study the molecular mechanisms of sphingolipid–protein interactions and their possible use as therapeutic tools, (3) neo-glycolipids with the oligosaccharide portion anchored to a hydrophobic tail different from ceramide, or to a hydrophylic tail making the compound soluble and (4) mimetics of sphingolipids having the same (or similar) steric conformation but with a different chemical composition. With regard to sphingolipids analogs particular interest was devoted to glycosphingolipids with glycosidic linkages not of the O-type, in order to acquire resistance to enzyme-assisted deglycosylations/ glycosylations, maintaining a conformation close to that of the corresponding natural compounds. Analogs of this kind could be useful in assessing cell behavior of individual (glyco)sphingolipids in the absence of any metabolic processing. The synthesis of C-, S,-N-glycosides in the perspective of applications in the glycosphingolipid field was first described by Nicolau group (Nicolau et al., 1984a, b). Further developments focused on S-glycosides and the following thio-analogs were synthesized: thio-Glc-Cer (Weis et al., 1985), thio-Gal-Cer/thio-Lac-Cer/thio-GM3 (Hasegawa et al., 1991), lyso-thio-Glc-Cer (Schwarzmann, 2000), thio-Glc-Cer carrying a [14C] or fluorescent probe in the fatty acid residue (Schwarzmann, 2000) and thio-GM1 (Schwarzmann et al., 1997), all with the S-glycosidic linkage to ceramide. A thio-Lac-Cer has been also synthesized with the S-glycosyl-linkage between the galactose and glucose moieties (Schwarzmann, 2000), as well as thio-gangliosides containing a-thioglycosidically linked sialic acid (thio-iso-GM4, thio-iso GM3 with the sialic acid linked to C6 of galactose, and the oligosaccharide

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chain S-linked to ceramide, and GD3 with S-linked terminal sialic acid) (Ishida et al., 1994). Examples of sphingolipid analogs are reported in > Figure 6-4. The field of synthetic un-natural sphingolipids is becoming crowded. Examples are given to emphasize the wide range of potential application of these compounds. a-Gal-Cer was proved to be a potent . Figure 6-4 Examples of analogs of glycosphingolipids, neo-glycosphingolipids and mimetics of sphingolipids

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stimulatory agent in an immunological process mediated by CD1 molecules (Plettenburg et al., 2002), opening the way to explore bioactivity of glycosphingolipids with the oligosaccharide chain a-linked to ceramide. Sialyl(a2!1)sphingosine has the property to de-stabilize the structure of glycosphingolipid signaling domains, inhibiting the clustering of GM3 in these domains and blocking GM3-dependent processes of cell adhesion and signaling (Zhang et al., 2000). In the course of studies on the mechanism of binding of Tetanus toxin to its ligand gangliosides GT1b and GD1b (particularly the fragment c of the toxin subunit H, responsible for the binding, TeNTHc), attempts were made to crystallize the complex between GT1b and TeNTHc. The crystallization could not be obtained with GT1b, but was successful with a synthetic GT1b containing a b-linked sialic acid to the inner galactose moiety (instead of the a-linkage of the natural compound) and, in addition, a 2-(trimethylsilyl) ethyl group substituting the entire ceramide portion (Fotinou et al., 2001). In a similar study, where the molecular requirements for the interaction between sialyl Lewis X ganglioside(s) and members of the selectin family were explored, it was observed that the fucose moiety had a pivotal role in the binding to E-and L-selectin: in fact the binding was completely suppressed using derivatives of sialyl Lewis X ganglioside having a deoxy-fucose in either one of positions 2,3 and 4, demonstrating that hydroxylation in all these positions is required for binding (Hasegawa et al., 1994). Concerning the enzymology of gangliosides, particularly in relation with the pathological syndromes deriving from inborn deficiency of some of the enzymes involved in ganglioside degradation, it became early known that ganglioside GM1 and GM2 are resistant to most of sialidases and GM2 requires the presence of an activating protein (a saposin) in order to be affected by hexosaminidase with liberation of N-acetylgalactosamine and GM3. In the course of the investigations regarding the mechanism of hexosaminidase hydrolysis of GM2 in the presence of the activating protein (HexAP), it was observed that the unnatural and synthetic 6-GM2, an isomer of GM2 where the terminal GalNAc is b1-6 linked to Gal (instead of b1-4), was easily hydrolyzed by hexosaminidase without any need of HexAP (Li et al., 2003). The GM2-oligosaccharide could be hydrolyzed by the same enzyme only in the presence of HexAP, whereas 6’-GM2-olisaccharide hydrolysis did not require HexAP (Li et al., 2003). NMR spectroscopy analysis and related conformational studies, carried out parallelly on GM2 and 6’-GM2, showed that in GM2 the trisaccharide bGalNAc(1!4) [aNeu5NAc(2!3)] bGal has a rigid core with a more mobile region corresponding to the external Glc residue, whereas in 6’-GM2 the above trisaccharide core displays a significant flexibility, likely depending on the 3,4-unsubstituted Gal residue, suggesting that the resistance of GM2 to hexosaminidase action is because of the rigid conformation of the oligosaccharide portion preventing the accessibility of GalNac to the enzyme active site (Li et al., 2003, 2008). The interaction with HexAP changes this conformation allowing enzyme anchoring and subsequent hydrolysis. Another example of un-natural glycosphingolipid mimicking the function of a natural one concerns the gangliosides that bind siglecs (sialic acid binding Ig-like lectins), particularly the neural siglec ‘‘myelin associated glycoprotein’’ (MAG), implicated in myelin–axon interactions (Schnaar et al., 1998). It is known that the best ligands for MAG are the a-series gangliosides, particularly GQ1a (Yang et al., 1996), and that the three sialic acid residues linked a-(2–3) to the terminal Gal, a (2–3) to the internal Gal and a (2–6) to GalNAc are responsible for highest binding affinity (Collins et al., 1997). The observation that each hydroxyl group on the exocyclic glycerol chain of the a (2–6)-linked Neu5Ac residue did not influence the binding capacity, led to explore the possibility that this internal Neu5NAc might be replaced by other anionic substituents. To this purpose three derivatives of GM1b (the backbone of GQ1a) were synthesized: one containing a sulfate group on C6 of GalNAc, one having a second sulfate on C3 of the internal Gal, and one being the isomer of the disulphorylated ganglioside with the external Galb (1–4) linked to GalNAc (Ito et al., 2003). All these compounds are highly active ligands for MAG, and disulphorylated iso-GM1b is the most potent MAG binding compound known to date. A promising group of glycosphingolipid mimetics is on the basis of chemical modifications at the level of the sphingosine moiety. It is known that ganglioside GM3 inhibits tyrosine kinase associated to the epidermal growth factor receptor (EGFR) and that Lyso-GM3 has the same, but stronger, effect (Hanai et al., 1988), although associated to cytotoxicity. Cytotoxicity was abolished by using lyso-GM3 dimer, a synthetic compound arising from the conjugation of two lyso-GM3 residues to glutaminyl-glutamine through the 2-aminogroups of sphingosine moieties (Murosuka et al., 2007). Both effects (strong inhibition of EGFR tyrosin kinase and absence of cytotoxicity) were obtained with a mimetic where sphingosine was

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substituted by 2-amino-dodecanol, with a marked simplification of the synthesis protocol (Haga et al., 2008). The chemical structures of some un-natural sphingolipids mimicking the function of natural counterparts are given in > Figure 6-4. Sphingolipids, particularly glycosphingolipids, have very peculiar physicochemical and aggregational properties attributing the ability to specifically interact with ligands through both the oligosaccharide and ceramide portions. The extreme heterogeneity of the oligosaccharide portion is the basis for a wide variety of specific interactions (saccharide–saccharide, saccharide–protein interactions) and diversity in the hydrophobic trails (long chain bases and fatty acids) is also the source of different interactions (hydrophobic interactions) (Hakomori, 1981; Hakomori and Igarashi, 1993). Other important features of sphingolipids are the following: (1) the ability to undergo phase separation on membrane surface participating in the formation of enriched domains (lipid rafts, glycosphingolipid enriched domains, detergent resistant domains), giving rise to additional interactive abilities (Sonnino et al., 2006); and (2) the possibility that the engagement of the oligosaccharide portion in binding induces conformational changes to the ceramide portion and binding through the ceramide portion influences the conformation and binding properties of the oligosaccharide portion (Crook et al., 1986; Stro˝mberg et al., 1991; Lingwood, 1996). Hence, the need of synthetic compounds capable to give information on the specific role separately played by the oligosaccharide and the ceramide portions of glycosphingolipids. In principle, these synthetic compounds are part of the so called neo-glycolipids (Tang et al., 1985), compounds containing an oligosaccharide (synthetically produced or obtained by proper deglycosylation of O- or N-linked glycoproteins) linked to a lipidic anchor, like L-1,2-dipalmitoyl-sm-glycero-phosphoethanolamine (DPPE) and L-1,2-dehexadecyl-sm-glycero-3phospho ethanolamine (DHPE). Before anchoring, the reducing sugar of the oligosaccharide is reduced to alcohol allowing the formed alcoholic hydroxyl to react with the ethanolamine amino group. These neoglycolipids are particularly used in studies on carbohydrate-dependent immunogenicity, receptor function, and lectin and toxin binding (Feizi and Childs, 1994; Feizi et al., 1994). The same rationale was employed to link oligosaccharides (also typical of glycosphingolipids) to phosphatidyl-ethanolamine carrying acyl (or alkyl) groups of different chains (lauroyl, myristoyl, palmitoyl, stearoyl) (Pohlenz et al., 1992). The obtained glycosylphosphatidylethanolamines can also be acetylated on the =NH group (N-acetyl species) (Yang et al., 1996). Neo-glycolipids of this type were obtained containing Glc-ol; bGal(1!4)Glc-ol;aNeu5Ac(1!3) bGal(1!4)Glc-ol;bGal(1!3)bGlcNAc(1!4)bGal(1!4)Glcol;dFuc(1!2)Gal(1!3)bGlcNAc(1!4) bGal(1!4)Glcol;bGal(1!3)bGalNAc(1!4)bGal(1!4)Glcol;bGal(1!3)bGalNAc(1!4)[Neu5NAc2!3)] bGal(1!4); and bGlc-ol; aNeuAc(2!8) aNeuA(2!3) bGal(1!4)Glc-ol, also as the N-acetyl species, and with acyl or alkyl residues of different chain lenght. These neo-glycolipids were successfully used as enzyme substrates, particularly for sialyltransferases. In a comparative study, where GM3 and GM1 and the corresponding neo-glycolipids were employed as substrates of Golgi sialyl transferase(s), it was observed that in both cases the neo-glycolipids were better substrates for the enzymes, the ones having the most positive effect (about 16–18 fold) being the N-acetyl forms carrying a C16 alkyl chain (Pohlenz and Egge, 1994). A particular group of neo-glycolipids are the synthetic soluble mimics of glycosphingolipids designed as tools to investigate aspects of the interactions of glycosphingolipids with various toxins and viruses. One approach was to substitute the fatty acid of glycosphingolipids with a rigid condensed hydrophobic frame (adamantane or norburnane)in order to generate a soluble mimic retaining the general hydrophobic character of the interface domain but presenting a more globular structure to disfavor lateral interactions within a hydrophilic environment (Lingwood and Mylvaganam, 2003). In order to synthesize the adamantyl-mimic, glycosphingolipids are first selectively de-N-acylated by the general procedure already reported. The adamantyl-mimics of Gal-Cer, O3S-Gal-Cer, Gb3-Cer and Gb4-Cer have been produced and used in ELISA and TLC overlay assays as inhibitors of the binding of verotoxin (Mylvaganam and Lingwood, 1999) and HIVgp120 (a glycoprotein of HIV 1 surface envelope) (Mamelak et al., 2001) to glycosphingolipids (particularly Gb3/verotoxin and O3S-Gal-Cer/HIVgp120) (De Rosa et al., 2008). A second approach starts from lactose that is reductaminated to amino lactitol and then submitted to N-acylation with both a fatty acid of different chain length and a carboxylated medium chain (Fantini et al., 1997), with formation of a soluble mimic of Gal-Cer. This mimetic Gal-Cer was able to interact with an anti-Gal-Cer antibody better than Gal-Cer and featured a marked inhibition of the infection of exponentially growing HT-29 cells by HIV-1 virus (Fantini, 2000). Details on the preparation and use of soluble mimics of glycosphingolipids

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are reported in dedicated reviews (Fantini, 2000; Lingwood and Mylvaganam, 2003; De Rosa et al., 2008). The chemical structures of some neo-glycolipids, particularly hydrosoluble neo-glycolipids, are presented in > Figure 6-4. A particular aspect of the soluble forms of sphingolipids concerns short chain ceramides. Naturally occurring ceramide is a strongly hydrophobic molecule. Owing to its peculiar bio-signaling properties a mandatory experimental approach was to induce an increase of intracellular ceramide concentration and record the following cellular responses. To this purpose one strategy was to stimulate the metabolic production of ceramide (for instance by stimulation of endogenous sphingomyelinase or application of exogenous sphingomyelinase) and another one was to supplement cells with a form of ceramide that could be easily taken up. Hence the use of ceramides shortened in their fatty acyl chain (particularly C2 and C6 ceramides) or in both the sphingoid base and the fatty acyl chain (C8–C8 ceramide). These ‘‘soluble’’ ceramides can be prepared starting from either natural C18 and C20 sphingosines, acetylated or acylated with a C6 fatty acid, or synthetic C8 sphingosine acylated with a C8 fatty acid. The reasons to use short chain ceramides, when, and how to use them, as well as the abundance of reliable information from their use and the matters of caution are described in a dedicated review (Luberto and Hannun, 2000). The findings that the permeability and solubility restrictions of the long chain ceramides can be overcome by delivering the molecules in a dodecane/ethanol solution (Ji et al., 1995), and that short chain ceramides, after uptake by cells, undergo metabolic processing with formation of the regular long chain forms from released sphingosine (Ogretmen et al., 2002), have solved most of the possible caveats for the use of short chain ceramides. The accurate definition of the conformational features of the oligosaccharide portion of glycosphingolipids stimulated the search of glycomimetics of potential therapeutical sygnificance. An example of this approach regards the mimics of GM1-oligosaccharide, starting from the well known notion that Cholera toxin binds GM1 (with by far the highest affinity) engaging the sialyl and terminal Gal residues, and that the GM1-oligosaccharide per se interacts as well with the same toxin although with low affinity (Scho˝n and Freire, 1989). The studies on the conformation of various ganglioside head groups agree that all the head groups containing bGal(1!3)bGalNAc(1!4)bGal trisaccharide or its bGalNAc (1!4) bGal fragment feature a single conformation whereas the aNeu5Ac (2!3)bGal fragment can assume two conformations reflecting two different orientations of the Neu5Ac residue. In the cases of GM2 and GM1 where the internal Gal is linked both to GalNAc and Neu5Ac (Gal-3,4 branched) the conformational freedom of the gangliosides around the 3,4 branched Gal is extremely low and the structure rigid, rendering the molecules less accessible to enzymes, like sialidase and hexosaminidase. In other words 3,4 branched Gal acts as a scaffold holding in site the remainder saccharide units of the oligosaccharide. On this basis the central 3,4-disubstituted Gal residue of GM1 was replaced by dicarboxy cis-1,2- cycloexanedial, which possesses the same configuration of galactose and is locked in a single chain conformation (Bernardi et al., 1999). This GM1 mimic shares with GM1 almost the same steric conformation and the ability to bind cholera toxin. Further studies led to mimics where the sialic acid residue is substituted with a hydroxyacid (lactic acid, glycolic acid). The mimic containing lactic acid displays the strongest affinity for cholera toxin (Bernardi et al., 2002). The chemical structure of some glycomimetics of GM1 is given in > Figure 6-4.

3

Compositional and Developmental Profiles of Sphingolipids in the Nervous System of Different Animals

Since the first explorations on the chemical composition of neural tissues, the evidence emerged that sphingolipids, particularly gangliosides, are largely abundant in the gray matter, cerebrosides and sulphatides in the white matter, and sphingomyelin in both brain portions. Hence, the consolidated notion that in the brain gangliosides are chemical markers of neurons, whereas cerebrosides and sulphatides of glial cells or, more precisely, myelin. Peculiarly, the content of gangliosides, as nmoles bound Neu5Ac/g fresh weight, is highest in neural tissues: brain gray matter, 3,000–4,000; brain white matter, 1,000–1,500; spinal cord, 500–700; peripheral nerves, 150–250; retina, 100–150; followed by spleen, 250–350; liver, 150–250; thyroid,

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100–200; kidney, 75–150; skeletal muscle, 50–80; skin, 30–40; adipose tissue, 10–15; small intestine (mucosa, 5–10; muscular layer, 40–80) (Svennerholm, 1984).

3.1 Analytical Approaches for the Detection, Structural Characterization and Quantification of Sphingolipids 3.1.1 Analytical Biochemistry of Sphingolipids The analytical biochemistry of neural sphingolipids is particularly laborious, owing to the extreme variety of their chemical composition and the complex morphological architecture of the nervous system at the regional, cellular and subcellular levels. Fundamental was the development of very sensitive micro-procedures for the detection and chemical characterization of sphingolipids. The methods for extraction, purification, separation and chemical characterization of the individual sphingolipids have been the object of extensive and critical reviews in different volumes of ‘‘Methods in Enzymology’’ (Merrill and Hannun, 2000a, b; Lee and Lee, 2003a, b). Sphingolipids constitute a real ‘‘analytical’’ challenge. Once extracted from biological specimens (for details see: Radin, 1988) the separation into individual entities is most commonly achieved by high performance thin layer chromatography (HPTLC) on silica gel plates, using different solvent systems and mono-dimensional or bi-dimensional runs (Ledeen and Yu, 1982; Yates, 1988; Ladish and Li, 2000; Van Echten-Deckert, 2000; Yu and Ariga, 2000). A bidimentional HPTLC with ammonia treatment between the first and second run enables to recognize alkali-labile ganglioside species (ganglioside lactones and ganglioside containing O-acetylated sialic acid residues) (Sonnino et al., 1983). HPTLC is the simplest available technique provided with a good resolution power (mainly sensitive to differences in the saccharide moieties) and suitable to the analysis of small samples. Detection of the separated substances is accomplished colorimetrically by spraying appropriate reagents [resorcinol-HCl (Svennerholm, 1957) or p-dimethylaminobenzaldehyde (Chigorno et al., 1982a) reagent for gangliosides; sulfuric acid for phospholipids-sphingomyelin (Kundu, 1981); anisaldehyde reagent for all glycosphingolipids including neutral glycosphingolipids (Partridge, 1948)] and heating. Adopting standard conditions an acceptably precise quantification can be obtained by TLC-chromatoscanning. Identification of the individual substances is accomplished by the use of standards of known structure concomitantly submitted to HPLC runs and colorimetrically detected. An alternative procedure for detecting glycosphingolipids is overlay immunostaining where a specific anti-glycosphingolipid antibody binds to the correspondent glycosphingolipid and the complex is then recognized and quantified by an immunometric technique. This innovative procedure was introduced by Magnani et al. (1980) and further developed. The rationale, procedural details and conditions of this approach are exposed in dedicated reviews (Ishikawa and Taki, 2000; Kannagi, 2000). This method, although not devoid of false positive/negative results (Suetake and Yu, 2003), may enable to recognize, in a mixture, the presence of components otherwise undetectable, owing to their very minute amounts, or to incomplete resolution from the major components. It is worth a comment concerning HPTLC procedures employed for quantitative purposes. The sensitivity (referred to the amount of sample submitted to HPTLC) relies on the detection method used and is of the order of 0.1–0.2 nmol with colorimetric methods and 0.1–1.0 pmol with immunometric methods (Svennerholm, 1984). Improvement of sensitivity can be obtained under conditions (for instance cell cultures) where a metabolic radiolabeling of sphingolipids at steady-state is reached with a precursor of very high specific radioactivity (for instance [3H]sphingosine, 2Ci/mmol or 3-[3H]sphingosine, 20Ci/mmol), and radioscanners of extremely high sensitivity (Radiospace b-imager) are employed. Adopting these conditions the sensitivity can be shifted to 1–5 f mol (Prinetti et al., 2000; Riboni et al., 2000). More complex procedures are also available to fractionate sphingolipid mixtures into sub-families of components or individual components. One of these is high performance liquid chromatography (HPLC, both normal and reversed phase-HPLC), where the resolution power works on differences in both the saccharide moiety and the hydrophobic tail of glycosphingolipids. In other words, the same sphingolipids can be separated into molecular species differing only for a different sphingoid base or fatty acid.

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This procedure is particularly useful for analysis of very complex sphingolipid mixtures, that are separated into sub-families (for instance mono-sialogangliosides, di-sialogangliosides, tri-sialogangliosides, etc.), each of them being further sub-fractionated by HPTLC or HPLC. Detailed descriptions of the application of HPLC to sphingolipids analysis are available (Sonnino et al., 1984a, 2000a; Ullman and McCluer, 1985; Mu¨thing, 2000). Recently, the conditions were worked out to separate glycosphingolipids by off-line and on-line capillary electrophoresis (El Rassi, 1999; Zamfir et al., 2000). This technique proved to have a high resolution power (Zamfir et al., 2000), in some cases superior to that of HPLC. The advantage of HPLC as well as capillary electrophoresis, is that each eluted fraction can be submitted to mass-spectrometry analysis and each component structurally characterized. In other words, identification of the individual components is direct, not by comparison with known standards. The elucidation of the primary and secondary structures of sphingolipids was very strictly dependent on the evolution of the conventional methods of analytical and structural chemistry of oligosaccharides [permethylation, periodate oxidation, Smith degradation, partial acid hydrolysis, enzymatic hydrolysis, ozonolysis, optical rotation measurements, etc. (see reference Whistler and Melville, 1976, for details)] and of gas–liquid chromatography (GLC), mass spectrometry (MS) and proton nuclear magnetic resonance (NMR) spectroscopy techniques, capable to provide unequivocal information on sugar, long chain base and fatty acid composition, sugar sequence, anomery of glycosidic linkages and conformation of sphingolipids, using the most minute amounts of samples (see Whistler and Melville, 1976 and Hakomori, 2008, for details). GLC procedures were firstly applied to sphingolipid structural analysis in the late fifties and sixties of the last century, with the estimation of saccharides (Vance and Sweeley, 1967), long chain bases (Sweeley and Moscatelli, 1959) and fatty acids (Klenk and Gielen, 1961; Radin and Akahori, 1961; Rapport et al., 1961). The general strategy, perfected thereafter, consisted of the following steps: (1) acidic methanolysis of the sample (that had the side effect to N-de-acylate N-acetyl-hexosamines and sialic acids); (2) removal of the fatty acid methyl esters by hexane extraction, and their analysis by GLC; (3) neutralization of the residue and re-N-acetylation of hexosamines and sialic acids; (4) transformation of the saccharide residues into O-trimethylsilyl (TMS) derivatives and; (5) GLC analysis on 20% SE-30 or OV-1 columns using a flame ionization detector. Long chain base analysis required preliminary periodate oxidation followed by GLC of the released aldeydes (Sweeley and Moscatelli, 1959). For detection and quantification of N-acetyl- and N-glycolylneuraminic acid by GLC, mild acid methanolysis conditions had to be employed, in order to avoid N-de-acetylation or N-de-glycolylation, respectively, and the sialic acid TMS-derivatives were isothermally separated on 3% OV-1 or 3% OV-225 columns (Yu and Ledeen, 1970). An alternative procedure consisted of preliminary acid methanolysis, followed by N,O-trifluoroacetylation (TFA) (Ando and Yamakawa, 1971). The TFA-derivatives of saccharides are more stable to acid condition and heat and have higher volatility than the TMS-derivatives. With both TMS and TFA derivatization, each sugar may provide more than one peak. A single peak per saccharide unit can be obtained when the alditolacetate derivatives of hexoses and hexosamines are used for GLC analysis. To this purpose the glycosphingolipid sample is subjected to acid hydrolysis and the sugars are reduced to their alditol derivatives, which are subsequently acetylated and then submitted to GLC. This method, although not applicable to sialic acid analysis (Torello et al., 1980), facilitates the determination of molar ratios within saccharides. The sensitivity of this procedure for quantitative purposes is in the range of 0.03–0.05 mg/individual saccharide/single injection, that is 0.1–0.5 mg of starting sphingolipid sample. Details of these procedures, interesting also from the historical point of view, are reported in excellent reviews (Sweeley and Siddiqui, 1977; Yates, 1988). The first applications of MS for structural analysis of sphingolipids were in the form of electron impact mass spectrometry (EIMS) and served to elucidate the structure of different ceramides (Samuelsson and Samuelsson, 1968) and simple neutral glycosphingolipids (Sweeley and Dawson, 1969). This approach required derivatization of the sphingolipids as trimethylsilyl or permethyl esters (in order to increase their volatility) and made use of high energy ionization method (electron impact) that ineluctably causes extensive fragmentations of molecular species, precluding analysis of sphingolipids carrying large saccharide heads. When milder ionization techniques became available, such as fast atom bombardment (FAB) and liquid secondary ionization mass spectrometry (LSI-MS), sample derivatization was no more necessary and, by the use of multiple stages of mass analysis (tandem mass spectrometry, MS-MS), product ions

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6

diagnostic of the carbohydrate head groups, sphingoid bases, or fatty acids could be obtained (Adams and Ann, 1993). However, these methods require the use of a matrix to solubilize and ionize the sample, which limits the sensitivity and makes difficult quantification of the sample. These problems were solved with the introduction of matrix-assisted laser desorption ionization (MALDI) time- of -flight (TOF) (MALDI-TOF) mass spectrometry that markedly facilitates sample ionization. The addition of delayed ion extraction (DE) (DE-MALDI/TOF-MS) enabled to increase the sample mass (>100 kDa), sensitivity ( 100 f mol) and accuracy ( 0.1%). This method was applied to analyze mixtures of glycosphingolipids of the different series, sphingomyelins and ceramides (Taketomi and Sugiyama, 2000). More recently, electron spray ionization (ESI) was introduced, providing a soft ionization of the analyte, reduced chemical noise and enormous increase of sensitivity. These analytical performances enabled to distinguish the individual components of mixtures of complex glycosphingolipids differing, for instance, only for the position of the sialic acid attachment sites, or the nature of the sphingoid base or fatty acid. Examples of application of ESI technique are the following: (1) the nano-electrospray ionization quadrupole time-of-flight (nano-ESIQTOF) MS or MS/MS, that succeeded in differentiating in ganglioside mixtures from small brain samples, the two isomeric monosialogangliosides GM1a and GM1b, the three isomeric disialogangliosides GD1a, GD1b and GD1a, and the three isomeric trisialogangliosides GT1a, GT1b and GT1(d) the last one carrying a Neu5Ac-Neu5Ac-Neu5Ac trisaccharide linked to C-3 of the terminal galactose moiety (Metelman et al., 2001), and (2) negative ion nano-electro spray ionization Fourier transport ion cyclotron resonance mass spectrometry [(-) nano-ESI-FTICR-MS, and sustained off-resonance irradiation collision-induced dissociation mass spectrometry (SORI-CID-MS-MS), by which sialylated, sulfated and glucuronylated glycosphingolipids obtained from bovine cauda equine, or contained (as contaminants) in a purified mixture of trisialogangliosides from normal adult human brain, could be recognized and structurally defined (Vukelic et al., 2005). The sensitivity of these techniques is in the order of few pmol of analyzed sample. Nano-ESI-QTOF-MS was also applied with success to identify and structurally characterize gangliosides separated by off-line capillary electrophoresis, although the sensitivity was around 100 pmol of analyzed sample (Zamfir et al., 2002). Owing to the extremely high resolution power and sensitivity, the most sophisticated MS techniques coupled to HPLC and capillary electrophoresis, enable to establish the sphingolipid composition of a mixture and determine the chemical structure of the individual components. Moreover, they can serve to assess the degree of homogeneity of a sphingolipid preparation obtained from a natural source. Finally they constitute a powerful tool to identify new sphingolipids in lipid extracts of biological materials. Reviews on the application of MS to study sphingolipid structures are available (Sweeley and Dawson, 1969; Adams and Ann, 1993; Costello et al., 1994; Klein and Egge, 1994; Sullards, 2000; Taketomi and Sugiyama, 2000; Metelman et al., 2001; Zamfir et al., 2002; Vukelic et al., 2005). It is worth mentioning that in some instances, the use of glycohydrolases that specifically act on particular glycosidic linkages (a-sialidases, a- and b-glucosidases, a- and b-galactosidases, a- fucosidases, a- and b-N-acetylglucosaminidases, a- and b-N-acetylgalactosaminidases, a- and b-mannosidases, etc.) can easily provide information on the nature of glycosidic linkages and the sequence of saccharides in the glycosphingolipid molecule. The strategy of this analytical approach and the working procedures are described in dedicated reviews (Li and Li, 1977, 1982). Proton nuclear magnetic resonance (NMR) spectroscopy started being successfully employed in sphingolipids (particularly glycosphingolipids) structural investigations when sophisticated computers, high-field superconducting magnets and pulsing programs (two-dimensional NMR spectroscopy, 2D-NMR) became available (Nagayama et al., 1980). It should be reminded that the natural allocation of protons within oligosaccharide chains is potentially well-suited to yield all or most of their primary and secondary structure. However, almost all resonances of the oligosaccharide portions occur between 3 and 5 ppm, regardless of the solvent and, in addition, water derived resonance occurs substantially within this narrow region; therefore, attribution of signals to individual saccharides is practically impossible. Only by reducing spectra into subspectra, each attributable to a single monosaccharide residue, that is by the use of 2D-NMR spectroscopy, it is possible to decipher overlapping envelopes of resonance. This happened in the early eighties of the last century (Bernstein and Hall, 1982; Dabrowski et al., 1982; Prestegard et al., 1982; Gasa et al., 1983). NMR spectroscopic methods succeeded to provide complete and independent

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information concerning the primary structure of sphingolipids, particularly glycosphingolipids, such as: (1) number and chemical characteristics of components (saccharides, long chain bases, fatty acids, phosphocholine in the case of sphingomyelin); (2) saccharide sequence; (3) characteristics of the glycosidic linkages, including a- and b-configurations. Moreover, they enable to inspect the secondary (tridimensional) structure of the molecules and the dynamics around a flexible linkage (presence of different conformers). All of this can be possible with relatively small amounts of sample (order of magnitude, mg), that can be recovered, and without any derivatization. For the inspections concerning primary structure, the glycosphingolipid is dissolved in an organic solvent, like dimethylsulfoxide (DMSO), where the molecules are in the form of dispersed monomers. When the secondary structure and molecular dynamics are under investigation, it is important to insert the glycosphingolipid into micelles of low molecular weight that are suitable to be analyzed by high resolution NMR methods and constitute a good model of biological membranes, the usual residential sites of glycosphingolipids. These small spherical micelles (molecular weight about 12–16 kDa, against the 300–18,000 kDa of a regular ganglioside micelle) can be obtained by mixing the ganglioside with perdeuterated dodecylphosphocholine (molar ratio 1:40), or by transforming the ganglioside into its corresponding lysoderivative that spontaneously aggregates in the form of small spherical micelles (Acquotti and Sonnino, 2000). The present availability of ultra-high field (920 MHz) NMR approaches opens the field to inspections at the secondary structure of sphingolipids and the sphingolipid/ligand interactions (see for details Siebert et al., 2003; Kato et al., 2008). A simple and rigorous introduction to the application of NMR spectroscopy to sphingolipid structural analysis has been given by Yu (1987). Procedural details, practical recommendations and cautions on NMR spectroscopic methods applied to sphingolipids structural and conformational analysis are exposed in authoritative reviews (Dabrowski et al., 1982; Sweeley and Nunez, 1985; Dell, 1987; Koerner et al., 1987; Yu, 1987; Klein and Egge, 1994; Acquotti and Sonnino, 2000; Siebert et al., 2003; Kato et al., 2008).

3.1.2 Immunochemical Methods for ‘‘In Situ’’ Detection of Sphingolipids An early acquisition in molecular immunology was that carbohydrate sequences, present in complex glycoconjugates and homo- or hetero-polysaccharides, can behave as antigenic determinants eliciting the formation of antibodies that are capable to recognize the same sequences (anti-carbohydrate antibodies) (Graf et al., 1965; Cisar et al., 1975). With reference to glycosphingolipids, it was also observed that the immunogenic activity is much higher with neutral glycosphingolipids than with acidic glycosphingolipids, particularly gangliosides (Graf and Rapport, 1965; Rapport and Graf, 1969; Rapport and Huang, 1984). The first approaches used to produce antibodies against sphingolipids, particularly glycosphingolipids, were to inject animals (mice, rats, rabbits, goats, chicken) with crude extracts of sphingolipids, or cells, cell fragments, isolated membranes, or tissue fractions rich in certain sphingolipids, and then purify the produced antibodies (generally polyclonal antibodies) by affinity chromatography or other procedures (Bogoch, 1960; Naiki et al., 1974; Eisenbarth et al., 1979; Rapport and Huang, 1984; Svennerholm, 1987). The immune response was generally much higher using, as immunogens, cell or tissue materials than using glycosphingolipid mixtures, and the specificity difficult to be warranted (Naiki et al., 1974; Kannagi, 2000). Following the introduction of the hybridoma technique for the preparation of monoclonal antibodies (Ko¨hler and Milstein, 1975), the immunization principle to adsorb glycolipids (or bacterial lipopolysaccharides) to cells and to use these cells as immunogens (Galanos et al., 1971) was reintroduced (Young et al., 1979), perfected and standardized to become a general method to produce monoclonal antibodies with high degree of specificity against any glycosphingolipid (Ozawa et al., 1992). This method consists in absorbing the pure glycosphingolipid (a ganglioside in the original description) into acid-treated Salmonella minnesota mutant R595, and inject the antigen-bacteria complex to mice of the inbred strain C34/HcN five times on different days. The spleen cells obtained 3 days after the last injection are fused with a myeloma cell line, the hybridoma fusion colonies screened against the glycosphingolipid used as immunogen, and the positive hybrid cloned by limiting dilution. By this procedure monoclonal antibodies were produced against acidic and neutral glycosphingolipids of the ganglio-, emato-, lacto-, neolacto-, and globoseries, as well as sulfated glycosphingolipids and carbohydrate sequences O-linked to proteins.

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6

Lists of the different anti-glycosphingolipid antibodies, and of the corresponding antigens, are reported by Svennerholm (1984, 1987), Schwarz and Futerman (1996), Kannagi (2000) and by Kawashima et al. (1994) for monoclonal antibodies specific for ganglioside lactones. The great majority of anti-glycosphingolipid antibodies belongs to the IgM class of immunoglobulins, very few to the IgG class (Kannagi, 2000). It is worth mentioning that anti-glycosphingolipid (better anti-carbohydrate) antibodies may be present in the blood serum of normal human individuals (Marcus, 1990), as well as in patients affected by several neuropathies (Quarles, 1989; Latov, 1994). It was also observed the occurrence of molecular mimicry between brain gangliosides and the terminal chains of the lipopolysaccharides of Campylobacter jejuni, that are the same, or very similar to those of gangliosides (Yuki, 1998). The saccharidic terminal chains of the bacterium are potent immunogens (like the gangliosides absorbed to the surface of Salmonella minnesota) and give rise to antibodies that recognize and bind gangliosides of human nervous tissue with a possible pathogenetic effect (Seikh et al., 1998; Yuki, 1998; Goodyear et al., 1999). A crucial point in the application of anti-glycosphingolipid antibodies is their specificity. From the methodological point of view when a single glycosphingolipid, for instance ganglioside GM1, is used as the immunogen the resulting monoclonal antibody is expected to recognize only GM1. This expectation is correct in the majority of cases, supporting the rationale that the entire GM1-oligosaccharide is the epitope responsible for the immune response. However, in other cases the molecular basis of the antigen/antibody interaction is different: the immunogen is a saccharide sequence shared by different gangliosides or glycosphingolipids; therefore, the antibody recognizes different antigens. For example, individual gangliosides of the b-series (carrying a di-sialosyl residue on the inner galactose moiety), namely GD3, 9-O-AcGD3, GD1b, GT1b and GQ1b, were used to generate specific monoclonal antibodies, and the corresponding antibodies assayed with a number of pure gangliosides (Ozawa et al., 1992). The antibodies against 9-O-Ac-GD3, GD2, GD1b, GT1b and GQ1b were found to recognize only the corresponding ganglioside used as immunogen; therefore, they appeared to be quite specific. Conversely, the antibody against GD3 exhibited a strong reactivity against GD3, a moderate activity against GT1a (which belong to the ganglioside of the a-series), and a weak reactivity toward 9-O-Ac-GD3 and GQ1b. Other examples are available. Monoclonal antibodies raised against Fuc-GM1 reacted also with GM1, although with a much lower affinity, making difficult to distinguish in tissue studies Fuc-GM1 from GM1, where the concentration of GM1 is 100-fold higher, or more, than that of Fuc-GM1 (Fredman et al., 1986). The monoclonal antibody A2B5 initially and cautiously proposed to react specifically with ganglioside GQ1b (Eisenbarth et al., 1979) and largely employed to recognize this ganglioside, was later reported to react with a number of different gangliosides (GT3, 9-O-Ac-GT3, GD3, GD2, GQ1c) although with lower affinity (Fredman et al., 1984; Fenderson et al., 1997). A second relevant issue is that antibodies against certain saccharide sequences carried by glycosphingolipids react also with the same sequences linked to glycoproteins (Hakomori et al., 1983; Schwarting et al., 1992). Typical examples are the monoclonal antibodies HNK-1 (Chou et al., 1985) and NGR50 (Yamawaki et al., 1996) that recognize sulfo-glucuronosyl-neolacto-tetraosyl (or exaosyl) sequences present in both glycosphingolipids and glycoproteins particularly myelin-associated glycoprotein (MAG), with a higher specificity for MAG by NGR50. A third important issue is the not unusual cryptic behavior of the carbohydrate head groups of glycosphingolipid, in other words the possible masking effect on glycosphingolipid accessibility at the cell surface exerted by proteins and glycoproteins (Hakomori et al., 1968; Kannagi et al., 1983; Yamawaki et al., 1996), or other glycosphingolipids (Lloyd et al., 1992; Allende and Panzetta, 1995). This behavior explains the experimental evidence (rare but observed) of a negative response of a glycosphingolipid to a ‘‘specific’’ antibody, against the positive finding of the same glycosphingolipid by chemical methods. A final matter of caution, when using anti-glycosphingolipid antibody for direct detection of glycosphingolipids in tissues or cultured cells, is the fixation procedure that should be accomplished only when necessary, also with the purpose to un-mask cryptic glycosphingolipids, but avoiding any loss of glycosphingolipids (Schwarz and Futerman, 1996). For details on the historical background, technical procedures, cautions and recommendations on the use of anti-glycosphingolipid antibodies see the many review articles dedicated to this topics (Rapport and Huang, 1984; Quarles, 1989; Svennerholm, 1987, 1988; Schwarz and Futerman, 1996, 1997, 2000; Kannagi, 2000). Information on the structure of

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anti-glycosphingolipid (particularly gangliosides)-antibodies is also available (Weng et al., 1992; Marcus and Weng, 1994). Notwithstanding the great care needed to interpreting data on glycosphingolipid detection and identification by the use of anti-glycosphingolipid-antibodies, these antibodies constitute anyway a powerful tool to the following: (1) acknowledge the presence of new glycosphingolipids in biological samples, as a first step to elucidate their structure; (2) localize glycosphingolipids in cultured neural cells at the subcellular level, and determine the distribution of glycosphingolipids in the different cells of different areas and regions of the nervous system during the embryonic and postnatal life; and (c) explore the possible function of the individual glycosphingolipids by complexing them with the corresponding specific antibody, in both in vivo and in vitro experimental approaches. Immune-associated localization of glycosphingolipids is accomplished by immuno fluorescence technique using a light fluorescence microscopy, using the specific antibody (more often of murine origin) toward the single glycosphingolipid as the primary antibody, and as the secondary antibody an anti-mouse IgM (or IgG) antibody carrying a fluorescent probe (Molander et al., 1997; Schwarz and Futerman, 2000). When the anti-glycosphingolipid is not of murine origin, the secondary antibody is an antibody toward the antibodies (generally IgM) of the animal used for immunization. Another immune-assisted methodology employed to detect a particular glycosphingolipid carried by cells is flow cytofluorimetric analysis, based on the same principle to recognize the glycosphingolipid carried by the cells with the corresponding antibody and the latter one with a fluorescent anti-antibody against the antibodies of the immunized animal. It should be reminded that besides antibodies, other specific ligands of glycosphingolipids can be employed to detect glycosphingolipid in situ. The most known of these is Cholera toxin holo-enzyme, or its 5-subunits B, that specifically binds very avidly to ganglioside GM1, the KD value ranging from 4.6  1012 (Kuziemko et al., 1996) to 7.3  1010 (MacKenzic et al., 1997). Using GM1, Cholera toxin and a toxin specific peroxidase, conjugated monoclonal antibody (with the enzyme substrate), electron opaque precipitates can be obtained, thereby enabling to explore the cell location of GM1 by electron microscopy (Hansson et al., 1977). Also in the case of Cholera toxin the specificity is not absolute: in fact it can also bind Fuc-GM1 with an affinity comparable to that of GM1 (Masserini et al., 1992), and other glycosphingolipids (GD1b, GM2, GT1b, GD1a, GM3, asialo-GMD) although with a much lower affinity (see Yanagisawa et al., 2006). Therefore care is to be taken, when assessing GM1 expression in cells with the use of Cholera toxin, to confirm the presence of, and to quantify GM1, by chemical methods (Yanagisawa et al., 2006). A final consideration is regarding the use of immune-assisted methods for the quantification of glycosphingolipids on HPTLC plates or solid phase immunometric procedures (ELISA or else). The detection limit is 0.1–1.0 pmol for HPTLC and 0.01–1.0 pmol for solid phase procedures. As a comparison the detection limit on HPTLC using colorimetric methods is 0.1–0.2 nmol. The most sensitive assay procedure, to be applied only to ganglioside GM1 (or Fuc-GM1) is on the basis of the use of Cholera toxin coupled to Horse radish peroxidase: the detection limit is in the range 2.0–5.0 fmol with solid phase method and 0.02–0.05 pmol with HPTLC.

3.2 Regional Cellular and Subcellular Localization of Sphingolipids in the Nervous System of Different Animals 3.2.1 Sphingolipid Composition and Distribution in the Central and Peripheral Nervous Tissue of Vertebrates Systematic explorations on the sphingolipid composition of the central and peripheral nervous tissue of vertebrates started in the middle sixties of the last century and proceeded there after, concomitantly with the development of more and more refined procedures for the isolation, separation and chemical characterization of the individual sphingolipids. Hence, the difficulty in many cases to compare data from the same animal and the same nervous tissue region provided by different authors and the lack of complete comparative patterns from one animal to another one. This situation is exemplified in > Table 6-5, where data from different reliable sources (given in the table legend) regarding the sphingolipid

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6

. Table 6-5 Major sphingolipids present in the nervous system (cerebrum and peripheral nerve) of human, bovine and rat origin. The data refer to adult subjects and are expressed as mmol g1 fresh tissue (/g fresh myelin in the case of myelin). The molecular weight of ceramide, Gal-Cer, sulphatide and sphingomyelin was established as suggested by Norton and Poduslo (1973). Data were obtained from the references given below and were treated in order to be expressed as specified above. This mode of expression was adopted with the aim to facilitate comparison of contents on a molar basis. As reference, cholesterol, the most abundant individual component of neural cell membranes, was considered Cerebrum (frontal lobe)

Human Ganglioside (as bound Neu5Ac) nLc4Cer SGPG (SGGL-1) SGLPG (SGGL-2) Gal-Cer Sulphatide Sphingomyelin Ceramide Cholesterol Bovine Ganglioside (as bound Neu5Ac) Gal-Cer Sulphatide Glc-Cer Lact-Cer Sphingomyelin Cholesterol Rat Ganglioside (as bound Neu5Ac) SGPG (SGGL-1) SGLPG (SGGL-2) Gal-Cer Sulphatide Sphingomyelin Cholesterol

White matter 0.9–1.2

White matter 1.3–2.1

64.5

8.8 2.4 21.0 2.6 56.1

43.5 9.9 6.3–8.8 3.9 59.4–117.7

Absent Absent 31.6–59.6 4.0–11.3 16.9 3.5 142.5–153.5

3.1–3.8

3.3

1.08

1.6

2.8

2.3 0.5 0.06 0.27 10.7

27.7 6.0 0.1 0.52 20.2 172.8

81.6 8.3 0.22 0.85 47.8 138.6

In toto 2.8–3.3

16.7 4.7

Gray matter 3.1–4.6

Peripheral nervea

Myelin (from) Peripheral nerve 0.4–0.75

0.2–0.44

0.027–0.029 Absent Absent

0.015–0.016 0.05–0.09 0.01–0.014

49.9

0.75

Total brain 2.4–3.8 0.04 0.007 11.1–15.8 4.4 1.9–3.6 48.3–66.8

0.52–0.58 0.01–0.11 60.5–75.4 14.8–18.2 8.1–8.6 143.6–151.4

41.2 13.2 29.7 152.6

References: Vanier et al. (1971), Norton and Poduslo (1973), Tettamanti et al. (1973), Ueno et al. (1978), Nagai and Iwamori (1984), Chou et al. (1987), Kohriyama et al. (1987), Ariga et al. (1990), Ogawa-Goto et al. (1992), Yoshino et al. (1993), OgawaGoto et al. (1993), Kotani et al. (1995), Ledeen (1983), Abe and Norton (1979), Fong et al. (1976), Galli and Fumagalli (1968), Smith and Curtis (1979), O’Brien and Sampson (1965), Spritz et al. (1973), Kishimoto et al. (1965), Suzuki (1965a, b) a Sensory or motor nerves, including in some cases the spinal cord

composition of central (cerebrum or full brain) and peripheral nervous system, and isolated myelin, are reported expressing the individual sphingolipids as mmol g1 fresh tissue (or myelin preparation) in order to facilitate comparisons. In human tissues, surely the most completely studied, the most abundant sphingolipid (in moles terms) is Gal-Cer followed by sphingomyelin, sulfatide, ceramide and gangliosides,

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with gangliosides being more enriched in the gray matter. Characteristic is the presence of nLc4Cer, and glucuronyl containing sphingolipid (SGPG/SGG1-1 and SGLPG/SGGL-2) in peripheral nerve (Chou et al., 1987; Kohriyama et al., 1987; Ariga et al., 1990; Ogawa-Goto et al., 1992, 1993; Yoshino et al., 1993). The human pattern is roughly reproduced in bovine and rat nervous system, although with some gaps. Notably, N-glycolylneuraminic acid covers about 40% of ganglioside bound sialic acid in bovine peripheral nerve myelin, whereas it is completely missing in the corresponding human preparation (Fong et al., 1976). Particular attention has been paid to the ganglioside content and patterns of the vertebrate nervous tissue, owing to the peculiar wide compositional variability of these sphingolipids. As shown in > Table 6-6 the total ganglioside content of brain, expressed as mmol Neu5Ac/g fresh tissue, varies from 1.6–3.8 in mammals, to 1.6–3.6 in birds, 0.9–1.4 in amphibians, 0.37–1.70 and 0.07–2.0 in cartilagineous and bony fishes, respectively, indicating a general trend of diminution from higher to lower vertebrates, and great species variability inside each order. The differences become more pronounced when the patterns of individual gangliosides are considered (see > Table 6-7). GM1, GD1a, GD1b, GT1b and GQ1b are the major gangliosides, followed by GD3 and GD2 in mammals and birds, while GT1c, GQ1c, GT3, GD2, GD1b and over-sialylated gangliosides are more represented in amphibians and fishes. Peculiar is the presence in considerable amounts of GM4 in human brain, of fucosylated gangliosides . Table 6-6 Gangliosides content in the brain of different animals, expressed as mmol bound Neu5Acg-1 fresh tissue. Whole brain (including cerebellum) for small animals or small brains; whole cerebrum for human, pig, bovine, rabbit, rat, cat, horse, sheep brains. Data were obtained from the following references: Suzuki (1965a, b), Tettamanti (1968), Vanier et al. (1971), Avrova (1971), Tettamanti et al. (1973), Lees et al. (1977), Ueno et al. (1978), Hilbig and Rahmann (1980), Iwamori and Nagai (1981), Saito and Tamai (1983), Rahmann and Hilbig (1983), Nagai and Iwamori (1984), Avrova et al. (1986) Animal Human Macacus rhesus Pig Cat Bovine Sheep Rabbit Horse Rat Mouse Dormouse Mole Hamster Guinea pig Chicken Thrush Sparrow Pigeon Frog Salamander Triton Tortoise Lisard a

Bound Neu5Ac mmol g1 fresh tissue 2.8–3.3 3.0 3.1 3.3 3.5 3.1 2.0–3.8 3.1 2.8–3.8 3.4 2.75 2.8 1.6 2.2 1.6–3.6 3.6 3.17 2.27 0.9–1.52 0.9 1.0 1.6 1.38

Number of different species examined

Animal Cartilagineous fishes: Squalomorpha: Squaliformes (4)a Batomorpha: Rajiformes (4)a Dasyatiformes (2)a Galeomorpha: Orectolobiformes (1)a Carchariniformes (2)a Bony fishes: Clupeomorpha (11)a Percomorpha (19)a Parapercomorpha (4)a Goldfish

Bound Neu5Ac mmol g1 fresh tissue From 0.37 to 1.28 From 0.57 to 0.98 From 1.61 to 1.70 1.65 From 1.65 to 1.69

From 0.067 to 1.27 From 0.35 to 1.98 0.92–1.4 0.8

3.7 0.60 1.4 1.4 0.5 1.1 1.0 4.9 5.8

3.1

0.3

1.0

3.5 0.4 tr 1.8 0.6 3.0 2.2 2.5 1.0

0.5

11.4

1.5

17.3 13.6 17.5 17.4 9.4 4.3 5.3 8.8 6.1 4.7 4.1 2.5 1.1 16.8 24.6

1.0 1.3

1.0

0.5

0.3 1.05 2.98

20.5 38.0 38.1 22.2 29.9 22.2 31.7 9.2 11.6 4.2 8.2 1.0 1.2 2.5 6.8

0.7

17.8 9.4 9.6 19.7 12.3 12.2 9.2 3.9 7.0 2.9 7.8 18.9 1.4 3.2 9.7 0.40 1.50

1.5

2.0

14.8 16.3 15.9 22.6 24.7 16.9 18.7 14.5 11.2 19.2 28.5 7.3 11.2 19.2 5.1 2.8 7.5 2.8 1.0

6.3 6.9

6.9

6.5

4.2 5.7 2.0 7.9 6.8 9.5 7.8 12.8 17.4 5.6 17.4 8.5 25.7 31.1 16.5 10.1

20.7

1.1 1.0 4.1

6.0 5.5 2.4 3.5 7.0 8.8 10.1 2.6 6.6 1.3 4.3

FucGD1b GD1b GT1a GT1b GT1c GQ1b GQ1c GD3

1.8 6.0 4.1 3.5

3.7 3,5

2.5 3.5

5.8 5.8 3.9 3.6 5.4 5.8 6.9 18.1 6.8 3.3 4.1 4.0 3.2 15.3 10.3

GT3 GD2

3.5 3.2 1.8

3.7 3.2

3.1 6.2 9.2 8.4 15.7 8.5 9.5 18.1 7.6 9.1

0.5 11.8 4.0 8.2 5.4

0.6

0.6

Over-siaGT2 GP1b lylateda

Figures in bold character: individual gangliosides covering more than 10% of the total ganglioside bound Neu5Ac. Figures in bold italic character: individual gangliosides covering from 7 to 10% of the total ganglioside bound Neu5Ac a With six or more sialosyl residues

Animal: Human 2.0 Bovine Pig Rat Mouse Chicken Pigeon Lizard Tortoise Frog Salamander Belone Salmon Torpedo Shark

FucGD1aGM4 GM3 GM2 GM1 GM1b GM1 GD1a GalNac

Individual gangliosides

. Table 6-7 Example of quantitative patterns of the individual gangliosides contained in the brains of different adult animals. Whole brain (including cerebellum) for small animals (or small brains); whole cerebrum for bigger animals (or brains). The relative content of each ganglioside is expressed as % of bound Neu5Ac on the total ganglioside bound Neu5Ac content. Ganglioside quantification was accomplished colorimetrically after (HP) TLC separation. The data were obtained from the following references: Suzuki (1965a, b), Ishizuka et al. (1970), Avrova (1971), Vanier et al. (1971), Tettamanti et al. (1973), Ueno et al. (1978), Avrova et al. (1986), Yu and Iqbal (1979), Chigorno et al. (1982b), Rahmann and Hilbig (1983)

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6 131

132

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Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

(Fuc-GM1 and Fuc-GD1b) in pig brain, the huge amounts of GT1c in fish brain, and the unusually high amounts of GM1 (as compared to other fishy species) in salmon and torpedo brain. An interesting but yet poorly understood issue concerns the alkali-labile species of gangliosides constituted by O-acetyl and lactonized forms of gangliosides (see > Table 6-8). The presence of alkali-labile gangliosides was recognized . Table 6-8 Content of total alkali-labile gangliosides and individual alkali labile gangliosides (9-0-Ac GT1b, 9-0-Ac GQ1b, GD1b-lactone, GT1b-lactone) in the nervous system of different animals. With the exception of human, rabbit and normothermic or hibernating dormouse, the whole brain was analyzed. The contents are expressed as % of bound Neu5Ac on total lipid bound Neu5Ac. Ganglioside quantification was accomplished colorimetrically after (HP) TLC separation. Data were obtained from the references: Sonnino et al. (1983, 1984b, 1988), Chigorno et al. (1984), Riboni et al. (1984)

Human (adult): Temporal lobe (cortex) Cerebellum (whole) Rabbit: Cerebrum (whole) Cerebellum (whole) Rat Mouse Pig Pigeon Bovine Japanese quail Zebra finch Lizard Blindworm Frog Trout Bass fish Cod fish Salmon Gobiidae family Dasyatis pastinaca Raja clavata Shark Scorpena porcus Lamprey Dormouse Normothermic: Cerebrum Olfactory bulb Cerebellum Spinal cord Hibernating: Cerebrum Olfactory bulb Cerebellum Spinal cord –: not detectable; tr: traces

Total alkali Labile gangliosides

GD1b Lactone

GT1b Lactone

9-0-Ac GT1b

9-0-Ac GQ1b

22.67 10.90

3.7 0.55

3.9 tr

2.3 3.3

0.3 1.75

8.1 13.8 11.5 15.9 6.2 10.9 0.5 2.1 3.5 85.0 78.0 64.0 28.0 64.6 24.7 55.0 45.4 40.6 54.1 64.2 75.0 90.0

– – – – – – – – – – – – – – – – – – – – – –

– – – – – – – – – – – – – – – – – – – – – –

2.7 3.8 3.4 4.2 2.4 1.8 – – – – – – – – – – – – – – – –

1.4 3.2 2.3 2.9 0.9 Tr – – – – – – – – – – – – – – – –

12.8 10.2 17.9 30.1

– – – –

– – – –

3.7 4.4 6.7 10.3

0.8 1.2 2.6 2.8

0.0 0.6 0.0 0.0

– – – –

– – – –

– – – –

– – – –

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6

practically in all brain samples that were analyzed for this purpose. It is surprising that in many animals the percentage of alkali-labile forms of ganglioside is more than 60% of the total, reaching 90% in lamprey brain. In some cases (human and several mammals) 9-O-AcGT1b and 9-O-AcGQ1b were recognized and quantified. In the case of human brain (temporal lobe cortex, and cerebellum) also GD1b-lactone and GT1b-lactone were detected. An intriguing matter, worth being further investigated, is the almost complete disappearance of alkali-labile gangliosides from different portions of the nervous tissue of dormouse when moving from the normothermic to the hibernating condition (Sonnino et al., 1984b). The fatty acid composition of brain gangliosides of different vertebrates is presented in > Table 6-9. The most represented species is by far C18:0 that covers from 50 to 92% of the total fatty acid ganglioside content, followed by C16:0 (from 1 to 19%), C20:0 (from 1 to 9.5%), C18:1 (from 1 to 7%), C22:0 (from 1.5 to 5%) and C21:1 (from 0.5 to 11.3%). In human brain C18:0 and C20:0 cover 87.5 and 9.2% of the total fatty acid content, respectively. The predominant long chain bases are dC18:1 and dC20:1 that cover 75–95% of the total long chain base content, followed by C18:0 and C20:0 (Trams et al., 1962; Rosenberg and Stern, 1966; Avrova and Zabelinski, 1971; Ledeen et al., 1973; Ogawa-Goto et al., 1990). Remarkably, as very accurately demonstrated in adult human whole brain (see > Table 6-10), the heterogeneity of the fatty acid and long chain base composition observed in unfractionated gangliosides is also carried by the individual gangliosides. In other words, a single ganglioside, homogeneous in its carbohydrate composition is heterogeneous in the fatty acid and long chain base composition and, most intriguingly, the lipid composition of a single ganglioside (GM3, GM2, GM1, GD1a, etc,) differs from that of another one. Characteristically, GM3 has the lowest relative content of C18:0 (55.5%) and the highest content of C24:1 (up to 11.4%). A feature that is shared by all gangliosides, with the only exception of GM4, is the complete lack of 2-hydroxylated fatty acids (> Table 6-10). Similarly to gangliosides, the fatty acid composition of sphingomyelin is characterized by the prevalence of C18:0, and the complete lack of 2-hydroxylated fatty acids. In sphingomyelin C24:1 is the second most represented fatty acid, as in the case of GM3. Gal-Cer and sulphatide in human brain have a very similar fatty acid composition characterized by a preponderance (about 60% of the total content) of the 2- hydroxylated species, with C24:1 as the most abundant species (30–35% of the total content), followed by C24:0 (the richest in the 2-OH species), C18:0 and C25:1. Owing to the presence of 2-hydroxylated fatty acids, the abundance of C24:1 and C24:0, and the occurrence of C25 and C26, species, the fatty acid composition of GM4 is much more similar to that of Gal-Cer and sulphatide than that of the other gangliosides. Remarkably, as reported by Ogawa-Goto et al. (1990), both the fatty acid and the long chain base compositions of the main gangliosides (GM1, GD1a, and GD1b) of human peripheral nerves are markedly different from that of the same gangliosides of human whole brain. In the case of fatty acids, C18:0 is much less abundant, whereas shorter (C14:0, C16:0, C16:1) and longer chain (from C22:0/1 to C25:0/1) fatty acids are much more represented. Regarding the long chain bases, the proportion of the 18:0 species is higher. Moreover, clear differences in fatty acid and long chain base compositions are also present between motor and sensory nerves: the differential trend described above is much more pronounced in sensory than motor nerves. All this contributes to provide the message that the ganglioside expression in different regions of the nervous system of the same animal (human) is different at a higher or lower degree, in both the saccharide and lipid portions suggesting the occurrence of specific biosynthetic and degradative machineries that operate locally.

3.2.2 Regional, Cellular and Subcellular Localization of Sphingolipids in the Nervous Tissue of Vertebrates In addition to the investigations on the sphingolipid composition of gray and white matter of central nervous tissue, peripheral nervous tissue and isolated myelin, in adult animals, regional inspections were also performed on cerebellum, in comparison with cerebrum, on different cerebrum areas and on other brain regions with particular attention to the ganglioside components. This topic has been covered by excellent reviews, although not recent (Ledeen and Yu, 1982; Ando, 1983; Ledeen, 1983; Nagai and Iwamori, 1984; Svennerholm, 1984). The total ganglioside content, expressed as mmol bound Neu5Acg-1 fresh tissue,

133

Ray Carp Shark Alligator Frog Tortoise Chicken Pigeon Turkey Porpoise Rat Guinea pig Rabbit Pig Bovine Monkey Human 1.2 1.0 – 1.0 0.7 2.0 tr tr – – tr tr tr tr tr tr tr 6.4 14.3 5.0 7.0 19.3 12.7 5.5 4.6 – 1.0 3.0 3.2 2.8 4.0 3.1 1.8 1.1 0.7 1.4 – – 8.9 tr 0.5 1.0 – – tr 0.5 2.0 – tr – – 0.9 tr – – 1.9 0.7 tr tr – – tr 0.7 tr – tr – – 72.7 57.1 72.0 79.0 50.1 61.5 79.2 78.6 96.0 80.0 83.3 92.0 87.2 82.0 86.0 87.7 87.5 1.8 3.4 – – 7.0 3.3 2.6 2.1 – – 1.6 1.0 Tr – 0.9 – 0.1 3.3 2.9 1.0 4.0 2.5 6.6 5.3 2.0 2.0 14.0 9.5 1.3 6.2 7.0 3.2 6.5 9.2 2.4 2.5 5.0 2.0 2.1 2.9 2.3 1.5 2.0 – 1.6 1.3 1.2 2.0 1.0 1.8 0.6 4.7 tr 4.0 – tr 0.9 tr tr – 1.0 tr tr tr – 0.5 0.8 – tr tr – – Tr tr 0.5 tr – – 1.0 tr 0.7 – 0.6 – 0.1 0.9 6.1 – – 1.3 0.6 1.1 4.3 – 1.0 tr tr tr 3.0 2.4 – 0.1 4.0 11.3 13.0 7.0 6.3 8.8 3.0 5.9 – 2.0 tr tr tr 1.0 0.7 – 0.5 – – – – – – – – – – – – – 1.0 1.6 0.8 –

tr: traces; –: not detectable

Fatty acid C14:0 C16:0 C16:1 C17:0 C18:0 C18:1 C20:0 C22:0 C22:1 C23:0 C24:0 C24:1 C24:2

Animal

6

. Table 6-9 Fatty acid compositions of the brain gangliosides of different vertebrates. Each fatty acid is expressed as % of the total fatty acid content of gangliosides. Whole brains were employed. Data were obtained from the references: Trams et al. (1962), Rosenberg and Stern (1966), Avrova and Zabelinski (1971), Ando and Yu (1984). The most abundant fatty acids in the brain of each animal are given in bold characters

134 Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

/ / 3.5/ / / / 61.6/ 4.5/ 3.5/ / 1.6/ / 10.3/ / 6.4/ 0.5/ / / / /

55.6 2.5 40.2 1.7

/ / 8.5/ / / / 55.5/ 5.1/ 4.0/ / 2.3/ / 5.0/ / 3.5/ 11.4/ / / / /

80.1 3.1 16.0 0.8

GD3

42.0 18.9 32.3 6.9

/ / tr/ / / / 85.8/ tr 13.2/ / 0.2/ / / / tr 0.1/ / / / /

GM2

41.9 8.3 42.4 7.4

/ / tr/0 / / / 80.6 tr 15.9 / 1.5 / / / 0.1/ 1.1/ / / / /

GD2

55.5 2.0 40.0 2.0

/ / 0.4/0/ / / / 92.6/ tr 6.9/ / 0.1/ / / / / / / / / /

GM1

42.1 3.1 50.8 4.0

/ / tr/ / / / 89.4/ tr 10.1/ / 0.5/ / / / / / / / / /

GD1a

38.4 2.0 55.7 3.9

/ / 0.3/ / / / 88.5/ tr 10.1/ / 0.6/ / / / / / / / / /

GD1b

64.1 7.8 21.2 3.9

/ / 5.9/ / / / 73.8/ tr 15.2/ / 1.4/ /. / / / / / / / / 35.2 2,2 55.1 7.5

/ / 2.6/ / / / 86.8/ tr 9.2/ / 0.6/ / / / / / / / / /

GD1bfuc GT1a

35.2 2.2 55.1 7.5

/ / 2.3/ / / / 81.8/ tr 12.1/ / 0.8/ / / / / / / / / /

GT1b

47.1 1.4 48.8 2.7

/ / 0.9/ / / / 90.5/ tr 8.2/ / 0.3/ / / / / / / / / /

GQ1b tr/0.12 0.08/12 8.6/0.06 0.9/ 0.24/0.06 0.16/ 4.4/0.18 5.3/ 0.2/0.06 0.08/ 0.5/2.9 0.04/ 0.8/8.8 0.04/ 4.4/25.7 9.6/10.3 0.7/4.1 2.0/1.9 0.3/tr 1.7/3.1

GM4

Sulfatide

Grey matter /0.06 0.12/0.24 1.0/ / 0.08/0 / 0.24/0.12 0.8/0.36 6.8/ tr/ 0.3/ / / 0.04/ / / / / 5.5/0.7 1.4/0.36 61.9/ 0.08/ 0.6/ 2.6/ 0.4/0.12 0.12/ 2.7/ / 0.04/ / 1.2/4.8 0.6/3.1 1.1/ 0.04/ 0.12/ / 1.8/9.5 1.2/8.8 1.5/ 0.3/ 0.3/ 0.5/ 6.3/22.8 5.7/25.2 1.9/ 18.4/10.8 19.2/14.9 12.0/ 0.9/2.3 1.4/2.5 0.4/ 3.1/1.4 4.1/2.0 2.5/ 0.12/0.12 0.5/0.54 tr/ 1.8/6.2 3.6/2.3 2.4/ Gal-Cer

White matter tr/ / 10.2/ / / / 20.1/ 6.5/ 1.1/ / 1.7/ / 2.8/ 1.3/ 6.9/ 30.2/ 3.4/ 8.3/ tr/ 5.5/

Sphingomyelin

6

tr: traces; : not detectable

C14:0/C14:0 OH C15:0/C15:0 OH C16:0/C15:0 OH C16:1/C16:0 OH C17:0/C17:0 OH C17:1/C17:1 OH C18:0/C18:0 OH C18:1/C18:0 OH C20:0/C20:0 OH C20:1/C20:1 OH C22:0/C22:0 OH C22:1/C22:1 OH C23:0/C23:0 OH C23:1/C23:1 OH C24:0/C24:0 OH C24:1/C24:1 OH C25:0/C25:0 OH C25:1/C25:1 OH C26:0/C26:0 OH C26:1/C26:1 OH Long chain base dC18:1 dC18:0 dC20:1 dC20:0

GM3

Gangliosides

. Table 6-10 Fatty acid and long chain base composition of the individual gangliosides isolated from adult human whole brain (cortex gray matter and white matter, in the case of sphingomyelin). Each fatty acid is expressed as % of the total fatty acid content. The fatty acid composition of ganglioside GM4, Gal-Cer and sulfatide of myelin prepared from brain white matter is also given for comparative purposes. For each fatty acid the 2-hydroxylated derivative (OH) is also indicated. Data were obtained from references: O’ Brien and Sampson (1965), Ledeen et al. (1973), Ando and Yu (1984) Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids 135

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was found to be similar in cerebellum and cerebrum: 2.1 and 1.7 in mouse, respectively, 2.7–3.0 and 2.8 in rat; 2.8 and 2.7 in rabbit, and 2.6–2.8 and 2.6–2.9 in humans (Suzuki, 1965a, b, 1970; Roukema et al.; Martinez and Ballabriga, 1978; Seyfried et al., 1982; Saito and Tamai, 1983; Schaal et al., 1985; Svennerholm et al., 1989; Kracun et al., 1992). Differences in the total ganglioside content were also observed in different areas of cerebrum (frontal cortex, occipital cortex, hippocampus) (Suzuki, 1965a, b; Saito and Tamai, 1983; Schaal et al., 1985; Kracun et al., 1992; Svennerholm et al., 1989). Data concerning this topic, having a particular historical value, are presented in > Table 6-11. As shown, the examined brain regions rich in gray

. Table 6-11 Gangliosides content in different parts of the central and peripheral nervous system from adult human subjects. Brain specimens from the left hemisphere, with the exception of midline structures. Ganglioside content is expressed as mmol bound Neu5Acg-1 fresh tissue Tissue Frontal cortex Precentral gyrus Postcentral gyrus Superior temporal gyrus Visual cortex Cerebellum Caudate nucleus Globus pallidus Thalamus

Content 3.11 2.83 3.02 3.33 2.84 2.13 3.13 2.49 2.83

Tissue Uncus Trigonal gyrus Cingulate gyrus Centrum semiovale Centrum callosum Spinal cord Sciatic nerve Femoral nerve

Content 3.05 3.31 2.97 0.53 0.61 0.29 0.27 0.11

In frontal cortex, precentral/superior/temporal gyrus, visual cortex, cerebellum, caudate nucleus and globus pallidus, the right hemisphere carries a higher ganglioside content (from a minimum of 6% in superior temporal gyrus to a maximum of 25% in precentral gyrus), as compared to the left hemisphere. Elaboration from the original data provided by Suzuki (1965a, b), Yu and Ledeen (1970), Ueno et al. (1978).

matter have a ganglioside content ranging from 2.1 to 3.3 mmol g1 fresh tissue, whereas areas like centrum semiovale and centrum callosum, rich in white matter, have much lower amounts (0.15–0.6 mmol g1 fresh tissue). Peculiarly, retina was found also to be enriched in gangliosides, although less than cerebrum and cerebellum (in the same animal), the content ranging from 1.9 mmol bound Neu5Ac/g fresh tissue in calf to 0.6 in frog, with intermediate levels in rat (1.6), chicken (1.4) and duck (1.2) (Dreyfus et al., 1976). The ganglioside composition markedly changes from one to another brain region or area. An example, referred to human brain, is given in > Table 6-12. In gray matter GD1a and GD1b prevail and the total content of disialogangliosides covers 53.6% of total gangliosides (as bound Neu5ac), total tri-and tetrasialogangliosides and total mono-sialogangliosides covering 23.2%, each. In the white matter GM1 is the most abundant ganglioside and total mono-sialogangliosides rise to 37.5%. In isolated myelin, GM1 goes up to 47.8% and GM4 to 18.4%, with total mono-sialogangliosides covering 74% of total gangliosides. Neurons, isolated from cortex gray matter have a ganglioside composition similar to that of the gray matter, and oligodendroglial cells, prepared from white matter-rich brain areas feature a ganglioside composition close to that of white matter. This is consistent with the notion that GM1 and, particularly, GM4 are characteristic myelin gangliosides, as first proposed by Ledeen et al. (1973). The differences in the ganglioside composition between cerebrum and cerebellum were ascertained by many investigators and are summarized in > Table 6-13. The most evident features are that in all reported animals: (1) GD1a is the most abundant ganglioside in cerebrum and GT1b in cerebellum; (2) GM1 is more abundant in cerebrum than in cerebellum; (3) GQ1b is more abundant in cerebellum than in cerebrum; and (4) multi-sialylated gangliosides are more expressed in cerebellum. A further feature of cerebellum, as compared to cerebrum, is a much higher content of alkali- labile gangliosides (species containing O-acetylated gangliosides and gangliosides in the lactone form). In the case of rabbit, 17% of total cerebellum gangliosides (as bound

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6

. Table 6-12 Ganglioside pattern of adult human brain gray matter, white matter, isolated neurons, oligodendroglial cells and myelin. Gray matter and neurons were prepared from cerebral cortex, white matter and oligodendroglial cells from corpus callosum and centrum semiovale. The individual gangliosides are expressed as % of bound Neu5Ac over total ganglioside bound Neu5Ac. Elaboration from the original data provided by Yu and Iqbal (1979) Biological material Ganglioside GM4 GM3 GM2 GM1 GD3 GD2 GD1a GD1b GT1a GT1b GQ1b Monosialo-gangliosides Disialo-gangliosides Tri- and tetrasialo-gangliosides

Gray matter 1.5 2.7 4.1 14.9 5.5 8.0 21.8 18.3 1.8 16.3 5.1 23.2 53.6 23.2

Neurons 0.9 4.1 4.5 22.5 7.2 3.9 21.5 20.7 1.3 11.3 2.2 32.1 53.3 14.8

White matter 8.6 4.8 2.5 21.6 8.8 3.1 17.7 16.9 2.2 11.1 2.7 37.5 46.5 16.0

Oligodendroglia 5.9 8.1 5.7 20.1 11.9 3.3 16.2 14.9 2.7 9.3 1.9 39.8 46.3 13.9

Myelin 18.4 1.6 6.2 47.8 2.8 1.4 8.2 11.2 2.2 0.2 74.0 23.6 2.4

The most abundant gangliosides in each analyzed material are given in bold character

. Table 6-13 Distribution of major individual gangliosides in the cerebrum and cerebellum of different adult animals. The content of each ganglioside is expressed as % of bound Neu5Ac on total bound Neu5Ac content. Quantification was accomplished colorimetrically after (HP) TLC separation. Elaboration of data from: Suzuki (1965a, b), Seyfried et al. (1982), Ledeen (1983), Chigorno et al. (1984), Schaal et al. (1985), Svennerholm et al. (1989) Major individual gangliosides Animal Mouse Rat Rabbit Human

Brain region Cerebrum Cerebellum Cerebrum Cerebellum Cerebrum Cerebellum Cerebrum Cerebellum

GM3 – – 1.8 tr – – – –

GM1 9.4 5.2 17.4 4.5 18.5 7.8 17.4 5.5

GD1a 29.9 11.5 22.2 31.2 51 14.8 39.6 20.9

GD1b 12.3 10.7 19.7 8.8 7.2 16.1 19.8 26.1

GT1a 10.1 – – – 2.3 – –

GT1b 24.7 39.1 21.6 30 14.7 31.7 15.9 42

GQ1b 6.8 24.2 7.9 13.6 1.8 9.5 2.9 4.6

GD3 7 3 3.5 3.5 – – – –

GD2 5.4 – 3.6 7.9 – – – –

The most abundant gangliosides in each material are given in bold character

Neu5Ac) is represented by the alkali-labile species, against 8% in cerebrum (Chigorno et al., 1984). Moreover, the adult mouse cerebellum, that carries about 0.04 mmol g1 fresh tissue (as bound sialic acid) of disialosyl-lacto-N-neotetraosylceramide LD1 (a high concentration for this ganglioside !) (Chou et al., 1990), contains O-acetyl-LD1 (about half the content of LD1) and, in lower amounts, O-acetyl-GD1a, O-acetyl-GT1b, O-acetyl-GQ1b and likely GD1a-lactone, all of them being alkali-labile forms of gangliosides (Chou et al., 1990). Notably, all these ganglioside species are not present in adult mouse cerebrum

137

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Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

(Chou et al., 1990). Also, the glycosphingolipids containing sulfated glucuronic acid (SGGL-1, SGGL-2), although minor constituents in mouse nervous system, appear to be more abundant in cerebellum than in cerebrum (Nair et al., 1998). Considerable attention was also paid to the ganglioside composition of retina. As shown in > Table 6-14, where the reliable available data were collected, the most remarkable feature is the extremely high content of GD3 (from 12 to 50% of total bound Neu5Ac, depending on the animal). A second peculiar feature is the relatively low percentage of mono-sialogangliosides (from 2.3 to 18%) and high percentage of tri- and tetra-sialogangliosides (up to 70% in frog retina). The availability of specific antibodies (against individual sphingolipids) opened the possibility to directly investigate the cellular location of the same sphingolipids in different brain regions, owing to the high sensitivity and versatility of the immunochemical detection procedure, although quantification could be only approximate. An elegant example of this analytical approach is given in > Table 6-15, reporting the distribution of gangliosides in different cell layers or portions of rat cerebellar cortex, cerebral cortex, hippocampal formation, and spinal cord. It is a consolidated notion that all sphingolipids, with the exception of sphingosine, sphingosine-1phosphate and ceramide-1-phosphate, are typical membrane components, particularly of the plasma membrane. Glycosphingolipids, particularly gangliosides, are constituents of the cell glyco-calix, exposing their saccharide moieties on the external leaflet of the plasma membrane (Wiegandt, 1967). The better characterized plasma membrane preparations obtained from the brain of different animals are those surrounding isolated nerve terminals (synaptosomes). These plasma membranes (synaptic or synaptosomal membranes) have a ganglioside content ranging from 7.3 to 45.2 mg bound Neu5Acmg-1 mg protein, with a much higher content in cholinergic than non-cholinergic terminals (Wiegandt, 1967; Breckenridge et al., 1972; Avrova et al., 1973) and with a compositional pattern close to that of the brain from which they were prepared (Waki et al., 1994). Instead, synaptic vesicles have, in general, lower ganglioside contents (2.9–5.1 mg bound Neu5Acmg-1 protein) (Lapetina and De Robertis, 1968; 1973Breckenridge et al., ), with the exception of those prepared from Torpedo electric organ, which feature a ganglioside content doubling that of synaptosomes (Ledeen et al., 1988). The ganglioside composition of these synaptic vesicles is characterized by a large abundance (about 70%) of tri-, tetra- and multi-sialylated species (Ledeen et al., 1988). Further location sites for sphingolipids are the intracellular structures where their metabolic pathways and intracellular traffic take place (endoplasmic reticulum, Golgi apparatus, lysosomes, transport vesicles). Remarkable is also the reported presence of gangliosides, Glc-Cer and neutral glycosphingolipids, in the growth cone membrane prepared from 16 to 18 day old fetal rat brain (Sbaschnig-Agler et al., 1988). The ganglioside content almost doubled that of the synaptic vesicles obtained from the same source, and the content of Glc-Cer equalled that of gangliosides in molar terms. The ganglioside composition of growth cone membranes was similar to that of synaptic vesicles and resembled that of the rat whole brain of the same embryonal age (Yu et al., 1988). No more recent data on this very stimulating topic are available. The nucleus is also a site of ganglioside location mainly at the nuclear membranes. The most abundant nuclear ganglioside is GM1, followed by GD1a, GD1b, and GT1b, with smaller amounts of GM3 and GD3 (Wu et al., 1995; Saito and Sugiyama, 2002; Ledeen and Wu, 2006). A detailed and updated review on nuclear sphingolipids is the object of chapter 7 of this volume. The targeting of neo-biosynthesized individual sphingolipids to their residential sites or from the residential places to the degradation sites implies the presence of ‘‘ad hoc’’ transport systems, operating by means of vesicles or transport proteins. A peculiar transport protein is CERT, that carries ceramide and has been fully characterized (Hanada et al., 2003). Evidence for the presence of transport proteins for gangliosides has been also provided, as the cytosoluble gangliosides, which represent a very small but definite fraction of gangliosides in brain, are sensitive to the conventional procedures for protein fractionation and precipitation (Sonnino et al., 1979; Ledeen et al., 1981; Gammon and Ledeen, 1985). No ganglioside transporting protein has been isolated so far. It is not known whether transport vesicles or proteins are implicated in the antero-grade and retro-grade axonal flow processes operating in ganglioside intraneuronal delivery (Ledeen et al., 1981, 1987; Aquino et al., 1985). Details on the cellular and subcellular aspects of sphingolipid cell biology are available in dedicated reviews (Ledeen, 1978, 1979, 1983, 1989; Yamakawa and Nagai, 1978; Ledeen et al., 1981; Nagai and Iwamori, 1995).

nmol 1.0 20.7 n.d. 13.3 2.9 12.2 44.2 43.2 32.5 n.d.

(%) 0.6 12.2 – 7.8 1.7 7.2 26.0 25.4 19.1 –

nmol 53.0 65.3 3.0 n.d. 36.0 138.7 38.6 59.4 23.3 6.8

(%) 12.5 15.4 0.6 – 4.9 31.2 8.7 14.0 5.5 1.6

Chicken nmol 24.0 68.8 2.4 n.d. 17.8 113.6 31.7 65.1 29.5 11.3

Duck (%) 6.6 18.9 0.6 – 4.9 31.2 8.7 17.9 8.1 3.1

nmol 29.5 176.3 n.d. n.d 13.0 57.0 85.5 73.9 47.8 n.d.

The most abundant gangliosides in each animal are given in bold character. n.d. = not detectable *These data are approximate

Ganglioside GM3 GD3 GM2 GD2 GM1 GD1a GD1b GT1b GQ1b GQ1c (?)

Frog

Rat (%) 6.1 36.5 – – 2.7 11.8 17.7 15.3 9.9 –

nmol 28 287.5 – – – 63.2 74.7 87.4 – –

Rabbit* (%) 5 50 – – – 12 13 15 – –

nmol 5.8 258.2 n.d. n.d 19.1 79.3 86.3 93.8 36.5 n.d.

Calf (%) 1 44.6 – – 3.3 13.7 14.9 16.2 6.3 –

nmol (18) 183 – – – 43.9 47.6 54.9 – –

(%) (5) 50 – – – 12 13 15 – –

Human*

. Table 6-14 Distribution of individual gangliosides in the retina of different animals. Each ganglioside is expressed as nmol bound Neu5Acg-1 fresh tissue and as % of total ganglioside bound Neu5Ac. From the original data by Holm et al. (1972) and Dreyfus et al. (1975, 1976)

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6 139

Brain region Cerebellar cortex Molecular layer Parkinje cell layer Granular layer White matter Cerebral cortex Molecular layer (I) External granular layer (II) External pyramidal layer (III) Internal granular layer (IV) Internal pyramidal layer (Va/Vb) Polymorphic cell layer (VI) White matter Hippocampal formation Hippocampus: Alveus Stratum oriens Stratum pyramidale

GD1a 3+ – 1+ – 3+ 3+ 3+ – 1+/– 1+ – – 2+ –

GM1

1+ – 1+ 3+

1+ 1+ 1+ 1+ 1+/1+ 1+ 2+

2+ 2+ –

Major gangliosides

2+ – 1+

– 1+ 2+ 3+ 3+/3+ 3+ 1+

1+ – 3+ –

GD1b

2+ 1+ –

1+ – 2+ 1+ 2+/2+ 1+ 3+

2+ – 3+ +

GT1b

3+ 1+ 2+

– – – 1+ –/1+ 1+ –

– – 2+ –

GQ1b

3+ – 1+

– – – 1+ 1+ 3+ 2+

– – 1+ 3+

GM3

2+ – 3+

– 1+ 1+ 2+ 1+ 1+ 3+

– – 2+ 2+

GD3

Minor gangliosides

– 2+ 3+

– 1+ 1+ 1+ 2+ 1+ –

3+ 3+ – –

O–AcDGa

– 2+ –

1+ 1+ 1+ 1+ 2+ 2+ –

1+ – 3+ –

GD2

2+ – –

– – – – – – 2+

1+ – 1+ 1+

GM4

6

. Table 6-15 Distribution of major and minor gangliosides in different regions of rat nervous system. The approximate amounts of gangliosides were estimated by an indirect immunofluorescence technique, using specific monoclonal antibodies for each ganglioside. Data selected from references Kotani et al. (1993, 1994)

140 Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

2+ – 2+ – 2+ nd – nd 3+ nd – nd – nd –

– 2+ 1+ – 1+

nd 1+ nd 1+ nd 1+ nd 1+ nd 1+

nd 3+ nd 3+ nd 1+ nd 1+ nd 1+

– 2+ 1+ 2+ – nd 3+ nd 3+ nd 2+ nd 2+ nd 2+

1+ 1+ 1+ – 2+ nd 2+ nd 2+ nd – nd – nd –

– – – – –

a

3+, strong reactivity; 2+, moderate reactivity; 1+, weak reactivity; -, negative reactivity; nd, not determined O-AcDG, O-acetylated disialogangliosides

Stratum radiatum Stratum lacunosum–moleculare Dentate gyrus: Polymorphic layer Granular layer Molecular layer Spinal cord Gray matter: Dorsal horn Anterior horn Ventral horn Posterior horn Intermediate zone White matter: Anterior funiculus Ventral funiculus Lateral funiculus Dorsal funiculus Posterior funiculus 3+ nd 3+ nd 3+ nd – 1+ 1+ nd

– 3+ – – – 3+ nd 3+ nd 3+ nd 3+ 3+ 3+ nd

– 2+ 1+ 3+ – 1+ nd 1+ nd 1+ nd – – – nd

– 1+ 1+ 1+ – 3+ nd 1+ nd 1+ nd – – – nd

2+ – 1+ – 1+ 1+ nd 1+ nd 1+ nd 1+ 1+ 1+ nd

– – – – – Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

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3.3 Development Profiles of Sphingolipids in the Nervous System of Different Animals The development of the nervous system is one of the most complex events in morphogenesis and does not differ essentially with animal species. It can be divided into different stages characterized by the occurrence of more or less irreversible processes in temporal sequences. These stages are as follows (Ro¨sner and Rahmann, 1987; Ro¨sner et al., 1992): (1) neural induction with formation of neural tubes; (2) proliferation of neuroblasts and glioblasts; (3) neuronal differentiation, growth cone formation and neuritogenesis, short distance migration of newborn neurons, and beginning of neuronal/glial interactions; (4) axonal and dentritic arborization, synaptogenesis, fibre tract mapping over short and long distances, trophic dependency, selective apoptosis; (5) further extension of neuronal connections, second period of glial proliferation, onset of myelination, continuing rearrangement involving later-born microneurons; (6) maturation of the functional neuronal network and of myelin; and (7) ageing. In a previous, less detailed approach, four stages were proposed, where the above stages (3) and (4) were unified into a single one, and the above stages (5), (6), and (7) into a unique stage too. The extent and duration of these stages are different among animal species with shorter or longer periods of overlapping and different exposures of the various stages to prenatal and post-natal age. In humans the approximate duration of stage (1) is between 3rd/4th and 8th gestational weeks; stage (2) from 8th to 25th gestational weeks; stage (3) from 25th gestational week to 4 months of age; stage (4) and (5) from birth to 5 years; and (6) and (7) thereafter. Of course, owing to the heterogeneity of neuronal and glial cells, each cell type may have its own crucial period for proliferation, differentiation, maturation and survival. Furthermore, neural cells feature the highest surface/cell mass ratio and the richest network of functional cell–cell interactions. Hence, they offer the highest relative content of complex lipids and particularly of sphingolipids, including among them glycosphingolipids that are molecularly suitable to be engaged in cell–cell interactions.

3.3.1 Developmental Profiles of the Main Sphingolipid Components of Brain The first comprehensive attempt to establish, with the biochemical methodology available at that time, the sequential lipid changes occurring in the brain during development was performed by Brante (1949). FolchPi (1955) also correlated the lipid biochemical changes during brain maturation with the morphological events pertaining to gray and white matters. Wells and Dittmer (1967) were the first to propose the occurrence in brain development of four stages of morphological/functional events in an irreversible sequence, on the basis of the assumption – later proven not to be fully correct – that a given morphological structure has a constant lipid composition. The changes in the lipid composition of the nervous system during development were the object of hundreds of publications in the following couple of decades, with particular emphasis to sphingolipids, because of their highly diversified chemical compositions. The impact of neuronal growth and maturation and of the myelination process on the content of total phospholipids, cholesterol, sphingomyelin, Gal-Cer, sulphatide and gangliosides in rat whole brain during post-natal development is illustrated in > Figure 6-5. These parameters are expressed as mmol g1 fresh tissue in order to put in evidence their actual accumulation with age. As shown, the increase of concentration from birth to adulthood is about twofold for total phospholipids, threefold for gangliosides, fivefold for cholesterol, tenfold for sphingomyelin and 40-fold for sulphatide and Gal-Cer. The more typical components of myelin, sulphatide, Gal-Cer and sphingomyelin, exhibit the highest accumulation rate concomitantly with the formation and maturation of myelin, processes that occur in rat after birth. The post-natal accumulation of gangliosides and of sphingomyelin, at a lesser extent, is the continuation of a trend already present in the embryonal life, reflecting neuronal growth, neuritogenesis, dendride arborization, as well as glial proliferation. Total phospholipids cover more than 60% of the total lipids present in brain close to birth, and only 45% in the adult brain. A more detailed picture of the effect of age, particularly pre-natal and early post-natal (till 2 years), on the content of the same lipid parameters considered above in the gray and white matter of human brain (whole brain from fetuses of 10–12 gestational weeks; frontal lobe for all the other fetuses and all post-natal

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6

. Figure 6-5 Effect of post-natal age on the distribution of total phospholipids, cholesterol, Gal-Cer, sphingomyelin, sulphatide and gangliosides in the whole brain of rat. All substances are expressed as mmol g1 fresh tissue (in the case of gangliosides as mmol bound Neu5Ac/g fresh tissue). Elaboration from original data provided by Suzuki (1965a, b), Cuzner and Davison (1968), Roukema et al. (1970), Vanier et al. (1971), Rouser et al. (1972), and Norton and Poduslo (1973). The average gangliosides in rat brain contain 2 mol of Neu5Ac/mol of gangliosides; however this ratio changes with age

subjects) is shown in > Figure 6-6. In gray matter, total phospholipids, cholesterol and gangliosides undergo a continuously increasing accumulation from the 2nd gestational month till 2 years, whereas sphingomyelin accumulation starts being evident in the late second trimester of pre-natal life. This indicates that in the stages of more intensive proliferation and maturation of neurons and glial cells the ratio between ganglioside and sphingomyelin changes. In white matter the accumulation of cholesterol, total phospholipids, Gal-Cer, sphingomyelin and sulphatide begins around birth and continues till 2 years, with a steep rate from birth to the 3rd and 4th months, reflecting the impact of the myelination process. In white matter the presence of gangliosides becomes relevant after birth, again concomitantly with myelination. Around the 10th month after birth the accumulation rate of all the lipid parameters remains constant, likely corresponding to the consolidation of some basic morphological structures, beginning to increase there after.

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. Figure 6-6 Effect of age (pre-natal and post-natal) on the distribution of total phospholipids, cholesterol, Gal-Cer, sphingomyelin, sulphatide and gangliosides in the human cerebral cortex (gray matter and white matter). All substances are expressed as mmol g1 fresh tissue (in the case of ganglioside as mmol bound Neu5Acg-1 fresh tissue). Elaboration from original data provided by Suzuki (1965a, b), Vanier et al. (1971, 1973), Svennerholm and Vanier (1972), Martinez and Ballabriga (1978), Svennerholm et al. (1989, 1991), Soderberg et al. (1990). g.m.: gray matter; w.m.: white matter

3.3.2 Developmental Profiles of Individual Gangliosides and Other Glycosphingolipids The modifications of the brain ganglioside composition during pre-natal and post-natal life were studied by many investigators in different animals. Examples of the results obtained, chosen with the aim to put in evidence general trends, or peculiar features, are presented in the following figures. > Figure 6-7 deals with the development profiles of gangliosides in the brain of chicken, an animal species where the first four stages of neural development, including the onset of myelination, occur in pre-hatching life. The ganglioside content is expressed in nmol bound Neu5Ac/g fresh tissue. The general trend of the major gangliosides GD3, GT1b, GD1a, GQ1b, GP1c, GM1, GT2, GT3, GT1a, GQ1c (with the notable exception of GD3) and of

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6

. Figure 6-7 Distribution of individual gangliosides in whole chicken brain during pre- and post-hatching life. Each ganglioside is expressed as mmol bound Neu5Acg-1 fresh tissue. Elaborations from original data provided by Dreyfus et al. (1975), Ro˝sner (1980, 1982), Ro˝sner et al. (1988), Sonnino et al. (1990) and Lehmann et al. (2003)

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the minor ganglioside GM2, is a steep accumulation from the 6th day of pre-hatching life, picking up on the 8th day, followed by a sharp decrease till hatching, and maintenance of a constant level thereafter (only GT1b exhibits a slow decrease). All this reflects intense processes of neural and glial proliferation, maturation and formation of an intercellular network of arborization. Peculiar is the behavior of GD3 that features high levels from day 6 to 9 of pre-hatching life, followed by a marked decrease till day 14 and then a constant increase of accumulation up to 80 days of post-hatching life. GM3, GM1 and GD1a undergo a rapid increase of content close to hatching, and then GM1 maintains a constant level till the 80th day, whereas GM3 and GD1b display a constant and moderate increase till the 40th day followed by a further rise till the 80th day in the case of GD1b, or marked decrease in the case of GM3. In the early pre-hatching age the most abundant ganglioside is GD3, followed by GT1b and GQ1b; in the picking up period just before hatching it is GT1b, followed by GD1a and GD3; on the 80th day after hatching it is GD1a followed by GD3 and GT1b. In the middle pre-hatching age was observed the occurrence of multi-sialylated gangliosides, peaking around the 14th day, and of alkali-labile GP1c (presumably an O-acetylated derivative) in the late post- hatching age in the presence of minute amounts of GM3 and Fuc-GM1 (Sonnino et al., 1990). > Table 6-16 reports data on the changes in the distribution of the most abundant gangliosides of human brain depending on different regions (frontal cortex, cerebellum, caudate nucleus, thalamus and centrum semiovale) and pre-natal (only in the case of frontal cortex) and post-natal age. In the frontal cortex GM1 and GD1a feature a parallel developmental behavior with increasing accumulation picking up around the 8th year, and diminution thereafter; GD1b and GT1b, after a drop of concentration around the 5th–7th month of pre-natal life, continue to accumulate till adulthood; GQ1b concentration, after a drop around birth till the 5th year, increases and reaches the maximal level in the adult life; GD3 has the highest level in the early prenatal life. The most abundant ganglioside is GT1b, followed by GD3 and GD1b in the 3rd month pre-natally; GD1a followed by GM1 and GT1b from birth to the first 8 years; GT1b followed by GD1b and GD1a in the adult age. In the caudate nucleus and thalamus the developmental profiles of gangliosides are similar to those of the frontal cortex, with a preponderance of GD1b over GT1b; in cerebellum the main characteristic is the marked preponderance of GT1b; in centrum semiovale, richer in myelin, the accumulation of all the different gangliosides in post-natal life is much higher than that in the other brain regions, the most abundant gangliosides are GT1b and GD1b, and the proportion of GM1 over the other ganglioside is higher than that in the other brain regions. Noteworthy, evidence has been provided for the presence in human infant brain (age: 2 years) of small (0.5–2 nmol bound Neu5Ac/g fresh tissue) amounts of two gangliosides of the lacto-series, 3’-LM1 and 3’-isoLM1 (Li et al., 1973; Molin et al., 1987). > Figures 6-8 and > 6-9 report the developmental profiles of gangliosides in rabbit brain. In > Figure 6-8, the data are given as % of the individual gangliosides on total gangliosides (as bound Neu5Ac). In the pre-natal age the major gangliosides are GD1a and GT1b, and only at an early stage (21–22 days) GQ1b is the second most abundant one, after GT1b. After birth, in cerebrum the most abundant ganglioside is GD1a followed by GT1b and GM1, whereas in cerebellum GT1b prevails, followed by GD1a and GD1b. In both tissues after birth GD3 is barely detectable. This confirms the notion that the overall outcome of the biosynthetic machinery of gangliosides is different in cerebrum and cerebellum. In -1 > Figure 6-9 the data related to pre-natal age are expressed as nmol bound Neu5Acg fresh tissue and refer separately to membrane-bound and cytosoluble gangliosides. All membrane-bound gangliosides, with the exception of GQ1b, show increasing accumulation during the pre-natal period of life with higher rates at 21–22 days and toward the end of pregnancy. Conversely, cytosoluble gangliosides, after a peak around 21– 22 days of pregnancy, feature a gradual and marked decrease of concentration till birth. On the 21st day of pre-natal life and at birth the total concentration of cytosoluble gangliosides is 52 and 2.5 nmol bound Neu5Acg-1 fresh tissue, respectively, and that of membrane-bound gangliosides 730 and 1,970 nmol bound Neu5Acg-1 fresh tissue, respectively. Thus the ratio between the two ganglioside populations shifts from 1:14 on 21st day of pregnancy to 1:790 at term (Chigorno et al., ). The hypothesis has been proposed that the cytosoluble and membrane-bound gangliosides are metabolically correlated, the cytosoluble forms behaving as precursors of the membrane-bound ones (Chigorno et al., 1984). Likely, the deposition of gangliosides into the membrane of neural cells is the result of at least two processes: the biosynthesis of the individual gangliosides and their transport and insertion into the membranes. As the rate of membrane biosynthesis rapidly increases during pre-natal life it is conceivable that the half-life of cytosoluble

7 – 170 375 140 70 90 35

1/2 110 230 750 190 130 45 42

1 – 270 845 245 305 40 43

5 98 440 1,030 430 420 36 44

8 96 463 1,053 527 423 77 –

2 102 365 1,020 290 380 38 45

5 – 80 190 130 195 120 28

3 – 38 80 140 290 63 90

The most abundant gangliosides in each tissue are given in bold character

Ganglioside GM2 GM1 GD1a GD1b GT1b GQ1b GD3

Frontal cortex Post-natal age (years)

Pre-natal age (months) 44 47 361 681 877 942 171 –

73 45 339 604 675 827 135 –

8 >20 121 459 574 924 101 –

44 >20 164 331 547 874 187 –

73 >20 171 308 546 1,087 91 –

Cerebellum post-natal age years 8 32 470 1,130 499 560 101 –

44 37 335 867 848 876 159 –

73 31 291 886 890 704 136 –

Caudate nucleus post-natal age years 8 25 326 586 737 535 145 –

44 Table 6-17, which illustrates the changes of the ganglioside pattern of rat cerebellar granule cells during differentiation in vitro. The 2nd day in culture corresponds to the initial stage of neuronal differentiation and neuritogenesis; the 8th day in culture corresponds to morphologically and biochemically fully differentiated neurons connected by a complex net of fasciculated fibers; the 17th day in culture corresponds to a late stage of neuronal development with initial signs of age-induced apoptotic death. In general, the ganglioside pattern exhibited by these cells reflects the distribution observed

. Table 6-17 Changes in the ganglioside pattern of rat cerebellar granular cells during differentiation in vitro. The ganglioside content is expressed as pmol/106 cells, and as % on total ganglioside content. The most relevant compositional changes are indicated in bold characters. Quantification of gangliosides was performed by the radiometric method after steady-state metabolic labelling of cells with [1-3H] sphingosine Days in culture Gangliosides Total (%) GM1 (%) GD3 (%) GD1a (%) GD1b (%) O-Ac-GT1b (%) GT1b (%) O-Ac-GQ1b (%) GQ1b (%)

2 pmol 74 3 11 20 9 3 25 1 2

Elaborations of data from Prinetti et al. (2001)

(%) – 4.0 14.9 27.0 12.1 4.0 33.8 1.3 2.6

8 pmol 790 60 40 210 90 80 260 10 20

(%) – 7.6 5.0 26.6 11.4 10.1 32.9 1.3 2.5

17 pmol 1020 50 40 260 100 90 320 10 30

(%) – 4.9 3.9 25.2 9.8 8.8 31.4 1.0 2.9

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in rat cerebellum with GT1b as the major ganglioside (about 31–34%), followed by GD1a (25–27%). All gangliosides undergo a marked accumulation during differentiation, but with relevant differences. The increase of GD1a, GD1b, GT1b, GQ1b, and O-Ac-GD1b (from 10-fold to 15-fold) is in line with the increase of total gangliosides (14-fold). Instead, GD3 features the lowest degree of accumulation (3.7-fold) but the most pronounced decrease of relative percentage (from 14.9 to 3.9%), being the third most represented ganglioside at 2 days in culture, after GT1b and GD1b. This trend closely reflects the developmental behavior of GD3 in rat cerebellum. Notably, GM1 and O-Ac-GT1b undergo the highest accumulation (20-fold and 30-fold, respectively) and double their relative concentrations from the 2nd to the 8th day in culture, that is, in the period required by cells to reach full differentiation. This consolidates the notion that GM1 is not only a marker of myelination that is absent in these cells, but also of neuronal differentiation. Concluding, more research is needed, new experimental approaches should be designed, and, hopefully, novel and challenging work-hypotheses be conceived in order to better understand the links between distinct morpho-functional events occurring in nervous system development and involvement of specific sphingolipids.

3.3.3 Developmental Changes of the Fatty Acid and Long Chain Base Composition of Individual Sphingolipids, Particularly Gangliosides The first accurate investigation concerning the fatty acid compositional changes occurring in a brain sphingolipid during human brain development was performed by Sta¨llberg-Stenhagen and Svennerholm (1965), and was regarding sphingomyelin. This paper is of relevant historical value and it is worth to summarize the most important data herein contained (see > Table 6-18). The modifications observed in the fatty acid pattern, expressed as % value on the total fatty acid content were as follows: (1) C16:0 content decreases from 11.9% at the 29th week of pre-natal life to 2–3% at the older age; (2) C18:0, after a moderate increase along pre-natal life till a maximum in the first year after bith (from 70.5 to 85.6%) gradually decreases with age (73% in the old subjects); (3) C24:0 increases, particularly in the white matter, from 4.6% at 8 years to 8% at the old age; (4) C24:1 increases from 6.9% at the 29th fetal week to 41–42% in the white matter and 12–13% in the gray matter of the adult; (5) C25:1 and C26:1 also increase with age from 0.1/0.2% at birth to 7–8% in the white matter and 2.5–3.2% in the gray matter in the adults. Interestingly,

. Table 6-18 Fatty acid composition of sphingomyelin from normal human frontal lobe at different pre-natal and post-natal ages. Each fatty acid is expressed as % of total fatty acids. At the ages marked with an asterisk the unfractionated frontal lobe was used; in all other cases the frontal lobe was fractionated in gray and white matters. In these cases the figure before the slash refers to gray matter, the one after the slash to white matter. Elaboration from data reported by Sta¨llberg-Stenhagen and Svennerholm (1965) Age

Fetal weeks (months)

Post-natal life (years)

Fatty acid C16:0 C18:0 C20:0 C22:0 C24:0 C24:1

29* 11.9 70.5 2.0 2.7 2.5 6.9

33* 8.7 76.8 2.2 2.1 1.9 6.0

Birth* 7.3 84.8 1.6 1.8 1.2 2.2

8 5.6/2.9 85.5/49.5 1.6/2.2 1.0/2.7 0.9/4.6 3.8/28.3

17 7.1/3.1 85.6/34.9 1.7/1.7 1.0/1.7 0.8/9.7 2.5/30.2

15 2.2/1.2 82.1/25.2 2.5/1.0 0.8/2.0 1.0/7.0 5.1/42.3

33 2.7/2.1 69.1/26.0 2.6/0.9 1.6/1.8 3.0/8.1 12.6/41.2

C25:1 C26:1

0.4 0.6

0.1 0.2

0.1 0.2

0.2/1.5 0.2/3.4

–/3.4 –/6.0

2.0/7.4 0.8/6.0

2.8/6.8 2.0/5.0

77 3.1/2.1 73/23.3 2.6/1.1 1.4/2.0 2.9/8.1 13.4/ 38.8 3.2/7.8 2.5/6.9

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

6

also the ratio of the % contents between gray and white matter changes with age, indicating that there is a differential age effect for sphingomyelin metabolism in the two brain areas. A similar trend – diminution with age of C18:0 and increase of C20:0 and longer-chain fatty acids, and mono-unsaturated fatty acid – was also observed in the different major gangliosides present in the brains of different animals (Ando and Yu, 1984; Mansson et al., 1978; Palestini et al., 1990). The first report on the effect of age on the long chain base composition of gangliosides was given by Rosenberg and Stern (1966). The notion released was the diminuition of C18:1 with concomitant increase of C20:1 long chain bases during brain development and ageing. More detailed studies on this topic regarding individual gangliosides were performed later (Ando and Yu, 1984), particularly when advanced HPLC methods were developed for the separation of molecular species of the same ganglioside carrying different long chain bases. Surprising was the evidence that no ganglioside-linked C20:1 long chain base could be detected in human, rat, mouse, rabbit, cat, dog and chicken brains during pre-natal life (Palestini et al., 1991). Additional details on the behavior during post-natal life of long chain bases contained in the major gangliosides (GM1, GD1b, GT1b, GQ1b) from rat forebrain are presented in > Table 6-19. As shown the % content of C20:1 long chain base in all gangliosides undergo a continuous . Table 6-19 Changes with age of the proportion (as % of total) of the long chain base C20:1 contained in the most abundant gangliosides from rat forebrain. Elaboration of data provided by Palestini et al. (1990). The remainder % content is covered by C18:1 long chain base Ganglioside species C20:1, % on total Age Days 3 15 30 Months 6 24 Increase, fold

GM1

GD1a

GD1b

GT1b

GQ1b

8.1 12.2 25.6

4.2 9.4 21.3

6.0 14.1 30.3

5.3 18.0 35.3

9.0 18.1 32.4

30.3 38.4 4.7

24.1 28.0 6.7

40.2 45.4 7.5

38.2 41.4 7.8

45.5 56.4 6.3

increase from 3rd day after birth to adulthood. The increase is 4.7-fold for GM1, 6.7-fold for GD1a, 7.5-fold for GD1b, 7.8-fold for GT1b and 6.3-fold for GQ1b. In the adult age the highest percentage of C20:1 long chain base is featured by GQ1b (56.4%), followed by GD1b (45.4%), GT1b (41.4%), GM1 (38.4%) and GD1a (28%). Of course, in parallel with the increase of C20:1, there is a parallel decrease of C18:1 long chain base in all gangliosides. A further important issue concerns the fatty acid composition of the ganglioside molecular species carrying C18:1 or C20:1 long chain base (l.c.b.). As shown in > Figure 6-11 the fatty acid compositions of GD1a-C18:1 (l.c.b.) and GD1a-C20:1 (l.c.b.) are different. Both compounds contain as the major fatty acids C16:0, C18:0, C18:1 and C20:0, covering all together more than 95% of the total fatty acid content. In GD1a-C18:1 (l.c.b.), C16:0 and C18:1 fatty acids undergo a constant and relevant increase from 3 days to 24 months of age (from 7 to 14%, and from 3 to 13%, respectively), C18:0 a marked decrease from 88 to 71%, whereas C20:0 remains constant, although at low concentrations. In GD1a-C20:1 (l.c.b.), C16:0 and C18:1 maintain constant levels, with some small fluctuations, C18:0 decreases from 58 to 46%, and C20:0 increases from 13 to 18%. The conclusion is that in the course of the multifaceted process of brain development and ageing the chemical diversification of glycosphingolipids deals not only with the saccharide portion of the molecule, but also with the two constituents of the ceramide portion, fatty acid and long chain base. This provides an extremely diversified molecular array with an extraordinary high potential of interactions.

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. Figure 6-11 Effect of post-natal age on the fatty acid composition of the two different species of ganglioside GD1a carrying C18:1 and C20:1 long chain base, respectively. These ganglioside species were isolated from rat forebrain at different post-natal ages, from 3 days to 24 months. GD1a was chosen as the most abundant ganglioside in rat forebrain. Elaboration of data provided by Palestini et al. (1990)

4

Conclusion

Sphingolipids, particularly glycosphingolipids, are very versatile molecules, because of the wide diversification of their carbohydrate, fatty acid and long chain base components. Owing to these chemical features they can warrant specific interactions with proteins, other lipids, glycoproteins and possibly nucleic acids. Glycosphingolipids-mediated specific interactions can lead to: (1) modulation of receptors, enzymes, channels and carriers at the membrane level; (2) formation of more rigid membrane microdomains like lipid membrane rafts (also named sphingolipid enriched microdomains); (3) establishment of cell–cell adhesion or repulsion; (4) liberation of phosphosphingolipids, sphingoid fragments – sphingosine, ceramide – that directly, or after phosphorylation, behave as bioregulatory messengers. Knowledge of the complex sphingolipid chemistry has guided explorations on the mechanisms of their biosynthesis and biodegradation, and on the sites of regulation of their expression in cells and tissues. A further opportunity of sphingolipid chemistry is the possibility to produce analogs and derivatives of sphingolipids designed in order to recognize part of the sphingolipid molecule responsible for a particular biological effect, and identify the cellular and subcellular location of the individual sphingolipid. Moreover, the chemical features of sphingolipids, particularly glycosphingolipids,

Chemistry, tissue and cellular distribution, and developmental profiles of neural sphingolipids

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constitute a challenge and, concomitantly, a guideline to establish precise connections between individual molecules and physiological events like proliferation, maturation and apoptosis of single neural cell types. The survey on the cellular and tissue distribution of sphingolipids, and on their behavior during neural differentiation, maturation and ageing, shows an enormous amount of data, and clearly indicates some definite connections between individual molecules and morpho-functional events (for instance the association between ganglioside GM4 and GM1 and the process of myelination), besides a number of enigmatic issues. Efforts should be made to develop sub-micromethods for sphingolipid analyses, in vitro systems of cell differentiation in culture and co-culture of different neuronal and glial cells, applications of imaging procedure to in vitro culture systems and new approaches to induce cell differentiation, for instance by the use of small synthetic molecules. The bases for the furtherance of research in this field are solid, and the expectations reasonably optimistic.

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Cellular and Subcellular Localization of Neural Lipids

7

Nuclear Lipids and Their Metabolic and Signaling Properties

R. Ledeen . G. Wu

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175

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Nuclear Structure and Endonuclear Domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175

3 Isolation Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 177 3.1 Purification of Nuclei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 177 3.2 Isolation of Subnuclear Domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 177 4 4.1 4.2 4.3

Lipid Composition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 178 Lipids of Whole Nuclei . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 178 Lipids of the Nuclear Envelope . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 178 Lipids of Endonuclear Domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 179

5 5.1 5.2 5.3

Glycerolipid Metabolism and Signaling in the Nucleus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 180 Choline and Ethanolamine Phosphoglycerides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 180 Inositol Phosphoglycerides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181 Diacylglycerol and Its Kinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183

6

Nuclear Eicosanoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 184

7 7.1 7.2 7.3

Sphingolipid Metabolism and Signaling in the Nucleus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 184 Sphingomyelin and Related Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 184 Ceramide and Related Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 Glycosphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187

8

Concluding Remarks on Nuclear Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 191

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_7, # Springer ScienceþBusiness Media, LLC 2009

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Abstract: Nuclei have a relative paucity of lipids, perhaps accounting for the early assumption that their role in that organelle is limited to providing structural support to the nuclear envelope. In addition to the growing awareness that lipids of this double membrane structure have dynamic signaling properties, biochemists and cell biologists now recognize that lipids also occur in endonuclear compartments where they exert an astounding array of signaling capabilities with profound influence on cellular functioning. Gone also is the concept of the nucleus as passive recipient of lipids that are synthesized elsewhere in the cell and imported, since numerous enzymes have been discovered that synthesize and catabolize lipids within the nucleus. Phosphatidylinositol is synthesized in extranuclear compartments and transferred to the nucleus where it is phosphorylated to phosphatidylinositol bisphosphate, a substrate for phospholipase C. The latter generates the two second messengers, inositol trisphosphate and diacylglycerol, which draws protein kinase C to the nucleus and activates it; the polyunsaturated forms of diacylglycerol are especially active in that regard. Phosphatidylinositol bisphosophate can also react with phospholipase A2 to liberate arachidonic acid, which can in turn undergo conversion to eicosanoids within the nucleus. More saturated forms of diacylglycerol are formed by hydrolysis of phosphatidylcholine, some of which are disaturated in terms of the aliphatic chains. Phosphatidylcholine is synthesized within the nucleus, as evidenced in the presence of the Kennedy pathway enzymes. The diacylglycerol form of signaling is terminated by diacylglycerol kinase, several isoforms of which occur in the nucleus. Sphingolipids such as sphingomyelin also have a prominent role in nuclear signaling, the main metabolic product being ceramide that is generated by sphingomyelinase. Evidence has suggested that the ratio of ceramide to diacylglycerol is a form of regulatory control critical for homeostatic properties of the nucleus. Sphingomyelin comprises a significant component of chromatin lipids and its variation in relation to cholesterol and phosphatidylcholine indicated that nuclear matrix lipids are metabolized independently of chromatin lipids. Current and prior studies suggest several key processes involving RNA and DNA reactivity that are dependent on these lipid-initiated events. Considerable interest has focused on inositides whose activities include promotion of transcription through neutralizing histone-mediated repression. These and other lipids occur in specles, the microdomains that are believed to contain molecules involved in splicing of pre-mRNA. Nuclei from mammalian cells all have the same general structure consisting of the double-membrane envelope and various less well-defined endonuclear compartments. The two membranes that make up the nuclear envelope are quite different in lipid composition and function: the outer membrane is continuous with the endoplasmic reticulum and bears many similarities to the latter, whereas the inner membrane is unique and contains elements that mediate communication between nucleoplasm and the lumen of the nuclear envelope. The latter is a calcium storage site, continuous with that of the endoplasmic reticulum, from which calcium can be released in signaling mode to the nucleoplasm and to which excess calcium can be transferred via a sodium– calcium exchanger in conjunction with GM1 ganglioside. This was shown to exert an important neuroprotective function in neural cells. GM1 and GD1a are the primary species of gangliosides found in the nuclear envelope; those together with GD3 are also present in some endonuclear domains. These ganglioside characteristics of the nucleus were observed in neural cells of various types and in certain nonneural cells as well. List of Abbreviations: CGN, cerebellar granule neurons; CER, ceramide; DAG, diacylglycerol; DGK, diacylglycerol kinase; ER, endoplasmic reticulum; INM, inner nuclear membrane; Ins(1,4,5)P3, inositol trisphosphate; NCX, sodium–calcium exchanger; NE, nuclear envelope; ONM, outer nuclear membrane; PGHS, postaglandin H Synthase; PI3K, PtdIns(4,5)P2 3-kinase; PI-PLC, phosphoinositide specific phospholipase C; PL, phospholipid; PLA2, phospholipase A2; PLase, phospholipase; PLC, phospholipase C; PLD, phospholipase D; PtdCho, phosphatidylcholine; PtdEtn, phosphatidylethanolamine; PtdIns, phosphatidylinositol; PtdIns(4)P, phosphatidylinositol 4-phosphate; PtdIns(4,5)P2, phosphatidylinositol 4,5bisphosphate; PtdOH, phosphatidic acid; PtsSer, phosphatidylserine; S-1-P, sphingosine-1-phosphate; SM, sphingomyelin; SMase, sphingomyelinase. Nomenclature of individual phospholipids is in comformity with the recommendations of the IUPAC-IUB Commission on Nomenclature of Lipids. Ganglioside nomenclature is that of the JCBN recommendations [Eur. J. Biochem. 257: 293–298 (1998)]

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Introduction

Nuclei of all cells that have been examined to date, including those of the nervous system, contain a large variety of lipids that are localized primarily in the nuclear envelope (NE). The relative paucity of lipids in endonuclear compartments likely accounted for the early view that lipid function in this organelle is limited to that of providing structural support for the NE. However, this has changed dramatically in the last decade or two with explosive growth of research on lipids of endonuclear domains. This has demonstrated in significant detail the presence of glycero- and sphingolipids within the nucleus, which, although quantitatively minor, are metabolically quite active. Such endolipids are now recognized as important mediators of a complex array of signaling reactions and modulatory mechanisms that exert major influence on nuclear and cellular functioning. There were early indications of such phenomena in studies of interactions of various lipids with DNA (Manzoli et al., 1974), following which the concept of a distinct nuclear inositol lipid signaling system whose activity depends on the differentiation state of the cells (Cocco et al., 1987) emerged. Phosphatidylcholine (PtdCho) is synthesized and metabolized within the nucleus (Hunt, 2006a), a process not yet established for similar glycerophosphatides such as phosphatidylethanolamine (PtdEtn) and phosphatidylserine (PtdSer). Glycosphingolipids, such as gangliosides have come into the picture as regulators of nucleoplasmic Ca2+ (Xie et al., 2002) with the possibility of additional endonuclear roles. Recent progress in these areas has been sufficiently dramatic to warrant description of the nucleus as ‘‘a cell within a cell’’ (Bkaily et al., 2003). To date there have been few if any systematic comparisons of nuclear lipids from different tissues and species. Similar properties and derived mechanisms appear to apply in broad outline to nuclei of diverse origin, although unique features undoubtedly exist in regard to lipid signaling; these await systematic exploration. In addition to their capacity for independent metabolism, nuclei are also responsive to the stimuli of extranuclear and even extracellular origin. It is noteworthy that certain extracellular stimuli are able to induce phosphoinositide signaling in the nucleus only (Divecha et al., 1993a; Cocco et al., 2001) or in both nucleus and cytoplasm (Maraldi et al., 1994; Kleuser et al., 2001). Phospholipids (PLs), the predominant lipids of the NE and endonuclear compartments, were the principal focus in early studies and remain so today, although these now share attention with such signaling entities as ceramide (CER), diacylglycerol (DAG), gangliosides, and sphingosine phosphate. Each of the two membranes that comprise the NE is known to possess unique composition and metabolic/signaling patterns, including regulation of Ca2+ flux and other determinants of nuclear homeostasis. In this chapter, we will attempt to summarize some of the major findings in the area of nuclear lipid composition and function, while highlighting neural cell nuclei where such data are available. For additional details the reader is referred to a number of informative reviews that have appeared in recent years (Martelli et al., 2001; Tamiya-Koizumi, 2002; Irvine, 2003, 2006; Albi and Viola Magni, 2004; Hunt, 2006a, b; Ledeen and Wu, 2006a, b, 2007).

2

Nuclear Structure and Endonuclear Domains

The NE appears to contain the only well-defined membrane(s) of the nucleus, since transmission electron microscopy failed to reveal endonuclear membranous systems. Despite this apparent absence of internal membranes, nuclei of eukaryotic cells are viewed as structurally well ordered with discrete subnuclear domains throughout the nucleoplasm (> Figure 7-1) (Lamond and Earnshaw, 1998). However, these endonuclear compartments have also been described as functionally diffuse and dynamically variable in relation to metabolic function (Maraldi et al., 1998) in contrast to the relatively stable structure of the NE. One such domain is chromatin itself, the major repository of nucleic acids, which interacts with the nuclear matrix or nucleoskeleton whose main function is to organize chromatin. This matrix is considered analogous to the cellular cytoskeleton in maintaining shape and is operationally defined as the components that remain insoluble after extraction of the nucleus with nonionic detergents

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. Figure 7-1 Representation of nuclear structure with endonuclear domains, as presently conceived. The outer nuclear membrane (ONM) is continuous with the ER, while the inner nuclear membrane (INM) is closely associated with the nuclear lamina and has a unique lipid composition. These two membranes are joined at the nuclear pore complexes that are distributed over the nuclear surface and permit passive flow of small molecules between cytoplasm and nucleoplasm. The lumenal space between the two membranes of the nuclear envelope (NE) is a storage site for Ca2+, continuous with the ER lumen. In addition to the NE, lipids have been shown to occur in intranuclear compartments such as nucleolus, chromatin, heterochromatin, and nuclear matrix whose composition is dependent on isolation methodology (Reproduced from Figure 7-1 of Ledeen and Wu, 2006b with permission.)

and salts and treatment with nuclease. Its composition is consequently dependent on isolation methodology and has been described as including the nuclear lamina, inner matrix, elements of the NE, and various structural links between the internal matrix and peripheral lamina (Maraldi et al., 1998; Vlcek et al., 2001). It thus remains to be resolved whether the nuclear matrix is a distinct structural/functional domain and to what extent the isolated nuclear matrix corresponds to an in vivo existing structure (Martelli et al., 2002). The nuclear lamina comprises a meshwork of intermediate filaments located on the endonuclear surface of the inner nuclear membrane. The so-called heterochromatin regions, which contain relatively little DNA and are transcriptionally inactive, are nevertheless rich in specific nuclear proteins that regulate transcriptional activity. Heterochromatin can suppress the transcriptional activity of genes that are translocated adjacent to it (Lamond and Earnshaw, 1998). The nucleolus contains the ribosome-producing machinery and is one of the better defined structural units. Among the methods that have been described for characterizing lipids and lipid metabolizing enzymes within these nuclear domains are cytochemical, autoradiographic, and biochemical techniques

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(Maraldi et al., 1998). The two membranes of the NE can be isolated in relative purity (see below) and although detailed comparison of their lipid compositions has not been reported, some differences have already been noted. Cholesterol, for example, was readily detectable in the outer nuclear membrane (ONM) but not the inner nuclear membrane (INM) (Alroy et al., 1981; Kim and Okada, 1983), and GM1 ganglioside, which occurs in both membranes, is associated with a Na+/Ca2+ exchanger (NCX) only in the inner membrane (Xie et al., 2002). The outer membrane is continuous with the endoplasmic reticulum (ER) and contains similar lipids as the latter, but at different concentrations (Keenan et al., 1972a; James et al., 1981). The two membranes are joined at the nuclear pores by the pore membranes which are associated with the nuclear pore complexes. The latter are distributed over the entire nuclear surface and consist of multiprotein assemblies of 1000 polypeptides that allow passive transfer of small and middlesized molecules (95%), with resultant high buoyant density, has facilitated isolation of nuclei from both tissues and cultured cells in relatively high purity. Various methods of isolation have been described, most of them employing differential and discontinuous-gradient centrifugation through high-density sucrose media; often two successive such gradients are employed. One of the earliest such methods was that used for liver cell nuclei by Blobel and Potter (1966) in which the tissue was homogenized in isoosmotic medium containing sucrose, KCl, MgCl2, and Tris–Cl (pH 7.4) that served to stabilize the nuclei. The latter were then pelleted by centrifugation through high density sucrose. Procedures of this type have been utilized for nuclei from neurons and neuronal cell lines (Baker and Chang, 1990; Wu et al., 1995; Antony et al., 2000; Saito and Sugiyama, 2002). If exposure of the nuclei to such strongly hyperosmotic medium is undesirable, alternative methods using an isoosmotic discontinuous gradient can be used (Graham, 2001). Purity is confirmed by light or electron microscopy and assay of marker enzymes for potential contaminants, e.g., 50 -nucleotidase or Na/K-ATPase (plasma membrane), a-mannosidase or galactosyltransferase (Golgi apparatus), cytochrome c oxidase (mitochondria), and glucose-6-phosphatase or NADPH-cytochrome c reductase (ER). The low activity often found for the latter enzymes does not necessarily indicate contamination since, as mentioned, the ONM is continuous with the ER and shares many of its properties.

3.2 Isolation of Subnuclear Domains The NE is obtained by treatment of isolated nuclei with DNase or DNase + RNase, followed by 6M NaCl to release DNA fragments. The fact that the INM is intimately associated with the peripheral nuclear lamina often results in portions of the latter co-purifying with the NE (Georgatos and Blobel, 1987). The ONM of the NE can be selectively removed from whole nuclei by treatment with 2% Na-citrate (Gilchrist and Pearce, 1993) or 0.2% Triton X-100 (Gurr et al., 1963; Neitcheva and Peeva, 1995). The INM is then liberated from the resulting nuclei by treatment with DNase/RNase (Gilchrist and Pearce, 1993). Methodology for isolating NE with associated nuclear pore complexes has been described (Matunis, 2006) that is a modification of an

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earlier procedure (Dwyer and Blobel, 1976). It employs two-step extraction of chromatin with low magnesium at pH 8.0 and low concentrations of Heparin; various detergents can then be employed to selectively solubilize the nuclear pore complexes. An important advange of this procedure is that it facilitates analysis of the many proteins (nucleoporins) that comprise the complex. These structures have not yet been subject to lipid analysis. As mentioned earlier there are various problems attendant to defining and isolating the so-called nuclear matrix as a discrete morphological entity (Martelli et al., 2002). Similar questions arise in regard to other proposed subnuclear compartments in the absence of evidence for membranous structures within the endonuclear regions. However, spliceosomes, the regions responsible for mRNA formation, are now receiving attention as a result of improved isolation procedures. These highly conserved aggregates from yeast to mammals contain five small nuclear RNAs and numerous proteins and have been isolated as highly purified 40–60 nm particles with functional splicing activity (Zhou et al., 2002).

4

Lipid Composition

4.1 Lipids of Whole Nuclei Phospholipids comprise the large bulk of lipids in nuclei of neural cells, as for nuclei in general, with lesser amounts of sphingolipids, cholesterol, free fatty acids, and others. These have been described in whole nuclei as well as individual nuclear domains. The total PL content of rat liver nuclei was reported as 3.25% by wt, compared with 74.6% for protein and 22.2% for DNA (Neitcheva and Peeva, 1995). Several studies of liver nuclei have shown PtdCho to be the major PL, with lesser but still significant amounts of phosphatidylethanolamine (PtdEtn) and phosphatidylinositol (PtdIns) (Khandwala and Kasper, 1971; Keenan et al., 1972a; James et al., 1981; Neitcheva and Peeva, 1995). Phosphatidylserine (PtdSer) and sphingomyelin (SM) were detected at lower levels. Present at even lower levels were various lipids prominent in signaling, such as DAG and phosphatidate that increased during cell proliferation (Bocckina et al., 1989; Banfic et al., 1993), and sphingosine that increased during mitosis (Alessenko, 1995) or apoptosis (Alessenko and Krenov, 1999). With regard to glycosphingolipids, the initial studies showed ganglioside presence in whole nuclei of rat liver (Keenan et al., 1972b) and bovine mammary cells (Katoh et al., 1993), the latter study presenting evidence for GM3, GD3, and GT1b. More recently whole nuclei from rat brain were shown to contain GM1, GD1a, GD1b, and GT1b with lesser amounts of GM3 and c-series gangliosides; large nuclei had significantly higher concentrations of the same gangliosides compared with small nuclei (Saito and Sugiyama, 2002).

4.2 Lipids of the Nuclear Envelope As revealed in early studies, the large majority of nuclear lipids occur in the NE (Gurr et al., 1963), whose total lipid content was approximately half that of protein by weight (Keenan et al., 1972a). Phospholipids were reported to comprise 65% of NE lipids whereas cholesterol was 10% (3 that of ER); lesser amounts of other neutral lipids (cholesterol ester, diacylglycerol, triacylglycerol) were also detected (Khandwala and Kasper, 1971). Although a relatively high concentration of free fatty acids (15% of total lipid) was reported in the latter study, it was not clear how much of that resulted from breakdown of PLs during isolation. The PL content (per mg protein) of the NE was reported as 9 that of whole nuclei (Khandwala and Kasper, 1971). Comparison with microsomes revealed NE to have significantly less PL/mg protein and correspondingly more cholesterol (Keenan et al., 1972a). The cholesterol content, however, was significantly below that of plasma membranes. Despite the above-mentioned quantitative differences, the PL profiles (% composition) for NE and microsomes were similar. Analysis of fatty acid composition of individual PLs gave discrepant findings, but one such study that used antioxidants to minimize peroxidation showed the PLs of NE and ER to have similar fatty acid profiles with high levels of polyunsaturated fatty acids (principally 20:4 and 22:6) in the four major phosphoglycerides (James et al., 1981).

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A cytochemical study employing cholera toxin B subunit in conjunction with chemical analysis of isolated nuclei showed GM1, GD1a, and minor amounts of other gangliotetraose gangliosides to be present in the NE of rat central nervous system neurons and Neuro-2a neuroblastoma cells (Wu et al., 1995). GM1 was also detected in the NE of peripheral nervous system neurons and NG108–15 cells (Kozireski-Chuback et al., 1999a), their relative content increasing with the onset of axonogenesis (Kozireski-Chuback et al., 1999b). Ganglioside GM1 at those sites, although an order of magnitude less than PLs, was clearly observable when stained with cholera toxin B subunit linked to horseradish peroxidase (see below, > Figure 7-4). Further study of localization placed GM1 in the INM in association with NCX (see below) (Xie et al., 2002); this proved to be a high-affinity association in that it survived SDS–-PAGE. Similar NCX/GM1 complexes were observed in the NE of glia-derived C6 cells and astrocytes, the latter expressing NCX in both NE and plasma membrane while the former had NCX (in association with GM1) only in the NE (Xie et al., 2004a). Whereas a number of nonneural cells were found to contain the NCX/GM1 complex in the NE, this complex was not detected in Jurkat cells or a subgroup of T cells (Xie et al., 2004b). These studies revealed that GM1 also occurs, along with GD1a, in the ONM, but their function at that locus remains to be determined. The presence of gangliosides in endonuclear compartments has been reported (see below). Inositol-containing lipids within the nucleus have been of special interest due to their participation in multiple signaling reactions (see below). Phosphatidylinosistol was demonstrated in early studies to be a component of the NE (Khandwala and Kasper, 1971; Keenan et al., 1972a; James et al., 1981). Phosphatidylinositol 4,5-bisphosphate [PtdIns(4,5)P2], detected with specific monoclonal antibody, was found in the NE (Tran et al., 1993) as well as the nucleoplasm (Voorhout et al., 1992). The derived D-3 phosphoinositide, PtdIns(3,4,5)P3, was shown to occur at the nuclear surface (Yokogawa et al., 2000) and was transiently elevated in the nuclei of PC12 cells subjected to nerve growth factor stimulation (Neri et al., 1999). Relatively few studies have focused on the separated ONM and INM, perhaps due to the difficulty of obtaining sufficient quantities for chemical analysis. However, it is now clear that they possess very different lipid compositions. One report, based on filipin-sterol interaction, found unequal distribution of complexes that suggested higher cholesterol content in the ONM compared with INM (Alroy et al., 1981). In agreement, another study using similar methodology reported filipin-sterol complexes only in the ONM (Kim and Okada, 1983). The above-mentioned NCX/GM1 complex was shown to occur in the INM, suggesting a mechanism for maintaining Ca2+ homeostasis in the nucleoplasm (see below).

4.3 Lipids of Endonuclear Domains A few early studies indicated that nuclear lipids are not limited to the NE but also occur, albeit in limited amounts, in the intranuclear domains of chromatin (Goureau and Raulin, 1970), nuclear matrix (Cocco et al., 1980), and nucleolus (Cave and Gahan, 1970). Use of gold-conjugated phospholipase (PLase) as a cytochemical tool demonstrated intranuclear PLs in the interchromatin spaces and in the nucleolar domain (Maraldi et al., 1992a). Additional evidence came from a combined histochemical and biochemical study of rat liver nuclei, with methodology that ruled out contamination by the NE, showing total PL content of the chromatin to approximate one tenth that of whole nuclei (Albi et al., 1994). While the same PLs were present with similar fatty acid profiles, their relative concentrations differed. Each phospholipid had a unique fatty acid profile that was generally the same whether the PL origin was chromatin or whole nuclei. However, recent study of the endonucleus of IMR-32 neuroblastoma cells revealed enrichment of PtdCho with a high degree of diacyl/alkylacyl chain saturation (Hunt et al., 2001). This study provided an estimate of the nuclear volume occupancy of such disaturated PtdCho species as 10%, suggesting these lipids may be present as large complex aggregates or even as liquid crystalline phases. Biosynthesis of these species of PtdCho was reported to occur endogenously in the nucleus (see below). Cholesterol and SM were found to occur in similar amounts in rat liver nuclei, suggesting a complex of those lipids with proteins in the chromatin (Albi and Viola Magni, 2002). Phospholipids were found localized near the RNA in decondensed chromatin near the nucleoli and nuclear membranes (Fraschini et al., 1992; Maraldi et al., 1992b). Sequential treatment of isolated nuclei with DNase and RNase showed selective removal of PLs with the latter, PtdSer and SM being most affected; this suggested

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functional linkage as well as co-localization of these (and perhaps other) PLs with RNA (Albi et al., 1996). An earlier cytochemical investigation had also indicated co-localization of nuclear phospholipids with RNA-containing structures (Zini et al., 1989). Further study of RNA-PL interaction suggested that SM might represent a bridge between the two RNA strands of double-stranded RNA, thereby protecting them from RNase action (Micheli et al., 1998). Use of small angle X-ray diffraction has shown the presence of both tightly and loosely bound PLs in close association with DNA (Struchkov et al., 2002). A monoclonal antibody specific for PtdIns(4,5)P2 demonstrated occurrence of phosphoinositides at intranuclear sites, confirming the presence of this key signaling inositide in the inner nuclear matrix of in situ matrix preparations (Mazzotti et al., 1995). This correlated with the presence of both metabolizing and synthesizing enzymes for this PL at intranuclear sites (see below). Spliceosomes, the pre-mRNA processing machinery, appear to possess functionally associated lipids. Immunoprecipitates of PtdIns(4,5)P2 were found to contain intermediates and products of the splicing reaction and this lipid was stably associated with electron-dense particles that resembled interchromatin granule clusters (Osborne et al., 2001). These granule clusters are hypothesized to be sites of assembly or storage of factors required to synthesize pre-mRNAs (Spector, 1996). Monoclonal antibody staining revealed the presence of this lipid in ‘‘nuclear specles,’’ similarly characterized subnuclear domains that contain pre-mRNA processing factors; also present were the kinases that convert PtdIns to PtdIns(4,5)P2 (Boronenkov et al., 1998). Discovery of the y isoform of DAG kinase (DGK) in nuclear specles points to the likely co-occurrence of DAG and the kinase product, phosphatidic acid (Tabellini et al., 2003). The presence of galectin-1 and galectin-3 in spliceosomes has been noted (Davidson et al., 2006), and the fact that these proteins interact with galactose-containing lipids as ligands suggests the possible presence of such molecules. Gangliosides are the only glycosphingolipid with verified presence in the nuclei to date and these were shown to occur in the endonuclear domains at apparently low levels. Immunocytochemical evidence was presented for ganglioside GD3 co-localizing with nuclear chromatin in cultured rat cortical neurons subjected to b-amyloid peptide [25–35]; this occurred before the neurons entered S phase and apoptotic death (Copani et al., 2002). Treatment of HUT-78 lymphoma cells with pro-apoptotic anti-CD95 antibody induced nuclear localization of GD3 that correlated with rapid phosphorylation of histone H1 shortly after induction of apoptosis (Tempera et al., 2008). Endonuclear presence of GM1 was suggested in a study of nuclei from mouse intestinal epithelial cells that showed binding of both cholera toxin and anti-GM1 antibodies in the heterochromatin (Parkinson et al., 1989). Such studies are clearly at an early stage of exploration in relation to ganglioside metabolism and signaling within the nucleus.

5

Glycerolipid Metabolism and Signaling in the Nucleus

5.1 Choline and Ethanolamine Phosphoglycerides The nucleus was originally viewed as an organelle with limited capacity for intrinsic lipid synthesis and therefore dependent on extranuclear processes in conjunction with import mechanisms for its lipid components. The nucleus is now recognized as at least semi-autonomous with respect to lipid metabolism, although some of the relevant enzymes originate in the cytosol and are drawn into the nucleus in the course of physiological activity. One form of activity is the well-known acylation–deacylation cycle affecting primarily the sn-2 position of nuclear PLs, which was found to increase in proliferating fibrosarcoma cells (Neufeld et al., 1985). This cycle, studied in the NE of neural cells, arises from the combined actions of acyltransferase (Baker and Chang, 1981a,b) and phospholipase A2 (PLA2) (Tamiya-Koizumi et al., 1989a; Antony et al., 2001). The latter enzyme hydrolyzes fatty acid ester bonds at the sn-2 position of glycerol in phosphoglycerides, in some cases generating polyunsaturated fatty acids that serve as precursors to eicosanoids (see below). PLA2 activity in LA-N-1 neuroblastoma cells included two Ca2+-independent enzymes, one active toward PtdEtn and the other toward plasmenylethanolamine, the plasmalogen analog (Antony et al., 2001). Both enzymes were strongly stimulated by exposure of the cells to retinoic acid, a neuronal differentiating agent. Some isoforms of PLA2 translocate from cytosol to the NE (Fatima et al., 2003) where enzymes of eicosanoid generation are clustered (Surette and Chilton, 1998). The secretory

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form of PLA2 occurs in the nucleus, as shown in detection of group V sPLA2 in the nuclei of PC12 and U251 astrocytoma cells (Macchioni et al., 2004). A more recent study of brain itself revealed the same isoform of sPLA2 in the nuclei of neuronal and glial cells of rat brain (Nardicchi et al., 2007). The above-mentioned study with LA-N-1 cells (Antony et al., 2000) also suggested the presence of phospholipase C (PLC) and phospholipase D (PLD) activities toward PtdCho and the existence of a nuclear PtdCho cycle. A high level of PLD activity was detected in rat brain neuronal nuclei, which was significantly greater than that detected in nuclei of glia or extraneural cells (Kanfer et al., 1996). Isolated nuclei from LA-N-1 cells carried out synthesis of PtdCho, this activity being enhanced by phorbol ester (Antony et al., 2000). Synthesis of highly saturated forms of chromatin-associated PtdCho was shown to occur in endonuclear compartment(s) of IMR-32 neuroblastoma cells in a manner spatially separate and compositionally distinct from that occurring in whole cells (Hunt et al., 2001, 2002). Membrane-free nuclei from these cells were indicated to contain the enzymes that comprise the three reactions of the CDP-choline (Kennedy) pathway: (a) choline + ATP ! phosphocholine + ADP (b) phosphocholine + CTP ! CDP-choline + PPi (c) CDP-choline + DAG ! PtdCho + CMP The a isoform of CTP:phosphocholine cytidylyltransferase, the principal regulatory enzyme in the above pathway (reaction b), is confined to the nucleus throughout the cell cycle and was shown through temperature-sensitive mutation to be essential for cell survival (DeLong et al., 2000). Precisely how the disaturated forms of PtdCho with their unusual acylation/alkylation pattern produced in this nuclear reaction sequence aid nuclear function and cell survival is not known, although these aspects appear consistent with the tight homeostatic control that is evident (DeLong et al., 2000; Hunt et al., 2001). Some DNA-phospholipid interaction is Ca2+-dependent, suggesting a possible role in transcription modifying signals (Quesada et al., 2002). The neural phenotype, neuroblastoma cells, were found to have the most saturated forms of any endonuclear PtdCho pools examined to date, suggesting protection against oxidative damage in the longer lived cell types (Hunt, 2006b).

5.2 Inositol Phosphoglycerides Identification of DAG, PtdIns, and PtdIns phosphate kinase in the NE provided the first evidence for phosphoinositide signaling in that double membrane (Smith and Wells, 1983). Similar activities were later shown to occur in the endonucleus (Cocco et al., 1987; Divecha et al., 1991). It is now well established that nuclei of neural and other cells have a constitutive phosphoinositide cycle, in which all but the initial member of the cycle are synthesized and metabolized within the nucleus (> Figure 7-2). Phosphatidylinositol is synthesized at extranuclear sites after which it is translocated to the nucleus by means of PtdIns transfer protein alpha (Vann et al., 1997; Hunt et al., 2004). The other kinases that sequentially phosphorylate PtdIns and PtdIns4P (types I and II) have been proposed to occur in the nucleus, though not exclusively. In contrast to other nuclear glycerophospholipids, which occur primarily in the NE, PtdIns(4,5)P2 occurrence is predominantly intranuclear (Vann et al., 1997). The first phosphoinositide-specific PLase to be studied was PI-PLC-b1, this being detected in the nuclei of Swiss 3T3 cells (Martelli et al., 1992), rat liver (Divecha et al., 1993b), and PC12 cells (Mazzoni et al., 1992). This enzyme is phosphorylated by p42/44 MAP kinase and enters the nucleus following IGF-I and other mitogenic signaling at the plasma membrane (Xu et al., 2001). The resulting rise in DAG is proposed to attract PKC-a to the nucleus, which may initiate a negative feedback mechanism. In a prior study four different forms of phosphoinositide-specific PLC were isolated from nuclei of rat ascites hepatoma AH7974 cells, all of which required Ca2+ for activity; these hydrolyzed PtdIns, PtdIns(4)P and PtdIns(4,5)P2 but not PtdCho or PtdEtn (Asano et al., 1994). Among the PLC isoforms detected in the nucleus, PLC-d4 was proposed as specific to that organelle (Liu et al., 1996; Maraldi et al., 1999), although that has been questioned (Lee and Rhee, 1996). A study showing that membrane-deleted rat liver nuclei hydrolyzed inositol PLs as effectively as membrane-containing nuclei suggested an intranuclear locus for at least some, and possibly most, PLC activity (Kuriki et al., 1992). Selective extraction procedures showed PLC to occur

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. Figure 7-2 A map of phosphoinositide metabolism, indicating synthetic and hydrolytic pathways. All reactions shown occur within the nucleus except PtdIns synthesis, which occurs at extranuclear domains. ‘‘Turtle’’ figure at top indicates pneumonic device used for numbering system of inositol

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in the nuclear matrix (Payrastre et al., 1992). The b2 and b3 isoforms of PtdIns-PLC translocate from plasma membrane to the nucleus in differentiating HL-60 cells (Bertagnolo et al., 1997), in contrast to the b1 isoform that is exclusively nuclear (Bahk et al., 1998). The selective extraction approach, when applied to phosphoinositide-specific kinases, revealed PtdIns 4-kinase exclusively in the peripheral nuclear matrix and PtdIns(4)P 5-kinase in the internal matrix (Payrastre et al., 1992). Inositol trisphosphate [Ins(1,4,5)P3] is generated (along with DAG) through the action of PI-PLC on PtdIns(4,5)P2. Inositol trisphosphate can mobilize Ca2+ in the nucleus via Ins(1,4,5)P3 receptors on the INM of the NE (Humbert et al., 1996). At that site Ins(1,4,5)P3 receptors are strategically located to regulate nuclear Ca2+ transport to nucleoplasm from the NE, a storage site for Ca2+ continuous with that of the ER. That Ca2+ mediates several key signaling reactions in the nucleus is well established, although the relative contributions of Ca2+ from the NE versus cytosolic compartments remains somewhat controversial (Hardingham et al., 1998; Bootman et al., 2000). Following release from PtdIns(4,5)P2, Ins(1,4,5)P3 can also be converted via successive kinases to InsP6, which has been proposed to have a role in mRNA transport and transcriptional control (York et al., 1999; Odom et al., 2000). Highly phosphorylated inositols have also been implicated in chromatin remodeling (Shen et al., 2003; Steger et al., 2003). Neuronal nuclei and those of other cell types are known to contain D-3 phosphorylated inositol lipids, such as PtdIns(3,4,5)P3 (Martelli et al., 2001; Irvine, 2003). Unlike the ‘‘canonical’’ inositol PLs, members of this family are not susceptible to PLC but act themselves as second messengers. The kinase that forms the above D-3 derivative from PtdIns(4,5)P2, type III PI3K, has been detected in the nucleus and was shown to translocate there in response to agonists (Mejian et al., 1999). With PC12 cells, for example, stimulation by nerve growth factor caused translocation of PI3K to the nucleus where it reacted with PIKE, a phosphoinositide kinase enhancer (Ye et al., 2000). The latter binds to the regulatory subunit of PI3K with resulting activation of the p110 catalytic subunit. PIKE is expressed in a variety of tissues but most abundantly in brain. PTEN, a 3-phosphatase that acts on PtdIns(3,4,5)P3, has been proposed to be partly nuclear and to have a role in neuronal differentiation (Lachyankar et al., 2000). There is considerable evidence to suggest that specific nuclear proteins can interact with phosphoinositides in a manner that modulates chromatin structure (Jones and Divecha, 2004; Martelli et al., 2005). Several in vitro effects of various phosphoinositides on DNA polymerase and other nuclear proteins have been described (Tamiya-Koizumi, 2002). A number of nuclear proteins contain a PtdIns-binding consensus, which may explain the ability of proteins such as histones, DNA polymerase, RNA polymerase, and various transcription factors to bind to PtdIns(4,5)P2 and certain other lipids (Maraldi et al., 1999). It was suggested that the activities of such enzymes are masked by the bound phospholipid and reactivated by metabolic breakdown of the latter (Tamiya-Koizumi, 2002). As one example, PtdIns(4,5)P2 binding to histone H1 reduces binding of the latter to DNA, thereby canceling the inhibition of RNA polymerase II by histone H1 (Yu et al., 1998). Another mechanism for regulation of chromatin structure was suggested in the discovery that the interaction of the chromatin remodeling complex BAF could be regulated by the level of PtdIns(4,5)P2 in the nucleus (Zhao et al., 1998). As mentioned, PtdIns(4,5)P2 has also been proposed to have a structural and/or regulatory function in RNA splicing.

5.3 Diacylglycerol and Its Kinases Diacylglycerol is a metabolic product of both the inositol- and choline-phosphoglyceride pathways and is generated in the nucleus, as elsewhere, by two general mechanisms: (a) phosphoinositide-specific PLC (see above) and (b) hydrolysis of PtdCho, e.g., a different PLC or sequential action of PLD and phosphatidic acid phosphatase. The former produces a stearoyl-arachidonoyl-rich DAG that could serve as precursor to eicosanoids, while the latter yields DAG with a different (more saturated) fatty acid composition. It has been pointed out that DAG constitutes at least 50 structurally distinct molecular species whose fatty acyl groups can be polyunsaturated, di-unsaturated, mono-unsaturated, or fully saturated (Hodgkin et al., 1998). As a general rule the most potent stimulators of PKC are the polyunsaturated DAGs, such as that produced by PLC hydrolysis of PtdIns(4,5)P2. Various isoforms of this kinase, such as PKCa and PKCbII, are translocated to the nucleus following elevation of nuclear DAG. The more abundant subnuclear pool of

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DAG, resulting from hydrolysis of PtdCho, is predominantly disaturated and mono-unsaturated species (Santos et al., 1999). Diverging DAG pathways were in evidence after differentiation, during cell cycle progression, and upon receptor stimulation of various cell types. Thus, HL-60 cells induced to proliferate with IGF-I gave rise to PtdIns(4,5)P2-mediated DAG in the nucleus and a selective translocation of PKCbII, whereas differentiation toward a granulocyte-like phenotype produced DAG from PtdCho with nuclear translocation of PKCa (Martelli et al., 1999). A primary mechanism for switching off the nuclear DAG signal is conversion to phosphatidic acid by DAG-kinase (DGK), a key regulator of proliferation and other cellular changes (Goto et al., 2006). This activity was shown to be preferentially localized in the internal matrix of the nucleus (Payrastre et al., 1992). Several DGK isoforms are known to occur in the nucleus, one of the first to be discovered being DGKz (Goto and Kondo, 1999). Nuclear localization of this isoform in CNS neurons was demonstrated by immunohistochemistry (Hozumi et al., 2003). As mentioned, DGKy occurs in the above-mentioned speckle domains that contain splicing factors. Stimulation of IIC9 cells with a-thrombin produced a rapid rise in the level of nuclear DAG that was derived from hydrolysis of PtdCho (Jarpe et al., 1994). In terms of regulatory mechanisms, RhoA was found to downregulate DGKy (Houssa et al., 1999) and stimulate nuclear phospholipase D breakdown of PtdCho (Baldassare et al., 1997). These pathways involving DAG and its kinases show considerable variability in relation to the cell cycle (Irvine, 2003). The existence of 10 known isozymes of DGK, including several that occur in the nucleus, suggests a high degree of compartmentalization and molecular interaction with other signaling moieties in that organelle.

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Nuclear Eicosanoids

In addition to the above-indicated reaction of PtdIns(4,5)P2 with PI-PLC to produce polyunsaturated DAG, this and other PLs containing arachidonate at the sn-2 position have the potential of being hydrolyzed by PLA2 to free arachidonate which in turn has signaling potential through conversion to eicosanoids. Arachidonate with its 20 carbons and 4 double bonds arranged in the divinylmethane pattern belongs to the n-6 family of essential fatty acids and can be enzymatically metabolized along the 5-lipoxygenase (5-LO) pathway to produce leukotrienes, or along the cyclooxygenase pathway to generate prostaglandins. Enzymes that catalyze these reactions are found in the nucleus as well as other subcellular compartments (Luo et al., 2006). Such enzymes are able to move into the nucleus in a regulated manner, producing eicosanoids that find their receptor targets within the nucleus. Notable examples are the peroxisomal proliferator-activated receptors (PPARs), one of which (PPARd) proved to be a key molecule of prostaglandin I2 signaling (Fukumoto et al., 2005). PPAR a, b, and g isoforms are now recognized as members of the nuclear receptor superfamily of transcription factors that includes the steroid receptors and are considered capable of targeting the nucleus directly (Vamecq and Latruffe, 1999). Synthesis of prostaglandins from arachidonate is initiated by the cyclooxygenase (COX) enzymes, and it is the COX-2 isozyme that is able to process arachidonate to PGH2 within the nucleus. The enzyme 5-lipoxygenease, initiator of leukotriene synthesis, showed diverse localization which, depending on cell type, included cytoplasm, peri-nuclear membranes, and perhaps endonuclear domains. Its import to the nucleus is facilitated by three nuclear localization sequences, each of which can be regulated independently (Jones et al., 2003). Eicosanoids of neural cell nuclei remain a relatively unexplored area.

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Sphingolipid Metabolism and Signaling in the Nucleus

7.1 Sphingomyelin and Related Enzymes Sphingomyelin was recognized early on as a component of liver nuclear membranes (Keenan et al., 1972a; James et al., 1981) and those findings were later extended with demonstration of SM in the nuclear matrix (Neitcheva and Peeva, 1995) and chromatin (Albi et al., 1994). Whereas the latter study showed SM to comprise a significant part of chromatin phospholipids, its concentration at that site was approximately a

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third that in the nuclear matrix; the SM to cholesterol ratio was similar in the two compartments (Albi et al., 2003a). Both SM and cholesterol were found to increase at the beginning of S-phase during liver regeneration, during which time PtdCho decreased. That study demonstrated that nuclear matrix lipids are metabolized independently of chromatin lipids, and also suggested that the higher cholesterolsphingomyelin/phosphatidylcholine ratio in the matrix creates a less fluid environment in relation to DNA synthesis. Also thought to contribute to reduced fluidity in endonuclear domain(s) was the observed enrichment of saturated fatty-acid-containing PtdCho (Hunt et al., 2001). Sphingomyelinase (SMase), the primary enzyme that metabolizes SM, was first detected in the nuclear matrix of rat ascites hepatoma AH 7974 cells (Tamiya-Koizumi et al., 1989b), and subsequently in the NE (Alessenko and Chatterjee, 1995), chromatin (Albi and Viola Magni, 1997), and nuclear matrix (Neitcheva and Peeva, 1995; Albi and Viola Magni, 1997) of rat liver nuclei. The NE was described as the primary site of this enzyme activity in intact nuclei, with translocation to the nuclear matrix in regenerating/proliferating rat liver (Alessenko and Chatterjee, 1995). The NE and nuclear matrix enzymes were proposed to represent different isoforms since neutral SMase 1 was identified biochemically and immunocytochemically as unique to the nuclear matrix and absent from the NE, chromatin, and plasma membrane (Mizutani et al., 2001). The latter report indicated neutral SMase 1 possesses a nuclear export signal but no nuclear localization signal. If as claimed SMase is translocated from NE to nuclear matrix during DNA synthesis (Alessenko and Chatterjee, 1995), this could represent an isoform other than SMase 1. The metabolically significant product of SMase is CER, which can undergo further reactions in the nucleus (> Figure 7-3). Sphingomyelin synthase was detected in both chromatin and NE, the latter activity being significantly greater; the two enzymes showed distinctive properties in regard to Km and pH optimum (Albi and Viola Magni, 1999). An enzyme that carries out the reverse reaction of SM-synthase was recently described in rat liver chromatin through reaction of [14C]SM with DAG, resulting in transfer of [14C]phosphocholine from SM to DAG with formation of phosphatidylcholine (Albi et al., 2003b). Sphingomyelin synthase activity in chromatin was 7.5 that of reverse SM-synthase, and it was unclear whether the two reactions are catalyzed by the same or different enzymes. The reverse reaction thus elevates CER, as does SMase, but with the important difference that it also reduces DAG (while increasing PtdCho). As a result the CER/DAG ratio, viewed as a form of regulatory control, is somewhat higher in chromatin than NE. It was proposed that perturbation of CER-DAG equilibrium in the nucleus may be a key factor that initiates proliferation or apoptosis, depending on the direction and magnitude of the ratio change (2004). Sphingomyelinase and SM-synthase in their various isoforms and loci, together with reverse SMsynthase, are thus considered to function as autonomous regulators of SM-induced nuclear signaling in response to the metabolic requirements of the nucleus (> Figure 7-3). The activity of neutral SMase in ligated rat liver nuclei increased before onset of apoptosis, coincident with increase of CER, and this was followed by elevation of ceramidase and sphingosine in the nucleus (Tsugane et al., 1999); this was thought to reflect NE activity, and no changes in these factors were observed in the plasma membrane. Different results were obtained with chromatin of liver cell nuclei from rats subject to ciprofibrate, an agent promoting hepatocyte proliferation; this resulted in SMase increase in contrast to SM-synthase that was depressed, these changes occurring selectively in the chromatin (Albi et al., 2003c). Following drug withdrawal the same hepatocytes underwent apoptosis with resulting increase in chromatin SM-synthase and SM. Experiments with whole nuclei of an embryonic hippocampal cell line subjected to serum deprivation-induced apoptosis showed that as these cells entered the G1 phase, nuclear SMase was activated and SM-synthase inhibited along with ceramide increase and SM reduction (Albi et al., 2005). These changes likely reflected the more active enzymes of the NE, perhaps behaving in opposite manner to those of chromatin (Albi et al., 2003c). Nuclear SMase activation, presumably in the NE, was shown to have a role in radiation-induced apoptosis of radio-sensitive TF-1 cells (Jaffrezou et al., 2001). Association of nuclear PLs with RNA-containing structures was suggested in a series of cytochemical, biochemical, and ultrastructural investigations that indicated PL localization near RNA in decondensed chromatin (Zini et al., 1989; Fraschini et al., 1992; Maraldi et al., 1992a). Cholesterol and SM were found to occur at equivalent levels, suggesting a complex of those lipids with proteins in the chromatin (Albi and Viola Magni, 2002). In addition to their common localization within the nucleus, study of RNA-PL interaction suggested to the investigators that SM might represent a bridge between the two RNA strands

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. Figure 7-3 Sphingomyelin (SM), the major sphingolipid of nuclei, and its metabolic pathways in mammalian nuclei. All indicated reactions have been shown to occur in the nucleus, with the exception of pathways with dashed arrows, viz., ceramide kinase. In addition, possible catabolism of sphingosine-1-phosphate (S-1-P) via S-1-P phosphatase and/or S-1-P lyase, known to occur in other subcellular compartments, has not yet been detected in the nucleus. A single fatty acid component (C16) of ceramide is shown, but other chain lengths are possible. Abbreviations: PC, phosphatidylcholine; DAG, diacylglycerol; SM, sphingomyelin; SMase, sphingomyelinase (Reproduced from > Figure 7-2 of Ledeen and Wu, 2006b, with permission.)

of double-stranded RNA, providing protection from RNase action (Micheli et al., 1998). The complex was found to contain SMase which, upon activation, rendered the associated RNA sensitive to RNase. Possibly related to these phenomena as well as DNA replication was the finding that PLs detectable in the nucleus underwent significant concentration and reduction in all steps of the S phase, consistent with their conversion to signaling metabolites (Maraldi et al., 1993).

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7.2 Ceramide and Related Enzymes The primary enzymatic product of SMase is CER, which has important signaling functions in the nucleus and which can be converted to other signaling entities by the action of ceramidase, sphingosine kinase, and possibly ceramide kinase. As mentioned, the nuclear levels of CER and DAG are interrelated and their ratio is thought to be an important determinant of nuclear signaling. A form of crosstalk has been reported in chromatin involving PtdCho and SM metabolism which regulates the intranuclear CER/DAG pool (Albi et al., 2008). This kind of nuclear signaling may prove analogous to cytosolic signaling, wherein protein kinase C is activated by DAG and inhibited by CER and sphingosine (Mathias and Kolesnick, 1993; Spiegel et al., 1996). Specificity was indicated in that induction of apoptosis in rat liver led to activation of both SMase and ceramidase with concomitant increases of CER and sphingosine in the nucleus, changes that were not observed in the plasma membrane and were presumably localized in the NE (Tsugane et al., 1999). The role of CER as inducer of apoptosis, although controversial in some respects, has been described most fully in whole cell studies (Mathias and Kolesnick, 1993; Spiegel et al., 1996). Following the initial description of ceramidase activity in the nuclei (Tsugane et al., 1999), a more detailed study of this enzyme in the nuclear membrane of liver was reported including its possession of maximum activity over a broad neutral to alkaline range (Shiraishi et al., 2003). Further catabolism by ceramidase gives rise to sphingosine, which was shown to have modulatory properties in whole cell studies and to be phosphorylated to sphingosine-1-phosphate (S-1-P) (Spiegel et al., 1996). The latter product has become a substance of intense study due to its ability to act as both intracellular messenger and extracellular ligand for a family of G protein-coupled receptors (Le Stunff et al., 2004). It has been implicated as regulator of both cell proliferation and antiapoptotic processes. Of the two major kinases that lead to its synthesis, sphingosine kinase-2 was shown to be localized in the nucleus due to a nuclear localization signal at the N-terminus (Igarashi et al., 2003). Expression of this kinase in various cell types caused cell cycle arrest at the G1/S phase with resultant inhibition of DNA synthesis. On the other hand, Swiss 3T3 cells, when stimulated with platelet-derived growth factor, showed significant increase in the nucleoplasm-associated kinase leading to S-1-P formation that correlated with progression of cells to the S-phase and translocation of the kinase to the NE (Kleuser et al., 2001). That study also revealed S-1-P activity in cytosol to be simultaneously activated and translocated to nucleoplasm following long-term exposure to platelet-derived growth factor. Additional studies will be needed to determine the possible nuclear presence of hydrolase and lyase enzymes of the type that metabolize S-1-P at other loci. The same may be said of ceramide kinase and its product, ceramide-1-phosphate, which in the context of whole cell activity shows evidence of Ca2+ regulatory properties (Colina et al., 2005; Mitsutake and Igarashi, 2005). To our knowledge this has not yet been observed in the nucleus.

7.3 Glycosphingolipids Gangliosides are the only type of glycosphingolipid to have been identified with certainty in the nucleus to date. As outlined above, these glycolipids were readily detected in whole nuclei of primary neurons (Saito and Sugiyama, 2002) and the NE of neuroblastoma cells and primary neurons by a combination of cytochemistry and thin-layer chromatography (Wu et al., 1995). GM1 within the NE was barely detectable before differentiation, and then underwent strong upregulation concurrent with axonogenesis (> Figure 7-4). GM1 and its disialo-analoge, GD1a, were detected in both membranes of the NE, while the INM proved unique in containing GM1 in association with NCX. This association between GM1 and NCX is unusually tight and was found essential for promoting full activity of the exchanger (Xie et al., 2002). A possible function of GD1a in the inner membrane is that of GM1 precursor, undergoing conversion to the latter as needed by a sialidase present in the NE (Saito et al., 1996). Studies with 45Ca2+ showed Ca2+ transfer from nucleoplasm to the NE lumen, consistent with NCX/GM1 location at the inner membrane (Xie et al., 2002). Such activity was relatively limited in the absence of GM1 and could be

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. Figure 7-4 Cytochemical evidence for co-expression of GM1 and Na+/Ca2+ exchanger (NCX) in the NE of cultured neuronal cells. GM1 was detected with Ctx B-HRP, and NCX with anti-NCX antibody plus HRP-linked second antibody. All images are reproduced from original articles with permission of indicated publisher. GM1 expression in the NE of differentiated Neuro2a cells (a) and rat cerebellar granular neurons (b). GM1 expression in the NE of rat superior cervical ganglion neurons (c). GM1 expression in the NE of differentiated (d) and undifferentiated (e) NG108–15 cells, showing scant GM1 in NE of the latter and elevated GM1 in NE of the former. Expression of NCX in the NE of differentiated NG108–15 cells (f). Arrow heads indicate staining of nuclear envelope and arrows staining of plasma membrane (Reproduced from > Figure 7-3 of Ledeen and Wu, 2006b with permission.)

blocked by binding of the latter with cholera toxin B subunit. Ganglioside-induced potentiation of NCX activity was specific for GM1. In seeking an explanation for the high affinity association between nuclear GM1 and NCX (a phenomenon not shared by such molelcules in the plasma membrane), attention has been directed to topology within the membrane. As with plasma membrane NCX, uphill transfer of Ca2+ from regions of low to high concentration is driven by an Na+ gradient, the required intraluminal Na+ buildup occurring naturally by means of an Na+/K+ ATPase in the NE (Garner, 2002). This type of exchange, intrinsically reversible, mediates counter-transport across the plasma membrane of three Na+ ions for extrusion of one cytosolic Ca2+ (Philipson and Nicoll, 2000). A topological requirement of this so-called ‘‘forward mode’’

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is that the large polypeptide loop between transmembrane segments five and six of NCX reside on the low Ca2+ (cytosolic) side, which would place it on the plasma membrane side opposite to the GM1 oligosaccharide chain. Assuming the same loop requirement for NCX in the NE, this would extend into the nucleoplasm to facilitate transfer of nucleoplasmic Ca2+ across the inner nuclear membrane to the high Ca2+ concentration pool within the NE lumen. This would result in the GM1 oligosaccharide chain now being on the same (nucleoplasmic) side as the NCX loop (> Figure 7-5), a different orientation than in the plasma membrane.

. Figure 7-5 Proposed topology of GM1 and Na+/Ca2+ exchanger (NCX) in the inner nuclear envelope (INM, a) and plasma membrane (PM, b). In both cases the large loop between transmembrane units 5 and 6 is located on the low Ca2+ side, i.e., cytoplasm for PM and nucleoplasm for NE. This accords with demonstrated location of both GM1 and NCX in the inner membrane of the NE (INM) and occurrence of the large NCX loop in proximity to GM1 oligosaccharide chain. We have proposed that the high affinity association of GM1 with NCX arises from the negative charge of N-acetylneuraminic acid in GM1 interacting with the alternative splice region (ASR) of the NCX loop, some of whose isoforms are enriched in positively charged amino acids. Such association is not possible for the PM, since the NCX loop and GM1 oligosaccharide occur on opposite sides of the membrane. ONM, outer nuclear membrane (Reproduced from > Figure 7-4 of Ledeen and Wu, 2006b.)

In this configuration the negatively charged oligosaccharide of GM1 is able to interact with the large inner loop of NCX, some of whose isoforms are enriched in basic amino acids (Kofuji et al., 1994). The existence of splice variants of the NCX1 subtype, which predominate in many neural cells (He et al., 1998; Thurneysen et al., 2002), suggests the possibility that specific isoforms are targeted to the NE and others to the plasma membrane. There is evidence to suggest that this GM1-potentiated exchange process has a vital role in maintaining Ca2+ homeostasis in the nucleus. Regulation of nuclear Ca2+ is critically important in relation to cell viability and Ca2+-triggered signaling processes that govern virtually every aspect of cell behavior. Despite the existence of nuclear pore complexes that permit free diffusion of Ca2+ (and small molecules in general) between cytosol and nucleoplasm, some studies have suggested the existence of nuclear-cytoplasmic Ca2+ gradients along with independent regulation of nuclear Ca2+ (Al-Mohanna et al., 1994; Badminton et al., 1998); however, these questions remain controversial for the present (Gerasimenko and Gerasimenko, 2004). Na+/Ca2+ exchangers potentiated by GM1 in the NE could serve a cytoprotective role in shielding the nucleus against prolonged elevation of cytosolic Ca2+, a condition in which Ca2+ exit through nuclear pores would not likely succeed as a protective strategy. The NE lumen is continuous with the ER intermembrane space and thus resembles the latter as a Ca2+ storage site. The outer membrane of the NE contains

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SERCA-type Ca2+-activated ATPase, similar to that in the ER, which pumps cytosolic Ca2+ into the NE lumen (Gerasimenko et al., 1995), whereas the inner membrane contains a number of Ca2+-release mechanisms regulated by Ins(1,4,5)P3, cADP-ribose, and nicotinic acid adenine dinucleotide phosphate (Stehno-Bittel et al., 1995; Humbert et al., 1996; Gerasimenko et al., 2003). Calcium is well known to have a critical role in apoptosis, the nucleus being especially vulnerable to prolonged elevation of nucleoplasmic Ca2+ (Mattson and Chan, 2003). The protective role of gangliosides was suggested in studies of mice engineered to lack GM2/GD2 synthase, resulting in the absence of GM1 and other gangliotetraose gangliosides as well as GM2 and GD2 (Liu et al., 1999). Cultured cerebellar granule neurons (CGN) from such mice were shown to have lost the ability possessed by wild type CGN to regulate Ca2+ homeostasis, resulting in apoptotic death when the cells were exposed to high K+ (Wu et al., 2001). That this was due to the absence of GM1 was suggested in the fact that the mutant cells could be rescued from apoptosis-inducing levels of K+ and glutamate by bath application of this ganglioside; significantly, LIGA-20, a semisynthetic analog of GM1 that is membrane permeant, proved even more effective than GM1 itself (Wu et al., 2004). This correlated with the known efficacy of LIGA-20 to restore Ca2+ homeostasis in normal CGN (Manev et al., 1990) and in the mutant CGN as determined by fura-2 ratiometric determination of intracellular Ca2+ (Wu et al., 2004). In vivo studies pointed to nuclear involvement since the above ganglioside-deficient knockout mouse, when administered kainic acid, developed temporal lobe seizures of significantly greater severity and duration than did normal mice (Wu et al., 2005). Kainate-induced seizures are associated with Ca2+ dysregulation (Ben-Ari and Cossart, 2000), and LIGA-20 again proved significantly more effective than GM1 in attenuating such seizures; experimental results suggested this was due to its greater membrane permeant properties with enhanced ability to cross the blood–brain barrier, enter brain cells, and insert into the NE (Wu et al., 2005). There it was seen to activate the subnormally active NCX of the nucleus and serve as functional replacement for the missing nuclear GM1 in the mutant cells. LIGA-20 also reversed the kainite-induced apoptosis observed in the CA3 region of the hippocampus. The fact that exogenous gangliosides also exert multimodal neurotrophic effects at the plasma membrane (Mocchetti, 2005) suggests the benefits incurred by LIGA-20 in this model may not be limited to the nucleus. In addition to understanding the cytoprotective benefits to cells possessing the nuclear NCX–GM1 complex, it is worth considering the potential benefits that may accrue with the absence of this mechanism in certain normal cells. During development of the nervous system, for example, the importance of programmed cell death as a universal feature of embryonic and postnatal neuroproliferative regions has been well established (Blaschke et al., 1998; Yeo and Gautier, 2004), and the absence of nuclear GM1 at these early stages before neuronal differentiation (Kozireski-Chuback et al., 1999a) may be a factor rendering such cells vulnerable to the necessary apoptosis. This might also pertain to the subpopulation of lymphocytes lacking the NCX/GM1 complex in the NE, analogous to Jurkat T cells (Xie et al., 2004b). Calcium signaling in T cells is recognized as highly complex, Ca2+ entry in such cells being long lasting and necessary for T-cell function (Weiss et al., 1984; Lewis, 2001). A current question is the mechanism(s) by which immune effector cells disappear after eliminating foreign antigens, one proposal being that return of the immune system to rest is mainly due to programmed cell death of activated lymphocytes (Parijs and Abbas, 1998). It remains to be determined whether such lymphocytes are among those shown to lack nuclear NCX/ GM1 (Xie et al., 2004b). To further speculate, the absence of this complex in the NE might also be a factor in maintaining unresponsiveness, or tolerance to self-antigens. As mentioned, the majority of work on nuclear glycosphingolipids has dealt with gangliosides, and there appear to have been relatively few studies of other groups such as neutral glycosphingolipids in this organelle. However, indirect evidence has suggested the possible presence of globotriaosyl ceramide in the NE of human astrocytoma and ovarian carcinoma cell lines, functioning as receptor for the B subunit of Verotoxin/Shigatoxin (Arab and Lingwood, 1998; Lingwood et al., 1998). This is based on the observation that intracellular targeting of the toxin following endocytosis was directed to ER and the perinuclear region, thus defining a potential new retrograde transport pathway from cell surface to nucleus. Interestingly, this targeting appeared dependent on fatty acid composition of the ceramide unit. A systematic exploration of the neutral glycosphingolipids as well as other types of acidic glycolipids in the nucleus would appear warranted at this juncture.

Nuclear lipids and their metabolic and signaling properties

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Concluding Remarks on Nuclear Lipids

The presence of lipids as major participants in nuclear processes has been well demonstrated in a variety of neural and nonneural cells. This applies as much to endonuclear compartments, where lipid levels are relatively low, as to the NE where they are more prominent and provide important structural and metabolic support to the nucleus. A key role for phosphoinositides, especially PtdIns(4,5)P2, has become evident in relation to its ability within the nucleus to give rise to (a) second messengers that regulate PKC and Ca2+ flux, (b) D-3 phosphorylated inositol lipids, and (c) eicosanoids following PI-PLC-induced release of arachidonate. It is significant that the nucleus is semi-autonomous in regard to these biosynthetic and catabolic reactions, as is also true for nuclear PtdCho for which all steps of the Kennedy pathway are present. The high proportion of disaturated species that characterizes the latter phospholipid requires further elucidation as to functional significance. Future studies will likely expand this general area to focus on such phospholipids as PtdEtn and PtdSer, about which relatively little is known beyond the fact of their presence. Results suggesting interaction of phosphoinositides with such nuclear proteins as DNA polymerase, histones, and the spliceosome complex are suggestive of interesting functional roles, which are likely to receive priority in future work. Much remains to be learned about glycosphingolipids such as gangliosides, whose role as potentiator of NCX in the INM has been described but about which relatively little is known in relation to the ONM and endonuclear processes. A more general question for future studies concerns specificity, i.e., which signaling/regulatory processes mediated by nuclear lipids in neural cells, for example, are unique to such cells or more broadly representative. A firm basis has been laid for further experimentation on the highly complex role of lipids in this organelle that has been appropriately termed ‘‘a cell within a cell.’’

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mitogenic action of insulin-like growth factor I. Mol Cell Biol 21: 2981-2990. Ye K, Hurt KJ, Wu FY, Fang M, Luo HR, et al. 2000. Pike. A nuclear gtpase that enhances PI3kinase activity and is regulated by protein 4.1N. Cell 103: 919-930. Yeo W, Gautier J. 2004. Early neural cell death: Dying to become neurons. Dev Biol 274: 233-244. Yokogawa T, Nagata S, Nishio Y, Tsutsumi T, Ihara S, et al. 2000. Evidence that 30 -phosphorylated polyphosphoinositides are generated at the nuclear surface: Use of immunostaining technique with monoclonal antibodies specific for PI 3,4-P(2). FEBS Lett 473: 222-226. York JD, Odom AR, Murphy R, Ives EB, Wente SR. 1999. A phospholipase C-dependent inositol polyphosphate kinase pathway required for efficient messenger RNA export. Science 295: 96-100. Yu H, Fukami K, Watanabe Y, Ozaki C, Takenawa T. 1998. Phosphatidylinositol 4,5-bisphosphate reverses the inhibition of RNA transcription caused by histone H1. Eur J Biochem 251: 281-287. Zhao K, Wang W, Rando OJ, Xue Y, Swiderek K, et al. 1998. Rapid and phosphatidylinositol-dependent binding of the SWI-SNF-like BAF complex to chromatin after T lymphocyte receptor signaling. Cell 95: 625-636. Zhou Z, Sim J, Griffith J, Reed R. 2002. Purification and electron microscopic visualization of functional human spliceosomes. Proc Natl Acad Sci USA 99: 122203-12207. Zini N, Maraldi NM, Martelli AM, Antonucci A, Santi P, et al. 1989. Phospholipase C digestion induces the removal of nuclear RNA: A cytochemical quantitative study. Histochem J 21: 491-500.

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Lipids of Brain Mitochondria

L. Corazzi . R. Roberti

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 200

2 2.1 2.2 2.3 2.4 2.5

Biochemical and Functional Characterization of Purified Brain Mitochondria . . . . . . . . . . . . . . . 202 Lipid Composition of Brain Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 202 Cytochrome c–Cardiolipin Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205 Biosynthetic Origin of Mitochondrial Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 207 Import of Fatty Acids into Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209 Cholesterol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210

3 3.1 3.2 3.3

Physiopathology of Mitochondrial Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 Aging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211 Altered Lipids in Neurodegeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 213

4

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_8, # Springer ScienceþBusiness Media, LLC 2009

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Lipids of brain mitochondria

Abstract: Mitochondrial inner membrane is a dynamic structure that changes shape rapidly in response to variations of osmotic or metabolic conditions. The intrinsic curvature of its constituent monolayers contributes to flexibility, allowing the conversion from flat structures to inverted hexagonal phases. The anchorage of cytochrome c to the inner mitochondrial membrane is mainly due to the interaction with the peculiar mitochondrial lipid cardiolipin. Any cellular event perturbing the stationary state of the lipid may influence the stability of the anchored protein, thus initiating its release outside mitochondria and caspase activation. We describe biochemical and functional characterization of brain mitochondria, focusing mainly on lipid classes and fatty acid composition. The role of cardiolipin fatty acid composition on its interaction with cytochrome c is discussed. In addition, since mitochondria are not able to synthesize all the lipids they contain, with the exception of cardiolipin, processes of lipid translocation from the site of synthesis to the acceptor membranes are described. Emphasis has been given to cholesterol synthesis in brain and to the mitochondrial importation of this lipid in neuronal cells. Finally, some aspects of physiopathology of mitochondrial lipids in aging, ischemia, and neurodegeneration are reviewed. List of Abbreviations: AD, Alzheimer’s disease; CCCP, carbonyl cyanide 3-chlorophenylhydrazone; CL, cardiolipin; CNS, central nervous system; PI, phosphatidylinositol; CPT-1, carnitine palmitoyl transferase; Cyt c, cytochrome c; DΨm, mitochondrial membrane potential; EM, electron microscopic tomography; ETC, electron transfer chain; MAM, mitochondria-associated membranes; mPTP, mitochondrial permeability transition pore; mtDNA, mitochondrial DNA; NAO; 10-N-nonyl-3,6-bis(dimethylamino) acridine; nDNA, nuclear DNA; PAF, platelet-activating factor; PBR, peripheral-type benzodiazepine receptor; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PS, phosphatidylserine; ROS, reactive oxygen species; TIM, translocation of the inner membrane; TOM, translocation of the outer membrane

1

Introduction

A considerable amount of knowledge has been accumulating on the internal structure of mitochondria since the pioneering work of Palade and Sjo¨strand (Palade, 1952; Sjo¨strand, 1953). Structural and functional models of the inner mitochondrial membrane are evolved from the baffle model of a continued closed surface with cristae and connection with the outer membrane through the ‘‘contact sites’’ (Hackenbrock, 1966) to a model characterized by tubular structures, called pediculi cristae, connecting cristae to the intermembrane space (Daems and Wisse, 1966). Application of electron microscopic (EM) tomography to mitochondria in situ in different tissues revealed common features (Perkins and Frey, 2000). 3D imaging EM tomography reveals the cristae as swollen cisterns with narrow, tubular connections to the peripheral surface of the inner membrane (Perkins et al., 2001). The formation of tubular cristae is a dynamic process sensitive to the volume of mitochondrial matrix and to the energetics of protein– lipid membrane folding (Perkins et al., 2001). Mitochondrial structure provided by 3D images of EM tomography suggests a restriction in the diffusion processes between internal compartments with deep functional implications (Westerhoff, 1989). The number and shape of cristae junctions could regulate the diffusion of ions and substrates toward sites of transport or reaction. The close apposition of outer and inner boundary membranes generates nonbilayer lipid structures, the so‐called contact sites, that represent macromolecular assemblies for transport of proteins, ions, or metabolites across outer and inner membranes (Hackenbrock, 1966; Van Venetie and Verkleij, 1982). EM tomography also established the presence of mitochondrial clusters in the vicinity of multilayered ER membranes (Mannella et al., 1998; Mannella, 2000). Small vesicles, mitochondria‐associated membranes (MAM) firmly attached to the mitochondrial surface and possessing all the biochemical features of ER pieces, were purified (Vance, 1990; Camici and Corazzi, 1995). MAM bridge the narrow gap between the outer membrane and the ER. Cytochrome c (cyt c) is a component of the mitochondrial electron transfer chain (ETC) that can initiate caspase activation when released outside the mitochondria during apoptosis (Wang, 2001). Many factors are able to trigger the release of the protein, with the common feature of weakening protein–membrane interactions. Anchorage of the protein is mainly due to cardiolipin (CL) and any cellular event perturbing the stationary state of the lipid may influence the stability and the amount of

Lipids of brain mitochondria

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anchored protein (Ostrander et al., 2001; Piccotti et al., 2002). According to a recent report (Ott et al., 2002), freshly prepared mitochondria contain a small amount of free cyt c that can be modulated by the membrane potential and cyt c–CL interactions (Piccotti et al., 2004). This finding is in agreement with the observation that the shape and volume of the cristae can be expected to affect the diffusion of cyt c between intermembrane compartments as well as the fraction of cyt c bound to the inner membrane (Frey and Mannella, 2000). The inner membrane is a dynamic structure that changes shape rapidly in response to alterations in osmotic or metabolic conditions (Hackenbrock, 1966). A physical property of the inner membrane that contributes to its flexibility is the intrinsic curvature of its constituent monolayers. Minimum curvature energy is reached when the monolayer bends to shape equal to its intrinsic curvature (Epand et al., 2002). When a bilayer has a large intrinsic negative monolayer curvature, it spontaneously converts from a flat structure to an inverted hexagonal phase. Membrane monolayer curvature may have particular importance in relation to the functioning of mitochondria. It is known that mitochondrial lipids will convert from a lamellar to a hexagonal phase in the presence of Ca2þ (Cullis et al., 1980; Nicolay et al., 1985). Nonbilayer structures have been observed in intact mitochondria (Van Venetie and Verkleij, 1982). The propensity of the mitochondrial membrane to form hexagonal phases to modulate the movement of Ca2þ through the membrane (Wolkowicz, 1988) as well as the activity of certain mitochondrial enzymes (Li et al., 1995) has been suggested. The neuron, like all other cells, is enclosed by a plasma membrane, a double layer of phospholipid molecules that acts not only as a barrier preventing the contents of the cell from mixing with that of the extracellular space, but also acts as an effective electrical insulator, hindering the diffusion of charged ions in and out of the cell. Mitochondria supply the energy needs of the neuron. Because a great deal of energy is required to maintain the transmembrane ionic gradients that are essential for neuronal signaling, neurons tend to be particularly rich in mitochondria. Neurons are therefore intensively involved in the building of large amounts of membrane components, proteins, and lipids for the assembly and remodeling of mitochondrial structures. Of the approximately 80 proteins of the respiratory chain, 13 are encoded by mtDNA and the others, including cyt c, are encoded by nDNA (DiMauro, 2004). Proteins encoded by nuclear genes are imported from the cytosol in the form of precursor proteins that usually contain an amino terminal signal sequence recognized by receptors on the surface of the mitochondrial outer membrane. The classical mitochondrial preproteins carry positively charged amino‐terminal presequences that direct them to the translocase of the outer membrane (TOM) and subsequently to the presequence translocase of the inner membrane (TIM23 complex; Wiedemann et al., 2003). The manner in which apo‐cyt c passes across the mitochondrial outer membrane defines a specific pathway differing from other preproteins. Indeed apo‐cyt c does not carry a cleavable N‐terminal targeting sequence. Two terminal segments of the protein are known to be important for targeting (Jordi et al., 1989; Nye and Scarpulla, 1990; Sprinkle et al., 1990). Moreover, unlike most other mitochondrial preproteins an electrochemical potential across the mitochondrial inner membrane is not required for transport (Mayer et al., 1995). In the intermembrane space apo‐cyt c is converted to cyt c and anchors the inner membrane through acidic phospholipids. Genetic classification of mitochondrial diseases is related to mutations in mtDNA and nDNA. Interestingly, respiratory chain components are under the dual control of both nuclear and mitochondrial genome. Essential clinical features of mtDNA and nDNA mutations producing impairment in mitochondrial protein synthesis have been reviewed (DiMauro, 2004). Disorders due to mutations in nDNA are more abundant due to the high number of mitochondrial proteins encoded by nDNA, but also because the correct assembly and functioning of respiratory machinery is under nDNA control. Many of the components of the respiratory chain are strictly embedded and interact with the lipid bilayer of the inner mitochondrial membrane. Alterations of the lipid medium may cause disease, as in Barth syndrome, a clinical manifestation of disorders in CL metabolism. Barth syndrome describes a unique mutation in Xq28 (G4.5 or tafazzin) (Barth et al., 1999). The tafazzin gene is expressed mainly in cardiac and skeletal muscles and encodes a family of proteins that are homologous to phospholipid acyltransferase (Bione et al., 1996). In this syndrome, a single mitochondrial CL species was affected selectively whereas other phospholipids were normal. Tetralynoleyl CL species are dramatically reduced in skeletal muscle, platelets, and lymphoblast

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from patients with Barth syndrome (Schlame et al., 2002; Xu et al., 2005). Taking into account that linoleic acid represents more than 80% of total fatty acids of CL in normal tissues, it has been speculated that the dramatic decrease in CL is due to mutations of the G4.5 gene product and the consequent defective functioning of acyltransferase in remodeling of CL and its precursor, phosphatidylglycerol (PG) (Vreken et al., 2000). No data are available on brain CL in the pathology. However, linoleic acid in rat brain CL represents no more than 20% of total fatty acid, unsaturation being replaced in part by tetra‐ and hexa‐ unsaturated species (Piccotti et al., 2004). In heart mitochondria, only the newly synthesized PG pool is used for CL synthesis (Hatch, 1996). In addition, newly formed CL may be remodeled by a deacylation– reacylation pathway (Schlame and Rustow, 1990; Hatch, 1998). When a linoleic acid pool is available, it is reasonable to assume that the remodeling pathway is operating also in the brain.

2

Biochemical and Functional Characterization of Purified Brain Mitochondria

Purity of mitochondria is an essential prerequisite in the analysis of lipid composition. Cross‐contamination of isolated subcellular fractions complicates the interpretation of analytical data. Compared to other organelles, mitochondria of acceptable purity can be isolated from brain tissue. All purification procedures are performed through centrifugation on gradient medium. The Percoll gradient procedure described by Sims (1990) is rapid and gives metabolically active mitochondria, although with a low yield. Lipid composition in synaptic and nonsynaptic mitochondria from rat brain was determined after purification in dextran medium (Ruggiero et al., 1992). Preparation on sucrose gradients was performed by Butler and Morell (1983). In our laboratory, brain mitochondria are routinely prepared using sucrose as gradient medium, although Percoll or Ficoll‐400 (Lai and Sheu, 1985) have also been used. The procedure of mitochondria purification with sucrose yields highly purified preparations, also devoid of MAM, that are released from mitochondria and recovered at a sucrose concentration lower than that of mitochondria in a low ionic strength medium (Camici and Corazzi, 1995). The biochemical characterization of preparations indicated that NADPH:cyt c reductase (microsomal marker) was not detectable in mitochondria whereas monoamino oxidase and cyt c oxidase (mitochondrial enzymes) were enriched 3.9 and 5.7 times, respectively, in mitochondria compared with the homogenate. Naþ, Kþ‐ATPase, and arylsulfatase A were not detected in mitochondria. Other positive or negative enzymatic markers are reported (Monni et al., 2000). Mitochondria were metabolically active with a respiratory control ratio (state 3 to state 4) in the range of 5–6, a value in agreement with data reported in the literature (Lai and Rex Sheu, 1985). Functionality of mitochondrial preparations was assayed by monitoring Dcm. Complete Dcm collapse and a maximal Dcm value were obtained in the presence of CCCP and nigericin, respectively (0% and 100% in the potential value scale). Dcm was sensitive to respiratory substrates. The lowest value was measured in deenergized state. Dcm increased on the addition of phosphate, whereas the addition of ADP, malate, pyruvate, and phosphate produced a decrease in Dcm, compared with phosphate alone, due to the consumption of membrane potential in state 3. Permeabilization or removal of the outer mitochondrial membrane was achieved by treatment with digitonin (Schnaitman and Greenwalt, 1968). In our experimental conditions, digitonin acted on the outer mitochondrial membrane without significantly affecting the inside of mitochondria. Only 4–5% of total CL and 14–15% of total mitochondrial phospholipids were solubilized and recovered in the postdigitonin supernatant.

2.1 Lipid Composition of Brain Mitochondria Lipid to protein ratio for phospholipid or cholesterol in brain mitochondria is lower than in other organelles (> Table 8-1). At the same time, phosphatidylcholine (PC) and phosphatidylethanolamine (PE) are the major phospholipid species of mitochondria and of other organelles. Mitochondria specifically contain CL and PG (4–5% and 3–4%, respectively, of total phospholipid) that are not found in other

Lipids of brain mitochondria

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. Table 8-1 Lipid content of subcellular fractions from rat brain cortex Phospholipids Cholesterol

Homogenate 578  10a 300  11b

Microsomes 720  10a 446  7b

Myelin 960  15a n.d.

MAM 570  12a n.d.

Mitochondria 400  12a 71  5b

321.9  16.9c 114  9.3c

Note: Data are expressed as nmol lipid/mg protein; n.d., not determined a Corazzi et al., 1993; b Monni et al., 2000; c Ruggiero et al., 1992

subcellular membranes. Mitochondria do not contain sphingomyelin and glycosphingolipids. In our laboratory, phospholipid composition of the outer and inner membranes was determined after digitonin treatment and purification of mitoplasts (Camici and Corazzi, 1995). A distinctive feature of mammalian mitochondria, including mitochondria from brain, is that the phospholipid to protein ratio of the mitochondrial outer membrane is higher than that of the inner membrane (unpublished data from our laboratory). PC and PE are the major phospholipids of mitochondrial membranes, both being enriched in the inner relative to the outer membrane. Phosphatidylinositol (PI) and phosphatidylserine (PS) are almost equally distributed on both membranes, whereas CL and PG are concentrated mainly on mitoplasts (> Table 8-2). Plasmalogens of choline and ethanolamine are found in brain mitochondria (Eichberg et al., 1964), where a relevant plasmalogenase activity was also detected (Ansell and Spanner, 1968). Based on the estimates of total PE plasmalogen in mitochondrial membranes (Sun et al., 1988) and on the relative amount of plasmalogen in PE and PC fractions (Horrocks and Sun, 1972), about 30% PE is ethanolamine plasmalogen whereas about 15% PC is choline plasmalogen (Sun and Gilboe, 1994). Cholesterol in mammalian mitochondria is usually low compared with the other membranes. Characterization of subcellular fractions from rat brain indicates that cholesterol is about 70 nmol/mg protein, i.e., less than 16% of total mitochondrial phospholipids (Corazzi et al., 1993). As reported earlier, EM tomography has been used to redefine the membrane architecture of mitochondria in neurons. A structurally distinct type of contact site may play a role in the structural integrity of the outer and inner membranes (Perkins et al., 2001). In mammalian cells, these sites are places for protein and lipid passage from outside into the mitochondrion (Brdiczka, 1991). Ardail and colleagues (1990) identified two populations of mitochondrial membrane contact sites in liver mitochondria with characteristics of the inner and outer membrane, respectively, and similar phospholipid to protein ratios, but different cholesterol content. Surprisingly, CL is more highly enriched in these contact sites than in the mitochondrial inner membrane. The observation that adriamycin, which interacts with CL (Goormaghtigh and Ruysschaert, 1984), inhibits protein import in mitochondria (Eilers et al., 1989) suggests that in contact sites the negative‐charged CL may be involved in the import of proteins by interacting with the positive‐ charged signal sequence of the protein in importation. A genetic model system to study the role of anionic phospholipids in yeast mitochondria reveals that the complete lack of CL causes a decrease in mitochondrial membrane potential (Jiang et al., 2000) and inhibition of translation of protein components to the ETC (Ostrander et al., 2001). Isolation of rat brain mitochondria of a fraction enriched in boundary membrane contact sites (Sandri et al., 1988) and their electrophysiological characterization (Moran et al., 1990) were performed. However, to date, no information is available on phospholipid composition of contact sites purified from brain mitochondria. Nonyl acridine orange (NAO) was also used to probe CL of brain mitochondria. About 66% of NAO fluorescence was associated with mitoplasts, confirming that CL was localized mainly in the inner mitochondrial membrane (Piccotti et al., 2002). NAO fluorescence increased noticeably in mitoplasts when mitochondria were fused with exogenous CL, indicating that the lipid enriched the inner membrane. This finding is consistent with a model in which CL fuses with the outer mitochondrial membrane and flows inside through the contact points. The inhibitory effect of cyclosporin A on CL liposome fusion, the

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Lipids of brain mitochondria

. Table 8-2 Phospholipid composition of mitochondria from rat brain cortexa Phosphatidylcholine Phosphatidylethanolamine Phosphatidylserine Phosphatidylinositol Cardiolipin

Intact mitochondria 43.4 40.1b 38.6 37.0b 9.0 4.7b 4.4 4.2b 4.5 14.0b

Mitoplasts 45.5 39.6 6.1 3.9 4,9

Outer membrane 40 38.9 14.2 5.9 0.8

a

Unpublished data from our laboratory. Data are expressed as percent of total phospholipids. Phospholipid content: mitochondria, 393 nmol/mg protein; mitoplasts, 250 nmol/mg protein; outer membrane, 560 nmol/mg protein. Protein content of the outer membrane is about 35% of the total b Ruggiero et al., 1992. Data are expressed as percent of total phospholipids. Phospholipid content of mitochondria is 321.9 nmol/mg protein

localization of mPTP on the contact points (Crompton et al., 1999), and the effect of potassium phosphate on mPTP suggest that mPTP opening is necessary for CL molecules to migrate into the inner mitochondrial membrane (Piccotti et al., 2002). The transbilayer asymmetry of lipids in membranes has assumed an increasing number of roles in apoptosis (Balasubramanian and Schroit, 2003). In mitochondria, CL is sequestered on the inner leaflet of the inner mitochondrial membrane. However, during apoptosis the lipid translocates to the outer surface of the outer mitochondrial membrane (Garcia et al., 2002; Qi et al., 2003). Using a model system Epand and group proposed that the increase in the rate of transbilayer diffusion of CL could be mediated by an activated form of Bax (Epand et al., 2003). Transbilayer orientation of phospholipids in membranes has been studied using nonpenetrating chemical probes. Trinitrobenzenesulfonate and fluorescamine were used to probe PE and PS. Phospholipases used in mild conditions or lipid transfer proteins that selectively remove exposed phospholipids were used for other phospholipids (Crain, 1990). CL was probed with specific antibodies (Krebs et al., 1979) or with the fluorescent probe NAO (Piccotti et al., 2002). In liver mitochondria, PC and PE are equally distributed between the inside and outside leaflets of the inner membrane, whereas PI and CL face the matrix side (Daum and Vance, 1997). PC is evenly distributed between the two leaflets of the outer membrane (Daum and Vance, 1997). Hovius and group observed that the majority of PE is exposed in the cytosolic leaflet of the outer membrane, whereas PI and PS are oriented toward the intermembrane space (Hovius et al., 1993). However, studies performed on intact liver mitochondria with fluorescent pyrene–PE species indicate that the majority of PE is located on the inner leaflet of the outer mitochondrial membrane (Jasinska et al., 1993). Few and limited are the observations on the location and topography of lipids in the membranes of brain mitochondria. Data from our laboratory suggest an asymmetric distribution of PE across the outer mitochondrial membrane, 25–30% being located on the outer leaflet (Camici and Corazzi, 1995). Generally, the percentage of unsaturated fatty acids of mitochondrial lipids is always higher than in other subcellular fractions (Daum, 1985). In brain, the pattern of fatty acids in total synaptic and nonsynaptic mitochondria has been determined (> Table 8-3) (Ruggiero et al., 1992). In both synaptic and nonsynaptic mitochondria the majority of fatty acids belong to saturated and monounsaturated species (63% and 62%, respectively). Polyunsaturated fatty acids contain 18:2n–6, 20:3n–6, 20:4n–6, and 22:6n–3. In nonsynaptic mitochondria, a significantly higher content of three‐ and hexa‐unsaturated fatty acids was found in the replacement of 20:4n–6. Fatty acid composition of single phospholipid classes has been determined by different authors (> Table 8-4). The results, although characterized by high variability, indicate that, except CL, all phospholipid classes contain high percent of saturated fatty acids. Monounsaturated species are contained in PC and CL, whereas 18:2n–6 is prevalently in CL. Polyunsaturated fatty acids (20:4n–6 and 22:6n–3) constitute more than 20% in all classes, except PG.

Lipids of brain mitochondria

8

. Table 8-3 Pattern of phospholipid fatty acids in synaptic and nonsynaptic mitochondria from rata 16:0 18:0 18:1 (n–9) 18:2 (n–6) 20:3 (n–6) 20:4 (n–6) 22:6 (n–3)

Nonsynaptic mitochondria 29.6  1.6 15.0  1.2 17.4  1.0 15.0 1.1 1.7  0.4 15.4  1.0 5.9  0.6

Synaptic mitochondria 27.9  1.2 16.8  1.0 18.4  1.1 14.0  0.9 0.5  0.3 21.8  1.3 0.6  0.3

Note: Data are mean  SD and are expressed as percent (wt/wt) Ruggiero et al., 1992

a

Dietary fatty acids change the fatty acid profile of phospholipid classes in brain mitochondria (Dyer and Greenwood, 1991). Fatty acid analysis of brain mitochondrial PE, PC, and CL revealed that the largest dietary effect is on 18:2n–6, which is 30% higher in rats fed with a diet rich in essential fatty acids (Dyer and Greenwood, 1991). It has been shown that, despite the high baseline levels of 18:2n–6 in heart mitochondrial CL compared with brain, 18:2n–6 levels in CL increase also in brain in proportion to dietary 18:2n–6 supply (McGee et al., 1996). Adversely, n–3 fatty acid deficiency decreases PS selectively, in agreement with the observation that 22:6n–3 is highly enriched in this lipid in brain mitochondria (Hamilton et al., 2000).

2.2 Cytochrome c–Cardiolipin Interactions CL is the only mitochondrial phospholipid that is synthesized in mitochondria. During the isolation of mitochondrial proteins, CL coisolates with each protein that participates in oxidative phosphorylation. CL has been claimed to be a proton trap for oxidative phosphorylation (Haines and Dencher, 2002) and is necessary for cyt c insertion, retention, stability, and function (Rytomaa and Kinnunen, 1994; Choi and Swanson, 1995). A feature of liver and heart CLs is the peculiar presence of 18:2n–6 that always exceeds 80%. In contrast, CL from brain mitochondria contains about 48% polyunsaturated fatty acids of which only 20% is 18:2n–6. To date, there is no explanation for this different fatty acid composition in CL between tissues. CL is claimed to anchor cyt c through hydrophobic and hydrophilic interactions (Tuominen et al., 2002). Exposure of brain mitochondria to phosphate results in the release of cyt c outside mitochondria (Piccotti et al., 2002). We have used a reconstituted system of cyt c in CL liposomes as a model to investigate the effect of phosphate on the hydrophilic component of cyt c–CL interactions (Piccotti et al., 2004). As for intact mitochondria, phosphate‐dependent cyt c release from reconstituted liposomes requires at least 10 mM phosphate, indicating this value as the threshold phosphate concentration able to discharge the loosely bound cyt c pool. About 44% of cyt c is released by increasing phosphate concentration. The remaining cyt c could account for the tightly bound conformation characterized by hydrophobic interactions. The relevance of the hydrophobic interaction component in cyt c–CL association was further demonstrated by comparing the phosphate‐dependent cyt c detachment from liposomes made with CL extracted from heart or brain mitochondria, highly different in linoleic acid content. Binding of cyt c to CL from heart mitochondria was stronger than that to CL from brain mitochondria (Piccotti et al., 2004). In cyt c– CL interactions, the adoption of the extended conformation (Ryto¨maa and Kinnunen, 1995) requires the protrusion of the sn–2 acyl chain, in which cis double bonds allow for greater conformational flexibility (Kinnunen et al., 1994; Tuominen et al., 2002). Therefore, the stronger cyt c–heart CL hydrophobic interaction is imputable to 18:2n–6 that should represent the fit acyl chain in this type of interaction. Consequently, the weaker hydrophobic interaction of cyt c with CL purified from brain mitochondria can be explained by the presence of more saturated acyl chains and less linoleic acid (Piccotti et al., 2004).

205

PCb – 38.9  1.1 – 20.6  0.4 22.1  0.8 0.98  0.04 – 10.5  0.4 – 12.1  0.5 –

PCc 0.7  0.06 27.8  1.4 – 25.1  1.1 8.2  0.7 1.2  0.06 – 29.2  1.8 – 7.8  0.5 –

PEa – 6.1  1.0 – 28.2  0.6 9.1  0.9 – 0.4  0.2 18.2  0.9 4.8  0.4 32.7  0.5 0.4  0.2

a

Note: Data are mean  SD and are expressed as percent (wt/wt) Nakahara et al., 1991; b Sun and Gilboe, 1994b; c Unpublished data from our laboratory

14:0 16:0 16:1 18:0 18:1 18:2 18:3 20:4 22:4 22:6 others

PCa – 34.4  1.3 – 12.0  0.8 25.4  0.3 0.7  0.2 0.5  0.2 13.9  0.4 0.6  0.3 12.2  0.8 0.4  0.3

PEb – 13.7  0.6 – 24.7  0.3 18.6  0.6 2.1  0.1 0.4  0.2 19.5  0.5 – 22.8  0.6 –

PEc 0.4  0.02 7.9  0.4 – 37.1  1.9 7.0  0.3 0.2  0.02 – 17.2  1.3 – 27.0  1.4 3.4  0.2

PSa – 1.8  0.6 – 42.8  0.7 5.8  0.6 – 0.4  0.2 3.5  0.9 2.2  0.7 38.8  0.4 5.2  1.5

PSc 1.6  0.08 11.7  0.7 0.7  0.07 58.9  3.0 7.0  0.4 – 0.3  0.02 4.4  0.3 – 13.7  0.8 1.7  0.2

PIa – 10.1  1.3 – 33.1  1.6 3.8  0.2 – – 40.4  4.7 – 10.0  1.8 2.6  0.4

CLa – 4.1  0.2 – 1.3  0.5 26.0  1.1 8.1  0.8 – 20.8  2.2 0.8  0.4 17.3  0.9 21.6  4.6

CLc 1.4  0.1 12.8  1.0 5.1  0.4 11.7  1.0 27.6  2.5 12.5  0.6 – 19.3  1.1 – 8.0  0.6 1.6  0.1

PGc 2.1  0.1 28.4  1.7 2.4  0.2 37.2  2.9 10.1  0.8 3.7  0.3 0.8  0.04 7.6  0.6 – 4.9  0.4 2.8  0.3

8

. Table 8-4 Fatty acid compositions of mitochondrial phospholipids in rat brain

206 Lipids of brain mitochondria

Lipids of brain mitochondria

8

2.3 Biosynthetic Origin of Mitochondrial Lipids The majority of glycerophospholipids occurring in brain cells are synthesized within the endoplasmic reticulum. From this site of synthesis, phospholipids are distributed to the proper locations and inserted in the bilayer structures. Organelle biogenesis and intracellular lipid transport in eukaryotes have been reviewed (Voelker, 1991). In mitochondria, phospholipid synthesis is restricted to the formation of PG, CL, and PE. The other phospholipids are imported from the endoplasmic reticulum (as observed in experiments performed in vivo) (Butler and Morell, 1983; Bjerve, 1985). Several hypotheses have been put forth to better define the intracellular trafficking of lipids. All are consistent with a flow of molecules from the donor particles (microsomes) to the acceptor particles (mitochondria) through mediator proteins or through vesicle‐mediated transfer. However, a translocation of lipids from the endoplasmic reticulum to mitochondria may also occur because of physical contact and fusion of mitochondria with MAM (Rusin˜ol et al., 1994). Ultrastructural studies revealed continuity between the endoplasmic reticulum and the outer mitochondrial membrane (Katz et al., 1983) as well as possible points of fusion between these membranes (Cascarano et al., 1982). MAM have been putatively involved in the synthesis of phospholipids and in their direct transfer to mitochondria (Camici and Corazzi, 1995). Mitochondria contain low amounts of PS. In addition, this lipid is the substrate of PS decarboxylase, an enzyme localized on the outer surface of the inner mitochondrial membrane (Percy et al., 1983). Since PS is not synthesized in mitochondria, its importation is necessary. PS synthesis in brain occurs through a base‐exchange mechanism (Porcellati et al., 1971) localized in the endoplasmic reticulum, particularly in MAM (Monni et al., 2000). At least two serine‐base‐exchange enzyme isoforms are present in brain but their biochemical properties and regulation are still largely unknown (Mozzi et al., 2003). An enzyme with the capacity to synthesize PS by serine‐base‐ exchange with PE, but not with PC, was partially purified from rat brain (Suzuki and Kanfer, 1985). The existence of two mammalian PS synthase (1 and 2) was established with mutant CHO‐K1 cells (Kuge et al., 1985; Voelker and Frazier, 1986). These enzymes were localized to MAM (Stone and Vance, 2000). A PS synthase‐1 cDNA from murine liver was also cloned (Stone et al., 1998). Metabolism and functions of PS have been reviewed (Mozzi et al., 2003; Vance and Steenbergen, 2005). Import of PS in mitochondria occurs in a reconstituted system made by mixing microsomes prelabeled with 14C‐PS and mitochondria (Voelker, 1989). In this experimental model, translocation of phospholipids results from collision complexes formed between the endoplasmic reticulum and the outer mitochondrial membrane. PS translocation is enhanced by Ca2þ but not influenced by cytosolic factors (Corazzi et al., 1993). The specific radioactivity of PS transferred to mitochondria is higher than that of microsomal PS. This finding supports the hypothesis that the lipid is compartmentalized in microsomes and that radioactive newly synthesized PS is better exported than the bulk of microsomal phospholipids (Corazzi et al., 1993). We also demonstrated that the amount of PS transferred from MAM to mitochondria correlates with the energized state of mitochondria. During respiration MAM associate with mitochondria, thus promoting a flow of PS to mitochondria (Monni et al., 2000). However, the nature of the contact between MAM and the outer mitochondrial membrane and the identification of factor(s) involved in PS translocation are yet unclear. We tested the possibility that mitochondria acquire PS through a fusion process. 14C‐PS‐labeled liposomes fuse to the outer mitochondrial membrane at acidic pH. Fusion is associated with a protein factor localized on the mitochondrial membrane. The protein factor has been partially purified and some of its properties have been described (Camici and Corazzi, 1997). The presence of a fusogenic protein in mitochondrial membranes strengthens the morphological evidence that the contact and fusion points observed could be responsible for phospholipid transfer from the endoplasmic reticulum to mitochondria. Imported PS is translocated from the outer to the inner mitochondrial membrane where it is decarboxylated to PE through PS decarboxylase activity. This enzyme contributes to the synthesis of total PE in brain for at least 7% (Butler and Morell, 1983). In the liver, the PE formed flows back to the outer membrane and then reaches the endoplasmic reticulum (Vance, 1991; Hovius et al., 1992). Conversely, in brain PE produced by PS decarboxylation appears to be used in the assembly of the inner mitochondrial membrane (Carlini et al., 1993; Camici and Corazzi, 1995). PE can be imported also from the endoplasmic reticulum where it is formed by the CDP–ethanolamine pathway (Kennedy and Weiss, 1956) or by

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base‐exchange reaction (Corazzi et al., 1986). Movement of PE from endoplasmic reticulum to mitochondria was observed in hepatocytes and kidney cells in culture (Yaffe and Kennedy, 1983). Moreover, experiments in vivo showed that PE formed in brain endoplasmic reticulum is translocated to mitochondria (Butler and Morell, 1983). Factors influencing the importation of PE in brain mitochondria and its utilization for the assembly of mitochondrial membrane have been studied (Camici and Corazzi, 1995). PE was imported in mitochondria when a rat brain homogenate was incubated with [3H]ethanolamine and subcellular fractions were subsequently isolated. Ca2þ and the nonspecific lipid transfer protein purified from rat liver enhanced translocation in vitro. The importation of PE in mitochondria was also shown in a reconstituted system made of microsomes (donor particles) and purified mitochondria (acceptor particles). 3 H‐PE synthesized in MAM could also translocate to mitochondria in the reconstituted system. Experiments in which mitochondria were exposed to trinitrobenzene sulfonate and other experiments in which the mitochondrial outer membrane was selectively removed by digitonin treatment indicated that imported PE was localized mainly to mitochondrial outer membrane, whereas PE generated from the decarboxylation pathway was confined primarily to mitochondrial inner membrane (Camici and Corazzi, 1995). PC is the most abundant lipid of brain mitochondria, comprising about 40% of total phospholipids. Since neither inner nor outer mitochondrial membranes contain enzymatic activities for synthesis, all mitochondrial PC must be imported. PC is synthesized by the CDP–choline pathway (Roberti et al., 1980) whose last enzyme, CDP–choline:1,2‐diacylglycerol choline phosphotransferase, resides in the endoplasmic reticulum. Base‐exchange activity may also contribute to PC synthesis in the nervous tissue (Kanfer, 1972; Arienti et al., 1976). In yeast and hepatocytes, a second pathway of PC biosynthesis exists in which PE is methylated to PC by methylation reactions (Marinetti et al., 1976; Vance and Vance, 1988; Cui et al., 1993). Adversely, low methyltransferase activity was detected in the nervous tissue (Mozzi and Porcellati, 1979; Dainous et al., 1982). In brain, choline phosphotransferase and choline base‐exchange activities are localized on the outer surface of microsomal vesicles (Arienti et al., 1985). An asymmetric distribution of PC was found in brain microsomes (Dominski et al. 1983), the majority being exposed on the outer surface of microsomes that corresponds to the cytosolic compartment in situ. This localization may help in the exportation of the newly synthesized lipid toward mitochondria. Pulse‐chase experiments carried out in tissues different from brain demonstrated that PC migrated sequentially from the endoplasmic reticulum to the mitochondrial outer membrane and then to the inner membrane (McMurray and Dawson, 1969; Jungalwala and Dawson, 1970). Although translocation from endoplasmic reticulum to the mitochondrial outer membrane occurs rapidly, movement to the inner membrane is a slower process (Eggens et al., 1979; Wojtczak et al., 1990). Experiments performed with labeled PC introduced into the outer membrane of mitochondria using a PC‐specific transfer protein demonstrated that the lipid equilibrates rapidly over both leaflets of the outer membrane (Dolis et al., 1996). In in vitro systems made of fluorescent donor vesicles and acceptor mitochondria the imported lipid localizes exclusively in the outer membrane, suggesting that additional factors are required for the transfer of PC to the inner membrane (Nicolay et al., 1990). Despite these studies, very little is known about the mechanism(s) governing PC transport, with complete lack of information on PC import in brain mitochondria. The metabolic pathways for the synthesis of phosphatidic acid (PA) in the central nervous system (CNS) have been reviewed (since 1981 by Bazan and group). Despite its importance as diacylglycerol and CDP‐diacylglycerol precursor, this lipid is a quantitatively minor component in brain mitochondria. Diacylglycerol is used for the synthesis of PE and PC, whereas CDP‐diacylglycerol is the starting point for the synthesis of PI, PG, and CL. Glycerol‐3‐phosphate is the precursor for the synthesis of PA by two acyltransferases acting consecutively, acyl‐CoA:sn‐glycerol‐3‐phosphate acyltransferase and acyl‐CoA:1‐ acylglycerol‐3‐phosphate acyltransferase. In the liver, glycerol‐3‐phosphate acyltransferase exists in two forms, one is localized in the endoplasmic reticulum and the other in mitochondria (Bell and Coleman, 1983). Purification and characterization of glycerophosphate acyltransferase from rat liver mitochondria have been performed (Vancura and Haldar, 1994). The subcellular distribution of glycerol‐3‐phosphate acyltransferase between rat brain mitochondria and microsomes has also been investigated (Fitzpatrick et al., 1982). The activities associated with purified brain mitochondria and microsomes may be distinguished by differences in acyl‐CoA specificity and sensitivity to N‐ethylmaleimide (Fitzpatrick et al., 1982). The second acylation reaction is catalyzed by 1‐acylglycerol‐3‐phosphate acyltransferase, whose activity is

Lipids of brain mitochondria

8

localized mainly in the endoplasmic reticulum. It has been inferred that the product of the first acylation reaction (lyso‐PA) is exported from the mitochondrion to the endoplasmic reticulum to form PA via the second acyltransferase. Experiments carried out in liver, but not in brain mitochondria, demonstrate that lyso‐PA produced in mitochondria leaves the organelles and is converted to PA in the endoplasmic reticulum (Haldar and Lipfert, 1990). Vice versa, PA necessary for the synthesis of PG and CL moves from the reticulum toward mitochondria where the synthesis of these compounds is restricted. Little information is available on the transfer of PA from reticulum to mitochondria in the liver (Baranska and Wojtczak, 1984; Wojtczak et al., 1990), whereas there is none available on the brain. The pathway of CL biosynthesis in mammalian cells and the factors regulating gene transcription in CL synthetic pathway have been given little attention (McMillin and Dowhan, 2002). These are very important issues since limitations in CL levels have significant effects on electron carrier proteins and on the pathways that initiate programmed cell death.

2.4 Import of Fatty Acids into Mitochondria Fatty acids are a major source of energy for many animal cell types. The b‐oxidation of fatty acids, with the resultant generation of ATP, takes place in the mitochondrial matrix. Exceptionally brain mitochondria oxidize fatty acids poorly or not at all but obtain the greatest amount of energy from glucose metabolism. b‐oxidation enzymatic activities in rat brain and heart mitochondria were measured and compared (Yang et al., 1987). It was found that the low rate of [1–14C]palmitoylcarnitine degradation in brain mitochondria may be the consequence of the low activity of 3‐ketoacyl‐CoA thiolase. However, in vivo the brain of diabetic mice exhibits a decreased capacity for glucose oxidation and increased capacity for fatty acid oxidation. In fact, isolated cerebral mitochondria oxidize palmitate to CO2 at a rate almost twice that of control mitochondria (Makar et al., 1995). In brain mitochondria, fatty acid utilization is therefore restricted to biosynthetic purposes. Importation of exogenous fatty acids in brain mitochondria occurs through the mechanism common to all tissues, which requires carnitine and carnitine palmitoyltransferase (Kerner and Hoppel, 2000). The carnitine‐acylcarnitine translocase is one of the components of the carnitine cycle. The carnitine cycle is necessary to shuttle long‐chain fatty acids from the cytosol into the intramitochondrial space where activated fatty acids will be used. Through this mechanism, mitochondria may acquire essential fatty acids that are inserted in mitochondrial lipid molecules. Carnitine carrier was purified from rat brain mitochondria and reconstituted into PC vesicles. The activity of the carrier varied with age, being twice as high in suckling rats than in adults (Kaminska et al., 1993). Carnitine palmitoyltransferase and carnitine octanoyltransferase activities in brain mitochondria were three‐ to fourfold lower than in liver activities. CPT‐1, the overt form of carnitine palmitoyltransferase, was strongly inhibited by malonyl‐CoA (Bird et al., 1985). In addition, it has been shown that inhibition of CPT‐1 activity in brain caused a decrease of food intake in rats (Lavrentyev et al., 2004). mRNA expression of three known CPT‐1 isoforms in different brain regions of normal, fasting, and insulin‐dependent diabetic rats was examined. Compared with the expression observed in liver and heart, there was either little or no difference depending on the particular brain region examined, suggesting that regulation of CPT‐1 mRNA levels is different in the brain compared with other tissues (Lavrentyev et al., 2004). Clinical, biochemical, and genetic aspects of carnitine–acylcarnitine translocase deficiency have been reported. Brain and other organs are involved in this pathology and most patients become symptomatic in the neonatal period with a high mortality rate (Rubio‐Gozalbo et al., 2004). The deacylation–reacylation cycle is an important mechanism responsible for the introduction of polyunsaturated fatty acids into neural membrane glycerophospholipids. It involves four enzymes, namely acy‐lCoA synthetase, acyl‐CoA hydrolase, acyl‐CoA:lysophospholipid acyltransferase, and phospholipase A2. These enzymes have been purified and characterized from brain tissue (Farooqui et al., 2000a). Remodeling by deacylation–reacylation may be an important contribution in maintaining a specific lipid profile in mitochondria. In liver mitochondria, complete remodeling of tetraoleoyl‐CL to tetralinoleoyl‐CL by a specific phospholipid transacylation was observed (Xu et al., 2003). Brain mitochondria contain phospholipase A2 activity necessary for acyl group release from mitochondrial phospholipids (Macchioni et al.,

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2004). Although experimental evidence supports phospholipid remodeling by deacylation–reacylation (Kevala and Kim, 2001), no data are available on the presence of acyltransferase activity in brain mitochondria.

2.5 Cholesterol Cholesterol metabolism in the CNS appears to be distinct from that in other tissues because the CNS and plasma are separated by the blood–brain barrier. Many studies reported that all the cholesterol required for the development of the brain and spinal cord is derived from endogenous synthesis within the CNS (Jurevics and Morell, 1995; Turley et al., 1998). Compared with the other tissues, cholesterol is highly enriched in brain (Dietschy and Turley, 2004); sterols are found predominately in the unesterified form with small amounts of desmosterol (Vance et al., 2005). Both neurons and glial cells contain cholesterol, although myelin contains the major pool of cholesterol in brain. Surprisingly, many of the proteins involved in transporting cholesterol in the circulating plasma, as well as lipoprotein receptors, have been found in the CNS, suggesting involvement in cholesterol transport in brain cells. The major apolipoproteins in CNS are apo‐E and apo‐J (Gong et al., 2002), both inserted in cholesterol‐containing lipoproteins resembling the size and density of HDLs (Pitas et al., 1987). In CNS, they are proposed to bind to neuronal surface receptors after synthesis and secretion mainly from glial cells. Therefore, glial lipoproteins are thought to be the source of cholesterol, an essential compound for the stimulation of axonal growth of CNS neurons (Vance et al., 2005). In brain mitochondria, cholesterol represents no more than 16% of total lipids (Corazzi et al., 1993) and, since its synthesis does not occur in these particles, importation should be operative. Distinct cholesterol domains characterize the outer membrane of brain mitochondria, revealed also in liver mitochondria (Cremel et al., 1990). Removal of the outer mitochondrial membrane with digitonin produced mitoplasts containing a very low amount of cholesterol (unpublished results from our laboratory). The import of labeled cholesterol from unilamellar vesicles into isolated mitochondria has been evaluated in Saccharomyces cerevisiae (Tuller and Daum, 1995). Transfer of cholesterol to the mitochondrial surface was enhanced in vitro by cytosolic proteins. Translocation between inner and outer mitochondrial membranes was observed in the same incubation system. Labeled cholesterol was translocated to the inner membrane and also detected in contact sites between the two mitochondrial membranes, indicating that contact sites function as bridges where cholesterol molecules are en route to the inner mitochondrial membrane. One candidate protein found to be involved in targeting cholesterol to mitochondria is sterol carrier protein‐2, also known as the nonspecific lipid transfer protein (Wirtz, 1991). This 15‐kDa protein contains a 20 amino acid putative mitochondrial targeting sequence (Trzeciak et al., 1987; Moncecchi et al., 1996). The role of the peripheral‐type benzodiazepine receptor (PBR) in cholesterol import into brain mitochondria has been studied. This receptor is a mitochondrial protein consisting of three subunits: PBR, a voltage‐dependent anion channel, and an adenine nucleotide carrier (Levitt, 1990). The protein is involved in the regulation of cholesterol transport from the outer to the inner mitochondrial membrane, the rate‐determining step in steroid hormone biosynthesis. Molecular modeling of PBR suggests that it might function as a channel for cholesterol. Expression of specific transcription factors results in overexpression of PBR and increased cholesterol transport into mitochondria of tissues with a specialized function (steroidogenesis) (Papadopoulos, 2004). Steroid hormones are synthesized in the adrenals, gonads, placenta, and CNS. Regardless of tissue origin, a common feature of all steroid hormones is that their synthesis uses cholesterol as a common precursor. Cholesterol residing in the outer mitochondrial membrane must be delivered to the inner mitochondrial membrane, the site of the cytochrome P450 side chain cleavage enzyme, which converts cholesterol to pregnenolone. The delivery of cholesterol to the inner mitochondrial membrane is an assisted process were steroidogenic acute regulatory or StAR protein has also emerged as the best candidate (Stocco, 2000). This protein has been purified, cloned, sequenced, and expressed (Clark et al., 1994). Both the central and the peripheral neural tissues have the capability to synthesize steroids from cholesterol and to metabolize steroid hormones. Proteins involved in the intramitochondrial trafficking of cholesterol, such as PBR and StAR, are expressed in neural tissue (Garcia‐ Ovejero et al., 2005). StAR is expressed in many neuronal subtypes (Kim et al., 2002; King et al., 2002),

Lipids of brain mitochondria

8

whereas PBR is expressed predominantly in microglia and astroglia (Banati et al., 1997; Lacor et al.,1999; Raghavendra et al., 2000; Rao et al., 2001). Neurons, astrocytes, and oligodendrocytes express several enzymes involved in the steroidogenic process and consequently are able to produce different types of steroids (Akwa et al., 1993; Kimoto et al., 1997; Zwain et al., 1997; Mensah‐Nyagan et al., 1999; Mellon and Vaudry, 2001).

3

Physiopathology of Mitochondrial Lipids

3.1 Aging An increased formation of reactive oxygen species (ROS), particularly from mitochondria, is produced in aging brain. ROS have the greatest effects on the mitochondrial inner membrane, the site of their production, and also where iron‐ and copper‐containing enzymes may exert a catalyzing role in the formation of hydroxyl radicals. The age‐related increase in the formation of superoxide radicals and its dismutase product (hydrogen peroxide) is based on the concept that aging modifies the electron flux conditions in the components of ETC, thus enabling electrons to escape the normal flow sequence (Benzi and Moretti, 1995). The leakage of electrons from the energy‐transducing sequence can take place even in the young, indicating that the formation of radicals is associated with the normal respiratory process. However, since the increase in leakage is an age‐dependent event, ROS impair the activity of ETC components as the result of altered membrane lipids, oxidized protein, and damage of mtDNA (Hillered and Ernster, 1983; Zhang et al., 1990; Mecocci et al., 1993). In particular, damage to mtDNA induces alterations to the mtDNA‐encoded polypeptides of the respiratory complexes, with consequent decrease of electron transfer, leading to further production of ROS (Genova et al., 2004). In beef heart, submitochondrial particle generation of ROS affects the activity of complex III through peroxidation of CL, which is required for the functioning of this multisubunit enzyme complex (Paradies et al., 2001). CL is the major target for ROS in brain mitochondria. In neurons, this lipid is lost during apoptotic death, but antioxidants block peroxidation and loss (Kirkland et al., 2002). An age‐related decrease in CL content was also observed (Sastre et al., 2000). Lipid composition in synaptic and nonsynaptic mitochondria from rat brain during aging was determined (Ruggiero et al., 1992). Cholesterol and phospholipids decreased 27% and 12%, respectively, with age, resulting in the decrease of cholesterol/phospholipids ratio. Among phospholipids, only CL showed a significant decrease in nonsynaptic mitochondria from the brains of aged rats. The decrease of linoleic acid observed only in nonsynaptic mitochondria may be related to the decrease of CL. The molecular bases of aging are related to a progressive accumulation of changes caused by the modification of cellular constituents and producing an increased susceptibility to disease and cell death (Harman, 1991). Lipids are one of such constituents that undergo modification during aging. Changes in metabolism influence cellular functions since membrane‐bound enzymes, transport systems, as well as transducing systems are affected by alterations in the properties of the lipid bilayer. Brain aging is accompanied by changes in the activity of enzymes involved in the modification of the polar moiety of phospholipids such as base‐exchange enzymes, PE N‐methyltransferase, phospholipase D, and phosphatidate phosphohydrolase (Ilincheta de Boschero et al., 2000; Pasquare´ et al., 2001; Salvador et al., 2002). Mitochondrial PS decarboxylase shows high specificity toward more hydrophilic PS species (18:0, 22:6n–3) preferentially imported from the endoplasmic reticulum (Heikinheimo and Somerharju, 1998). In aging, the enzyme reduces its activity toward substrates containing these species, but increases its activity toward less‐hydrophilic PS species that are more available in this condition, suggesting that the enzyme adapts its activity to substrate availability (Salvador et al., 2002). Age‐associated changes in CNS glycerolipid composition and metabolism have been reviewed (Giusto et al., 2002).

3.2 Ischemia Preliminary studies showed that production of diacylglycerols enriches in arachidonate and stearate during early brain ischemia (Aveldano and Bazan, 1975). Degradation of phospholipids occurs mainly in

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mitochondrial membranes (Majewska et al., 1978) causing mitochondrial malfunction that affects cellular energy metabolism (Rehncrona et al., 1979). Factors that have been proposed to account for mitochondrial damage during ischemia and reperfusion include intracellular acidosis, Ca2þ‐induced membrane damage, and free‐radical‐dependent membrane lipid peroxidation (Fiskum, 1985). Degradation of mitochondrial phospholipids during experimental cerebral ischemia in rats produced a significant decrease in the amount of PI and CL (54% and 51%, respectively, compared with control) (Nakahara et al., 1991). The content of other phospholipids also decreased, although the decrements were not statistically significant. Changes in the composition of fatty acids were revealed in PC with a decrease in arachidonic and docosahexaenoic acid. Reduction in arachidonic acid content was also observed in PE, whereas docosahexaenoic acid decreased in PS and PI. In addition, ischemia caused a decrease in the amount of whole polyunsaturated fatty acids in each phospholipid class. In contrast, saturated and monounsaturated fatty acids were correlatively increased (Nakahara et al., 1991). The differential degradation of phospholipid classes and the preferential hydrolysis of the polyunsaturated molecular species during ischemia were not followed by reacylation of the mitochondrial phospholipids during long‐lasting normoxic reperfusion. The respiratory control ratio decreased significantly during 30 min of ischemia with no apparent recovery following 60 min of reoxygenation. The amount of phospholipids and the percentage of polyunsaturated fatty acid chains after ischemia lasting for 60 min decreased further after reperfusion, suggesting progressive disruption of phospholipids by phospholipase A2 (Nakahara et al., 1992). The degree of free‐radical‐mediated lipid peroxidation increased during ischemia and reperfusion (Sun and Gilboe, 1994a). In addition, the conversion of phospholipid breakdown products mediated an array of cellular reactions. Platelet activating factor (PAF), arachidonic acid cyclooxygenase, and lipoxygenase products can induce changes that jeopardize cell survival in reperfused tissue (Siesjo and Katsura, 1992; Siesjo et al., 1995). Ischemia‐induced accumulation of mitochondrial‐free fatty acids and loss of polyunsaturated fatty acyl chains from PC and PE were reversed by PAF antagonist, whereas mitochondrial respiration improved simultaneously (Sun and Gilboe, 1994b). In addition, hyperglycemic damage to mitochondrial membrane during cerebral ischemia was improved by PAF antagonist. Particularly, mitochondrial‐free fatty acid release was decreased, whereas reacylation of PC was promoted (Kintner et al., 1997). The incubation in vitro of mitochondria with arachidonic acid, known to be dramatically released in mitochondria during cerebral ischemia, promoted mitochondrial swelling (Hillered and Chan, 1989), an event connected to the activation of mitochondrial permeability transition pore and mitochondrial dysfunction (Kroemer, 1999). Swelling reversal occurred without the recovery of respiratory function. The inhibition of mitochondrial respiration activity by arachidonic acid has been claimed to be a possible mechanism of mitochondrial dysfunction during cerebral ischemia (Takeuchi et al., 1991). Arachidonic acid may inhibit mitochondrial ATP production during brain ischemia and act on the site(s) closely related to NAD‐linked respiration, in addition to its uncoupling effect (Takeuchi et al., 1991). Ischemia influences the fetus. In a model of fetal ischemia/reperfusion using rats at day 19 of pregnancy, ischemia followed by reperfusion was likely to induce fetal brain mitochondrial lipid peroxidation, which may inhibit respiratory activity (Wakatsuki et al., 1999). Multiple beneficial effects were exerted by cytidine 50 ‐diphosphocholine that restored PC, sphingomyelin, and CL levels as well as arachidonic acid content of PC and PE in gerbil hippocampus after 10 min forebrain ischemia/1‐day reperfusion (Rao et al., 2000). Cytidine 50 ‐diphosphocholine lowered the formation of hydroxyl radical by attenuating the activity of the secretory Ca2þ‐dependent PLA2, a predominant isoform in mitochondria (Rao et al., 2003). Forebrain ischemia/reperfusin in normoglycemic and hyperglycemic rats caused mitochondrial dysfunction that lead to cyt c release and caspase activation in hyperglycemia (Li et al., 2001). These findings support the notion that biochemical changes typical of programmed cell death occur after ischemia, thus contributing, in part, to hyperglycemia‐aggravated ischemic neuronal death. Both cyt c and CL were released from brain mitochondria of rats submitted to anoxia and reoxygenation. Mitochondrial membrane lipid peroxidation increased during reoxygenation and was proportional to the decrease in membrane fluidity (Morin et al., 2003). The rate of cyt c and CL release increased with reoxygenation time, whereas the antioxidant a‐tocopherol abolished all reoxygenation‐induced changes. In addition, a nitrone with free‐radical trapping properties inhibited the release of cyt c after focal cerebral ischemia, reducing brain damage through the suppression of the cell‐death pathway initiated by cyt c release (Han et al., 2003).

Lipids of brain mitochondria

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3.3 Altered Lipids in Neurodegeneration Glycerophospholipids in brain, metabolism, incorporation into membranes, and involvement in neurological disorders have been reported in an excellent review (Farooqui et al., 2000b). Alterations in glycerophospholipid composition and increases in lipid hydrolyzing enzyme activities have been reported in Alzheimer’s disease (AD) and other neuropathologies. The presence of elevated glycerophosphocholine concentrations in brain regions and in cerebrospinal fluid as a result of phospholipase activities is the hallmark of membrane phospholipid breakdown during neurodegeneration (Blusztajn et al., 1990; Nitsch et al., 1992; Klein, 2000; Walter et al., 2004). The collective evidence from many studies suggests that neural membrane phospholipid metabolism is disturbed in neural trauma and neurodegenerative disease. This disturbance is caused by different PLA2 isoforms, enzymes involved in signal transduction and whose activities are stimulated in neural trauma and neurodegeneration (Farooqui et al., 2004). A decrease in polyunsaturated fatty acids in the AD brain suggests the increase in free‐radical‐mediated lipid peroxidation. Polyunsaturated fatty acid levels in glycerophospholipids are significantly decreased in the hippocampus of AD patients (Markesbery, 1997). Moreover, neural membranes from AD subjects contain decreased amounts of plasmalogens. The decrease in the ratio of plasmalogen to nonplasmalogen glycerophospholipids produces changes affecting membrane stability (Ginsberg et al., 1995, 1998). Plasmalogens protect polyunsaturated fatty acids from peroxidation (Guan et al., 1999), thus protecting neural cells against oxidative stress in AD patients. 31P nuclear magnetic resonance analysis of lipid extracts from autopsy brain material showed significant changes in AD brain phospholipids compared with age‐matched, nondemented controls (Pettegrew et al., 2001). In AD brain, cells exhibit a membrane defect characterized by accelerated phospholipid turnover. The pattern of phospholipid metabolic changes was mimicked in PC12 cells after exposition to inhibitors of mitochondrial bioenergetics, indicating that mitochondrial function may underlie membrane defects in AD (Farber et al., 2000). Oxidative stress and mitochondrial oxidative damage, including alterations in mitochondrial lipids, have been implicated in the etiology of numerous diseases (Fariss et al., 2005). The consistent reduction of CL and of its precursor PA, synthesized in the endoplasmic reticulum and transferred to mitochondria, indicates that membrane defects caused by altered lipid synthesis in the endoplasmic reticulum may result in the impairment of mitochondrial lipid importation. Mitochondrial CL depletion may be related to the role this lipid plays in the activation of proteins located in the inner mitochondrial membrane (Corazzi et al., 2002). In rat cerebellum TNF‐a, acting through receptors linked to the sphingomyelin pathway, increased sphingomyelinase activity and ceramide levels as well as peroxide products, determining a connection of lipid peroxide with the sphingomyelinase pathway in the development of AD (Alessenko et al., 2004), were observed. Ceramide, accumulated in neurons during various disorders associated with acute or chronic neurodegeneration, induces apoptosis through mitogen‐activated protein kinases and causes the release of multiple mitochondrial proteins (Stoica et al., 2005). Alterations of mitochondrial phospholipids, consisting of anionic phospholipid and cholesterol decrease, and PC and sphingomyelin increase, occurred in picrotoxin‐induced epileptic rat brain, causing mitochondrial respiratory chain dysfunction (Acharya and Katyare, 2005). There is compelling evidence pointing toward a potentially important link between cholesterol, b‐amyloid, and AD (Lukiw et al., 2005). Cholesterol, a modulator of the biophysical state of the lipid bilayer, modifies the rate of b‐amyloid precursor protein cleavage, thereby regulating cellular production of amyloid b‐peptide (Gibson et al., 2003). In brain, aging and AD disease, characterized by the deposition of amyloid b‐peptide, alterations in sphingolipid and cholesterol metabolism resulted in the accumulation of long‐chain ceramides and cholesterol. In the sequence of events in the pathogenesis of AD, amyloid b‐peptide induces membrane‐associated oxidative stress producing perturbation in ceramide and cholesterol metabolism which, in turn, triggers a neurodegenerative cascade (Cutler et al., 2004). Mitochondrial neurosteroids affect neuronal growth and differentiation and modulate neurotransmitter receptors. Disordered cellular cholesterol trafficking, particularly toward the inner mitochondrial membrane, site of neurosteroid synthesis, may alter neurosteroidogenesis. In the brain of Niemann–Pick type C mouse, an experimental model recapitulating all the defects of the most severe forms of human disease, including a defect in cholesterol trafficking, reduced amounts of neurosteroids, and decreased expression of

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5a‐reductase and 3a‐hydroxysteroid dehydrogenase was observed, compared with wild‐type brain (Griffin et al., 2004). In Niemann–Pick mouse brain and neural cells, the amount of cholesterol within mitochondrial membranes is significantly higher. In addition, the mitochondrial membrane potential, the activity of ATP synthase, and the level of ATP are markedly decreased, indicating that altered cholesterol metabolism affects mitochondrial function (Yu et al., 2005).

4

Conclusion

Lipids of neural mitochondria have been reviewed. Since mitochondria are not able to synthesize all the lipids they contain, processes of lipid translocation from the endoplasmic reticulum to mitochondria have also been described. Most of the experiments concerning traffic routes of lipids have been performed in vitro, while their relevance in vivo remains uncertain. Likewise, the molecular mechanisms underlying the supply of lipids to mitochondria in CNS remain unclear. Here, we review data suggesting that the most likely mechanism of lipid import is mediated by proteins acting through membrane contacts. Although the mechanism of lipid transfer between MAM and mitochondria is yet unknown, several studies have suggested that proteins on the surface of one or both membranes participate in this process. Evidence has also been provided to suggest that proteins with fusogenic properties participate in lipid transfer reaction, although none of these components has been clearly identified. Diffusion mechanisms might have also been involved in the translocation of the less ‘‘lipophilic’’ lipids such as PA and lysophospholipids. There is no evidence that lipids are supplied to brain mitochondria by vesicle flow. In addition, little information is available on how lipids are exported from mitochondria, particularly when the process occurs in the brain. Many points need to be clarified concerning the mechanism of cholesterol homeostasis in the brain and how cholesterol is taken up by mitochondria. Increasing information is emerging that lipoprotein metabolism in the brain is important to normal functioning of neurons. The molecular mechanisms underlying intracellular lipid movement in brain are yet to be acknowledged. In vivo, more than one mechanism of lipid transport might function concurrently to have the regulatory mechanisms supply the correct amount of lipids to mitochondria. Open questions concern the mechanisms that coordinate extramitochondrial and intramitochondrial lipid synthesis. Moreover, how lipid synthesis influences the transport between organelles is still unclear. It would be essential to know how the stationary state of lipids in mitochondria is maintained and how lipid synthesis and degradation are regulated. Recent developments using a genetic approach in yeast have clarified which genes participate in the intracellular lipid movement. Finally, provided pure mitochondria preparations are readily available, mass spectrometry will represent a powerful technique to be used on lipidomics of mitochondria.

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Neuronal Membrane Lipids – Their Role in the Synaptic Vesicle Cycle

L. Lim . M. R. Wenk

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 224

2 Roles of Membrane Lipids in Synaptic Vesicle Recycling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 228 2.1 Glycerophospholipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 228 2.1.1 Phosphatidylinositol (PI) and its Phosphorylated Derivatives (Phosphoinositides, PIs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 228 2.1.2 Phosphatidylserine (PS) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 230 2.1.3 Phosphatidylethanolamine (PE) and Anandamides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 230 2.1.4 Phosphatidylcholine (PC) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231 2.2 Sterol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 232 2.2.1 Cholesterol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 232 2.2.2 Oxysterols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 232 2.3 Sphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 2.3.1 Ceramide and Glycosylated Ceramides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 2.3.2 Sphingomyelin (SM) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 2.3.3 Sulfated Ceramides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 2.3.4 Sphingolipids and Cholesterol in Lipid Microdomains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 234 3

Future Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 234

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_9, # Springer ScienceþBusiness Media, LLC 2009

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Neuronal membrane lipids – their role in the synaptic vesicle cycle

Abstract: The synaptic vesicle cycle requires a stringent interplay between many entities, traversing through temporal and spatial coordination in order to ensure successful and sustainable neurotransmission. Adding to this complexity of coordination, the complete cycle of neuronal synapse is rapid, estimated at 1 min. As early as the 1960s, various studies have taken on the task to characterize the compositions of synaptic vesicles, identifying both lipids and proteins. Clearly, even till now, there have been extensive debates over how the synaptic vesicle (SV) cycle occurs, which led to various imaginable models, loosely divided into clathrin-dependent and -independent pathways. There have also been many lines of evidence to support the existence and relevance of each model, continuously adding to the wealth of knowledge in identifying the roles of various proteins in the SV cycle. However, less is known about the roles of lipids. While the most and best studied lipids are glycerophospholipids, in particular phosphorylated forms of glycerophosphatidylinositol, the phosphoinositides, we still do not know if and how other lipids, such as cholesterol and sphingolipids, regulate the SV cycle. This chapter will therefore focus on our current understanding of lipid involvement in the SV cycle. We will review the main classes of lipids found in SV membranes and discuss their functions in the context of the SV cycle. List of Abbreviations: AA, arachidonic acid; ACh, acetyl-choline; Cer, ceramide; Chol, cholesterol; DAG, diacylglycerol; DHA, docosahexaenoic acid; Ins(1,4,5), inositol (1,4,5) triphosphate; LPC, lysophosphatidylcholine; NAE, N-acylethanolamine; NAPE, N-acylphosphatidylethanolamine; NPC, niemann-pick disease type C; PA, phosphatic acid; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PI, phosphatidylinositol; PI(4,5)P2, phosphatidylinositol-4,5-bisphosphate PI(4,5)P2; PIs, phosphoinositides; PLD, phospholipase; pPC, plasmenylcholine; pPE, plasmenylethanolamine; PS, phosphatidylserine; SM, sphingomyelin; SMase, sphingomyelinase; SV, synaptic vesicle

1

Introduction

While about 50% of the brain’s dry weight is lipid, neuroscientists still have a limited explanation to this observation. Perhaps the simplest explanation for a high lipid composition is that it is due to white matter, which is composed largely of myelin (> Figure 9-1). In neuronal architecture, myelin is a type of specialized membrane that acts as an electrical insulator, wrapping around axons of neurons to create higher resistance on the axonal membrane. It is composed mainly of cholesterol, glycerophospholipids, sphingomyelin, sulfatide, and ethanolamine plasmalogen (Breckenridge et al., 1972, 1973; Deutsch and Kelly, 1981; DeVries et al., 1981; Fedorow et al., 2006; Takamori et al., 2006; Simons and Trotter, 2007).

. Figure 9-1 Neuronal cell architecture and the neurological synapse. (a) Cartoon depiction of neuron, axon, and synapse with another neuron. Oligodendrocytes producing myelin act as electrical insulators are shown as ovals. (b) Enlarged area of a synaptic contact between two neighboring neurons ( dotted box in Figure 9.1a)

Neuronal membrane lipids – their role in the synaptic vesicle cycle

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. Figure 9-2 Lipid biochemistry of synaptic vesicles. The wheel chart represents the relative levels of different lipid classes found in synaptic vesicles. These values are compilations of data reported from independent studies in separate laboratories (Breckenridge et al., 1972, 1973; Takamori et al., 2006)

However, beyond acting as insulators, lipids also play important regulatory roles in neurological synapses. The fundamental properties of the SV cycle require the fusion event of two lipid membranes. In Heuser and Reese’s original model of the SV cycle, vesicles fully fuse, collapse, recycle by clathrinmediated endocytosis, and reenter as new SVs via endosomal sorting. At each point, this would require extensive lipid–lipid and lipid–protein interactions. It is in such a context that we shall discuss the known roles of glycerophospholipids, cholesterol, and sphingolipids, which together make up the bulk of biological membranes including those found in neurons and synaptic vesicles (> Figure 9-2). As the SV cycle progresses from one stage to the next, lipid distributions are reshuffled to allow for fusion and subsequent budding and re-formation of new SVs. Intracellular communication between the plasma membrane and synaptic vesicles must be achieved along side with structural changes. Different classes of lipids act as signaling molecules to recruit proteins, play important roles in membrane fluidity, and maintain the integrity of synaptic vesicle structure and architecture (see > Table 9-1 for summary). It is unclear as to how the tight size distribution of SVs (with diameters ranging from 35 to 50 nm) is achieved; but it is likely that both lipids and SV proteins play supportive roles in achieving this. For example, mutations in FAT-3, a gene which encodes for a fatty acyl desaturase in C. elegans, lead to reduced numbers of SVs and compromised neurosecretion (Lesa et al., 2003). Thus, in order for the SV cycle to be successful, various species of lipids and proteins must cooperate, communicate, and interact with each other. In addition, and unlike the general endo- and exocytosis in other cell types, synapse require not only speed but also consistency and tight coupling. If there is any misregulation, this could compromise transmission of neuronal signals. Phenotypes of a compromised synaptic vesicle cycle may include impaired cognition, mental retardation, and neurodegeneration.

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Ins(1,4,5) P3 DAG

PS

PE

PC

Inositol (1,4,5) triphosphate Diacylglycerol

Phosphatidylserine

Phosphatidylethanolamine

Phosphatidylcholines

Lipid name Phosphatidylinositol-4,5bisphosphate

Common Abbrev. PI(4,5)P2

Membrane structure, choline homostatsis, ACh precursor

C2 domain dependent protein recruitment, electrostatic interaction, DHA storage Membrane fluidity, AA storage

SV priming

SV priming, Ca2+ levels

Role(s) in synaptic vesicle cycle SV protein recruitment, SV recycling

Plasma membrane (hydrophobic region), Plasma membrane (inner leaflet), SV membrane Plasma membrane (inner leaflet), SV membrane Plasma membrane (outer leaflet), SV membrane

Location (mainly) Plasma membrane (inner leaflet), SV membrane, Cytosol

Schizophrenia, progressive epilepsy with mental retardation Binding to snake venom phospholipase A2, Alzheimer’s disease

Alzheimer’s disease

Examples of association with neuronal diseases/ toxicity Various reported: manic depression, schizophrenia, Alzheimer’s disease, etc.

Li et al. (2006), Vance et al. (2007), Li and Vance (2008)

Michaelson et al. (1983), Kim et al. (2004), Glomset (2006), Yeung et al. (2008) Hansen et al. (1998), Glomset (2006), Okamoto et al. (2007)

Reference Cremona et al. (1999), Di Paolo et al. (2004), Lee et al. (2004), Wenk and De Camilli (2004), Milosevic et al. (2005), Nakano-Kobayashi et al. (2007)

9

. Table 9-1 Summary of lipids and their roles in synaptic vesicle cycle

226 Neuronal membrane lipids – their role in the synaptic vesicle cycle

Cer

SM

Ceramide

Sphingomyelin

Sulfatides

Chol

Cholesterol

Unclear

Membrane structure via ceramide backbone

SV formation, fusion, structure, and membrane fluidity, signaling

Membrane fluidity

Metachromatic leukodystrophy

Alzheimer’s disease, metachromatic leukodystrophy

Brodin et al. (2000), Buccoliero and Futerman (2003), Futerman and Riezman (2005), van EchtenDeckert and Herget (2006), Chen et al. (2007), Fernandis and Wenk (2007) Han (2004, 2005)

Farber’s, Sandhoff, Gaucher diseases

Cytosol, plasma membrane (outer leaflet), SV membrane Cell plasma membrane (outer leaflet), SV membrane

Myelin membrane

Pellkofer and Sandhoff (1980), Deutsch and Kelly (1981), Goritz et al. (2002), Vance et al. (2006), Gylys et al. (2007), Wasser et al. (2007) Carrer et al. (2003), Rohrbough et al. (2004), Snook et al. (2006)

Niemann–Pick disease, metachromatic leukodystrophy

SV and cell plasma membrane (inner and outer leaflet)

Neuronal membrane lipids – their role in the synaptic vesicle cycle

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Neuronal membrane lipids – their role in the synaptic vesicle cycle

Roles of Membrane Lipids in Synaptic Vesicle Recycling

2.1 Glycerophospholipids 2.1.1 Phosphatidylinositol (PI) and its Phosphorylated Derivatives (Phosphoinositides, PIs) In mammalian cells, seven isomeric species of PIs, are currently known which are generated by phosphorylation processes on the inositol headgroup. This group of glycerophospholipids is rather low in abundance in most cell types, but heavily enriched in the brain (Rana and Hokin, 1990; Costa, 1994; Pacheco and Jope, 1996). Due to their important roles in signaling, they are one of the most extensively studied classes of lipids (for recent review Di Paolo and De Camilli, 2006). In the synaptic vesicle cycle, where rapid membrane fusions involve protein recruitment and restructuring of the cytoskeleton, the generation and catabolism of specific PIs at appropriate stages seem essential. At the plasma membrane of a resting neuron, most of the PIs exist as phosphatidylinositol-4,5bisphosphate (PI(4,5)P2, > Figure 9-3a). Many neuronal proteins bind to the inositol-phosphate headgroup of PI(4,5)P2, including clathrin adaptor proteins AP2, AP180, dynamin, epsin, Hip/Hip1R ARH/ Dab (see recent review Rohrbough and Broadie, 2005). In the classic phospholipase C-mediated signal transduction cascade, ‘‘metabolites’’ generated from PI (4,5)P2 are inositol(1,4,5)-tri-phosphate [Ins(1,4,5)P3] and diacylglycerol (DAG). Ins(1,4,5)P3 will cause an increase in intracellular Ca2+ (Berridge, 1984). For SV vesicles to be ‘‘primed’’ or ‘‘directed’’ to the presynaptic membrane, DAG recruits UNC13, or its mammalian homolog MUNC-13, to the presynaptic membrane, which allows for the commencement of the SV cycle (Rhee et al., 2002). To turn off the signal, DAG kinase (DGK) phosphorylates DAG to generate phosphatidic acid, which is subsequently converted to other lipids including PI(4,5)P2. Therefore, DGK is considered to be a regulator of MUNC-13 activity via the attenuation of DAG levels (Goto and Kondo, 1999). After SVs are released, they need to be recycled or endocytosed for the SV cycle to continue. Endocytosis by clathrin/AP2 dependent-pathway is controlled by the generation of PI(4,5)P2 at the nerve terminal by phosphatidylinositol phosphate kinase type I gamma (PIP5KIgamma) (Wenk et al., 2001; Krauss et al., 2003; Di Paolo et al., 2004; Di Paolo and De Camilli, 2006; Nakano-Kobayashi et al., 2007). One proposed mechanism is that AP2 translocates to the plasma membrane by recognition and interaction of (1) a receptor containing tyrosine and acidic dileucine motif and (2) the inositol headgroup of PI(4,5)P2. This

. Figure 9-3 Structures of various classes of lipids found in synaptic vesicles. Structures of various classes of lipids discussed in this chapter. (a) The headgroup of phosphatidylinositol-4,5 bisphoshate [PI(4,5)P2] binds to various neuronal proteins including adaptor and accessory factors of the clathrin coat. Metabolites from PI(4,5)P2 (e.g., Ins(1,4,5) P3 and DAG) function in neuronal signaling. (b, c) Glycerophosphatidylethanolamine (b) with ester-linked fatty acyls (b) and a plasmalogen (c) are found in high levels in brain. A major fatty acyl in brain is arachidonic acid (AA, 20:4), an omega-6 fatty acid. AA is a precursor to signal molecules such as prostaglandins. (d) Phosphatidylserine is an abundant phospholipid in neuronal and SV membranes. Its headgroup binds to C2 domains of many proteins (e.g., synaptotagmin). The major fatty acyl chain in brain PS is docosahexaenoic acid (DHA) (22:6), with double bonds starting at the omega-3 position (shown here). (e) Phosphatidylcholine is a very abundant lipid in many cells including neurons. Metabolism of PC at the headgroup via phospholipase D (PLD) leads to phosphatidic acid, a potent signaling phospholipid that activates lipid kinases. (f, g) Cholesterol (f) and an oxidized derivative (oxysterol, g) can influence membrane structure and fluidity. The oxysterol shown has an addition hydroxyl group at the C7 ring position. (h, i, j) Ceramide (h) forms the backbone for a large class of chemically diverse sphingolipids. Sphingomyelin (i), a ceramide which carries a choline headgroup. More complex glycosphingolipids are extremely abundant in brain and myelin. Sulfatide (j) is a complex glycosphingolipid that has a sulfate group on the 3-OH position of the galactose

Neuronal membrane lipids – their role in the synaptic vesicle cycle . Figure 9-3 (continued)

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requirement of both the membrane receptor and PI(4,5)P2 lead to the model of ‘‘dual-key recognition’’ or ‘‘coincidence detection’’ (for recent review Wenk and De Camilli, 2004). Since the SV cycle is a rapid and dynamic process, it is not surprising to see that some proteins, such as AP2, require intact PI(4,5)P2 while others, such as MUNC-13, require ‘‘breakdown’’ of PI(4,5)P2. For example, synaptojanin 1 is a neuronal polyphosphoinositide phosphatase, which breaks down PI(4,5)P2. Synaptojanin 1-deficient mice exhibit impaired synaptic functions and have neurological defects, along with increased levels of PI(4,5)P2 (Cremona et al., 1999). On the other hand, PIP5KIgamma, a major neuronal kinase, is found at the synapse to generate PI(4,5)P2. This enzyme is required to promote the recruitment of clathrin/AP2 coats (Wenk et al., 2001). Therefore, a tight regulation of PIs levels, and in particular PI(4,5)P2, is indispensable for synapse function.

2.1.2 Phosphatidylserine (PS) Phosphatidylserine (PS) is normally distributed on the inner leaflet of the plasma membrane (Jones and Rumsby, 1977; Kuypers, 1998; Yeung et al., 2008). Translocation to the outer leaflet of the membrane typically triggers a cascade of signaling events and often is accompanied by cell death via apoptosis (Voelker, 1988). In the brain, it is estimated that maintenance of this asymmetrical distribution employs about 20% of net brain ATP consumption (Purdon et al., 2002). Yet, the exact reason for this asymmetry is not entirely understood. Most likely, since PS is an anionic lipid facing the cytoplasm, it is believed to play important roles in electrostatic recruitment of various proteins in signal transduction and the SV cycle. At physiological pH, PS carries one negative charge, whereby it functions as an electrostatic mediator and also binds to the C2 domains of many proteins. Increasing evidence has shown that while modification of proteins, such as myristoylation, is necessary to recruit various G proteins and MARCKS to the membrane, it is not sufficient for these proteins to function (McLaughlin and Aderem, 1995; Murray et al., 1997, 2001). In the context of the SV cycle, synaptotagmin I, a synaptic vesicle membrane protein, is also a Ca2+ sensor with two C2 domains that functions only upon binding to the anionic charge of the headgroups found in PIs and PS (Shahin et al., 2008). Similarly, protein kinase C alpha, whose activity is Ca2+ dependent, can only bind Ca2+ upon specific binding to PS (Ochoa et al., 2002; Mozzi et al., 2003). The intrinsic binding of proteins to divalent cations, in particular Ca2+, tends to be of low affinity and noncooperatively. However, upon binding to negatively charged phospholipid membranes – specifically PS, overall affinity towards Ca2+ is enhanced in a cooperative manner (Ochoa et al., 2002). Other than recruiting proteins by electrostatic interactions, PS may also act as a storage buffer for bioactive fatty acids such as docosahexaenoic acid (DHA). DHA is obtained largely through the diet (e.g., from fish oil or in omega-3 fatty acids such as linolenic acid found in soybeans, which is later synthesized into DHA by the liver). DHA crosses the blood–brain barrier and is immediately esterified to membrane glycerophospholipids such as PS. Uniquely, PS levels can increase upon DHA enrichment. This is, however, not a universal mechanism, as it is only specific to neuronal cells (Guo et al., 2007). It is important to note that de novo synthesis of PS does not occur in animal cells but through the exchange of serine base by the enzymes PS synthase 1 and 2 (PSS1 and PSS2). It has been shown that PSS1 and PSS2 have substrate preference for DHA-containing phospholipids (Kim et al., 2004; Kim, 2007). The reason for such a preference is not clear. However, the relative ratio of fatty acyl chain with omega-3 (such as DHA) to omega-6 (such as arachidonic acid AA) in PS compared to phosphatidylethanolamine (PE) in brain is about 3 times higher in PS than that of PE (Svennerholm, 1968). Therefore, one of the possible roles for PS is that it acts as a buffer for DHA release and/or storage (Salem et al., 2001). Interestingly, while DHA is not a precursor for prostaglandins (an important class of signaling hormones in neurons), levels of DHA directly control its release (Strokin et al., 2007).

2.1.3 Phosphatidylethanolamine (PE) and Anandamides Phosphatidylethanolamine (PE), like PS, resides mostly in the inner leaflet of the plasma membrane (Kuypers, 1998). However, unlike PS, which has a relatively high level of DHA as its fatty acyl chain, PE

Neuronal membrane lipids – their role in the synaptic vesicle cycle

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is enriched in arachidonic acid (AA) (Martinez and Mougan, 1998). In brain PE (as well as PC, see below) ether linkages are common and particularly abundant (Andre et al., 2006), and they represent between half to two-thirds of PE (Nagan and Zoeller, 2001; Andre et al., 2006). Ether lipids can be classified into plasmanyl or plasmenyl, corresponding to the alkyl (C–O–C–C) or alkenyl (C–O–C=C) forms respectively. Specifically, plasmenylethanolamine or ethanolamine plasmalogen (pPE) is tenfold more abundant than plasmenylcholine (pPC) (Farooqui and Horrocks, 2001). One of the reasons for the abundance in pPE could be to facilitate membrane fusion. This is because at physiologically relevant temperatures, pPE forms inverse hexagonal phases, while the diacyl analog does not (Lohner, 1996). This means that there is an increase in membrane leakage, promoting membrane–membrane fusion. Although synthetic vesicles containing 45–50% of pPE fuse much more rapidly than those with the diacyl counterparts, there is no direct evidence that pPE is the determining factor of membrane fusion during the SV cycle. Additionally, the pPE content in SV is not so high (> Figure 9-2): a maximum of 15–20% on biological membranes. However, this is not to say that pPE does not contribute to the membrane fluidity, as pPE deficient cells show compromised membrane dynamics (Glaser and Gross, 1995). Other than membrane integrity, much like PS, which could function as storage for DHA, pPE and PE could also act as storage for AA, the precursors to anandamides and prostaglandins. N-Arachidonoylethanolamine, or anandamide, belongs to a class of bioactive long-chain N-acylethanolamines (NAEs), and is also a ligand to the endocannabinoid receptors. NAEs are formed by two enzymatic reactions: (1) N-acylation of PE to generate N-acylphosphatidylethanolamine (NAPE); (2) cleavage of NAPE by a phosphodiesterase to form NAE (Jin et al., 2007; Okamoto et al., 2007). During the SV cycle, the endogenous endocannabinoid system can inhibit neuronal synapses of excitatory neurons. This involves communication between the post- and pre-synaptic membranes. First, release of anandamide from the post-synaptic membrane will retrograde to the cannabinoid receptors (CB1) on the pre-synaptic membrane. Upon binding, activation of CB1 will inhibit N-type Ca2+ channel activity, which in turn reduces glutamate release (Huang et al., 2001; Melis et al., 2004). Furthermore, NAE derivatives, such as N-ethylmaleimide (NEM), have also been shown to inhibit N-type Ca2+ channels in sympathetic neurons (Shapiro et al., 1994). How exactly is this process coordinated has not yet been fully elucidated.

2.1.4 Phosphatidylcholine (PC) PC is a very abundant lipid in many cells and constitutes up to a third or more of membrane lipids (Li and Vance, 2008; van Meer et al., 2008). Thus, it has important roles in maintaining membrane structure. Many lipids have spontaneous curvature, categorized on the basis of their preference of curvature when placed in a monolayer. For example, if a lipid tends to bulge spontaneously towards the polar heads, it is considered to have an inverted-cone shape, with a positive curvature. If it bulges towards the hydrocarbon tail, then it has a cone shape, with a negative curvature. If it remains fully neutral in a monolayer, then it is considered cylindrical. PC is considered cylindrical, whereas PE inverted-cone shaped (Chernomordik and Kozlov, 2003; Wenk and De Camilli, 2004; Chernomordik et al., 2006). Changing the membrane shape can be achieved by either liberation of the headgroup of PC to generate phosphatidic acid (PA) (e.g., by PLD), or by liberation of the fatty acyl chains (e.g., by PLAs) to generate lysophosphatidylcholine (LPC). This will thus yield molecules with very different geometrical characteristics, whereby PA has a cone shape and LPC has an inverted-cone shape. Since PC is such an abundant lipid, such conversion, in particular if and when induced locally, may have profound implications for the membrane bilayer structure (Chernomordik and Kozlov, 2003; Piomelli et al., 2007). Other than membrane curvature, PC also has a role in controlling membrane fluidity. Maintaining the ratio of PC to PE levels is also crucial to prevent the membrane from being too leaky (Li et al., 2006; Vance et al., 2007; Li and Vance, 2008). As discussed previously, while high levels of plasmenylethanolamine can influence membrane fluidity, a low PC to PE ratio could also increase fluidity. In mice on choline-restrictive diets lacking the rate-limiting enzyme to synthesize PC, PEMT / , the PC to PE ratio is altered and levels

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of PE are increased (Li et al., 2006). In these mice, the membrane permeability is increased as evidenced by an increase of plasma alanine aminotransferase (an enzyme that is normally found only in the liver) (Li et al., 2006). Finally, PC also has a crucial role in choline homeostasis (see details in Li and Vance’s review 2008). Choline, while being an essential dietary nutrient for various cellular functions, is also toxic, with an upper tolerance limit of 3.5 g/day for humans (Li and Vance, 2008). In a cell, the importance of this balance and the adaptation associated with this imbalance become crucial to survival. To maintain choline homeostasis, a cell can directly do so by controlling the metabolism of membrane phosphatiylcholine. In neurons, choline is stored, after being taken up in the diet, as PC on the plasma membrane. A small percentage could be immediately converted to acetyl-choline (ACh), a neurotransmitter. In the event where choline is not readily available in the brain, the hydrolysis of PC can then give rise to choline for ACh production. However, if there is an inadequate supply of choline over a long term period, this might result in a net loss of membrane PC, followed by an impairment of membrane function and loss of cellular viability (Amenta and Tayebati, 2008). During such choline deprivation, choline can be transported from other tissues’ cellular membrane. In studies where PEMT / mice, which cannot synthesize PC, were deprived of choline in their diets, [3H]-choline labeling showed that choline was redistributed from kidney, lungs, and the intestines to the brain and liver (Li et al., 2005; Li and Vance, 2008).

2.2 Sterol 2.2.1 Cholesterol Uniquely, the brain contains about 25% of the body’s total cholesterol, which is essentially all synthesized independently within the brain. Glial cells produce apolipoprotein E-containing particles to deliver cholesterol to neurons and developing axons (Pfrieger, 2003a, b; Han, 2004; Goritz et al., 2005; Vance et al., 2006). Cholesterol varies in concentration depending on organelle type. In purified SVs, cholesterol levels are comparable to those found in neuronal plasma membranes (25%, > Figure 9-2, Breckenridge et al., 1972, 1973; Deutsch and Kelly, 1981; DeVries et al., 1981). While we know that cholesterol transport and regulation are important for axonal growth and proper function of the brain, its precise role in SV structure and dynamics is not entirely clear. Various groups have found that cholesterol can decrease membrane permeability and can stabilize the synaptic plasma membrane. Pharmacological inhibition of cholesterol synthesis results in a decrease of evoked synaptic transmission (Zamir and Charlton, 2006). More recently, in an elegant study combining cholesterol inhibitors and transgenic mice with defects in cholesterol transport, enhanced spontaneous fusion was observed upon cholesterol depletion (Wasser et al., 2007). Neuronal cholesterol is very important for proper brain development, and its misregulation in adult and aging brains has been associated with several neurological diseases such as Alzheimer disease (Pfrieger, 2003a, b; Puglielli et al., 2003; Karten et al., 2006; Vance et al., 2006). The fatal lipid storage disease, Niemann–Pick disease type C (NPC), is characterized by mutations in the protein that encodes for transport of cholesterol (Ribeiro et al., 2001; Karten et al., 2006).

2.2.2 Oxysterols Cholesterol (> Figure 9-3f ) can be oxidized at various positions, e.g., the C7 position (> Figure 9-3g) or the C24 position, to generate oxysterols. In many disease states, the generation of oxysterols can induce a complex mode of cell death exhibiting some characteristics of apoptosis. Structurally, these oxidized sterols may appear very similar to cholesterol, but it is clear that metabolism and clearance follow different routes. Formation of oxysterol is seen in many disease states such as Alzheimer’s. Thus, oxysterols might act in part by direct physical compromise of membrane fluidity, which thus affects synaptic function (Sagin and Sozmen, 2008).

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9

2.3 Sphingolipids Sphingolipids – ceramide (> Figure 9-3h), sphingomyelin (> Figure 9-3i), gangliosides, and their metabolites – have been implicated in structural and signaling roles of biological and neuronal membranes (Usuki et al., 1996; Buccoliero and Futerman, 2003; Rohrbough et al., 2004; Snook et al., 2006; van EchtenDeckert and Herget, 2006; Morales et al., 2007).

2.3.1 Ceramide and Glycosylated Ceramides Ceramide is a central molecule in sphingolipid metabolism. While we do not know how ceramide interacts with various SV proteins (Snook et al., 2006), mutants lacking ceramidase – the enzyme which cleaves ceramide to produce ‘‘soluble’’ signals, i.e., fatty acyls and sphingosine – have decreased ability to complete priming or fusion (Rohrbough et al., 2004). These mutants are termed as ‘‘slug-a-bed’’ (SLAB) for their impaired movement in drosophila. It is possible that dysregulation of ceramide levels could affect microdomains mentioned above by perturbation of sphingolipid/cholesterol assemblies (Rohrbough et al., 2004; Tani et al., 2007). Sphingolipids are linked via complex metabolic chains, which in addition to lipids also include carbohydrates. Thus, it is difficult to assess individual roles of ceramides, sphingomyelins, or gangliosides (for reviews of sphingolipid metabolism, see Buccoliero and Futerman, 2003; Futerman and Riezman, 2005; van Echten-Deckert and Herget, 2006). In vitro studies have found that a certain balance of ceramide, sphingomyelin, and gangliosides is required for small vesicle size and membrane integrity (Haque et al., 2001; Carrer et al., 2003). As an example, it has been long known that halothane and other anesthetics elevate membrane fluidity. This is accomplished by the breakdown of sphingomyelin and ganglioside (Pellkofer and Sandhoff, 1980). It is possible that such degradation products act as signaling molecules in the modulation of membrane fluidity (Sandhoff and Pallmann, 1978; Pellkofer and Sandhoff, 1980).

2.3.2 Sphingomyelin (SM) Sphingomyelins are enriched in myelin. They are typically found on the outer leaflet of plasma membranes. SM can be hydrolyzed to ceramide and choline by sphingomyelinases (SMase), which are classified into ‘‘acidic’’ and ‘‘neutral’’ forms according to their enzymatic characteristics. In cell-free systems, SMase activity has been shown to drive structural reorganization of membrane lipids (Fanani et al., 2002; Mattjus et al., 2002). In cells, disruption of SM leads to inhibition of Ca2+-triggered membrane fusion (Rogasevskaia and Coorssen, 2006). The roles of SMases in neurons are not overly well understood, in particular with respect to normal physiology. Acid SMases are involved in intracellular sphingolipid transport (see above). Neutral SMase2 (nSMAase2) is one of the two known SMAse forms and is primarily expressed in brain. It is found (at least in some cell types) at the periphery (Marchesini et al., 2004). SMase2 plays an important role in late embryonic and postnatal development. Disruption of its function results in a form a dwarfism (Stoffel et al., 2005).

2.3.3 Sulfated Ceramides In brain, a number of glycosphingolipids carry a sulfate group on the carbohydrate headgroup. ‘‘Sulfatides’’ are a class sphinglipids with a sulfate on the 3-OH position of the galactose of galactosylceramide (> Figure 9-3j, cerebroside sulfatide). Like SM, these lipids are highly enriched in myelin and are not very abundant in other tissues. These lipids appear to be important for cell–cell contact, but little is known about the potential mechanisms during SV formation and recycling. Abnormal sulfatide metabolism has been associated with many diseases such as Alzheimer’s disease, multiple sclerosis, and metachromatic leukodystrophy, which is caused dysfunction on arylsulfatase A, a sulfatide-degrading enzyme. In such neurons, there is a prolonged and reoccurring spontaneous hyperexcitability, with recurrent discharge lasting from 5 to 15 s (Eckhardt et al., 2007). Sulfatides have been

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successfully used as biomarkers for neurodegeneration since the levels are seen increased in cerebral fluid as well and in brain of postmortem patients with Alzhiemer’s disease (Han et al., 2002; Han, 2005). Antibodies against sulfatide have also been shown to increase in serum of patients with chronic immune-induced neuropathies (Nemni et al., 1994) (Kanter et al., 2006).

2.3.4 Sphingolipids and Cholesterol in Lipid Microdomains Sphingolipids and cholesterol are often mentioned in the context of detergent-resistant complexes. In biological membranes, lipid microdomains play important roles in many cellular processes including signal transduction, membrane trafficking, cytoskeletal organization, and pathogen entry (Munro, 2003; Kenworthy et al., 2004). Microdomains are thought to be transient in nature and their formation dependent on membrane lipid composition including cholesterol (Xu et al., 2001; Munro, 2003; Kenworthy et al., 2004; Gylys et al., 2007; Matsuura et al., 2007).

3

Future Perspectives

The SV cycle has been extensively studied for over four decades. Progress in ‘‘lipidomics,’’ however, is still lagging behind proteomics and genomics. In summary, there are many experimental evidences supporting the critical role of PIs during the SV cycle, in particular with respect to their association and recruitment of proteins to membrane surfaces. Much less, however, is known about the roles of other glycerophospholipids, cholesterol, ceramides, sphingomyelin, and sulfatides, even though their roles are indispensable in proper neuronal functions. We have guided our overview based on biochemical determination of lipids in SVs. However, we have not addressed the important issues of asymmetrical transbilayer distribution of lipids (Glomset, 2006). Based on biophysical considerations a vesicle of 35–50 nm in diameter will harbor 60% of its lipids in the other leaflet. Thus, it is likely that in SV fatty acyl composition, charge distribution, and protein–lipid interactions play important roles in determining asymmetry. How does maintenance or misregulation of lipids in SV affect the SV cycle? We have also not addressed, in detail, oxidized fatty acids and endocannabinoids and their potential role in the SV cycle. These lipids are emerging as potent regulators of very diverse cellular and physiological functions (Freund et al., 2003). Most importantly, we have yet to answer or elucidate how any misregulation of lipids during SV could lead to perturbations of SV cycle, and subsequently be linked to disease states such as psychiatric disorders, metal retardation, and neurodegenerative diseases. It is clear that many neuron-related diseases, such as schizophrenia, Krabbe’s disease, Farber’s disease, Sandhoff/Tay Sach disease, multiple sclerosis, Guillain–Barre syndrome, Alzheimer’s disease, are implicated by misregulation of lipids (> Table 9-1). Most recently, in the context of Alzheimer’s disease, neurons with oligomeric amyloid-beta peptide have been shown to display altered phosphatidylinositol-4,5-bisphosphate metabolism (Berman et al., 2008). Such molecular studies with a focus on lipids that play prominent roles in the SV cycle are likely to provide answers to our questions in the future. Therefore, the challenge for future research of neural lipids is not just teasing out the complexity or diverse roles of lipids, but also how we combine the analysis of lipids in neurons and the SV cycle to genetics, proteomics, and its relevance to medicine. To do so, we have to start with a basic understanding of neuronal cell types and organelle-specific lipids, their diversities, functions, and interactions with other proteins. Beyond that, the subsequent link towards proteomics, genetics, and medicine would take a due course.

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Functional Dynamics of Myelin Lipids*

S. N. Fewou . N. Jackman . G. van Meer . R. Bansal . S. E. Pfeiffer

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 240

2 2.1 2.1.1 2.2 2.3 2.3.1 2.3.2

Lipid Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241 Sphingolipid Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241 Ganglioside Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 242 Cholesterol Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 245 Phospholipid Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 246 Phosphosphingolipid Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 246 Glycerophospholipid Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 247

3 3.1

Sorting and Transport of Lipids During Myelin Assembly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 248 Lipid Transport in Polarized Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 249

4 4.1 4.2

Role of Lipids in the Regulation of Protein Sorting and Transport . . . . . . . . . . . . . . . . . . . . . . . . . . 250 Fatty Acids and the Regulation of Myelin Protein Sorting and Transport . . . . . . . . . . . . . . . . . . . . . . 250 Sphingolipids/Cholesterol and the Regulation of Intracellular Transport . . . . . . . . . . . . . . . . . . . . . . . 250

5

Role of Lipids in the Biogenesis and Maintenance of Myelin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 251

6 6.1 6.2 6.3

Role of Lipids in the Regulation of Oligodendrocyte Physiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 253 Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 253 Sphingolipids and the Regulation of Oligodendrocyte Lineage Progression . . . . . . . . . . . . . . . . . . . . 253 Gangliosides and Oligodendrocyte Physiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255

7

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 256

*This book chapter is dedicated to the memory of Prof. Steven E. Pfeiffer (1940–2007). May every reader of this chapter remember him as one of the scientists who largely contributed to the comprehension of the myelin physiology and oligodendrocyte development. G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_10, # Springer ScienceþBusiness Media, LLC 2009

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Functional dynamics of myelin lipids

Abstract: Biological membranes of living organism are composed of two fundamental components: proteins and lipids. Lipids are defined as water-insoluble biomolecules, which have high solubility in nonpolar organic solvents. They account for more than half of the total mass of myelin, which is an extension of oligodendrocyte plasma membrane that spirally enwraps axons and is critical for efficient nerve conduction. Because of the high lipid content of myelin, in particular glycosphingolipids and cholesterol, it was thought to play a central role in myelin/oligodendrocyte physiology. This view has been strongly supported by multiple approaches, most prominently the gene knockout studies that have significantly enhanced our understanding and appreciation of lipids in the overall function and structure of the CNS myelin. This chapter discusses the role of lipids in the regulation of myelin/oligodendrocyte physiology including oligodendrocyte development, myelin biogenesis and maintenance, and sorting and transport of myelin components. List of Abbreviations: ABC, ATP-binding cassette; CNP, 20 ,30 -cyclic nucleotide 30 -phosphodiesterase; CNS, central nervous system; CST, cerebroside sulfotransferase; DHAP, dihydroxyacetone-phosphate; GalCer, galactosylceramide; GD1a, Neu5Aca3Galb3GalNAcb4-(Neu5Aca3)Galb4GlcCer; GD1b, Galb3GalNAcb4(Neu5Aca8Neu5Aca3)Galb4-GlcCer; GD2, GalNAcb4-(Neu5Aca8Neu5Aca3)Galb4GlcCer; GD3, Neu5Aca8Neu5Aca3Galb4GlcCer; GlcCer, glucosylceramide; GM1a, Galb3GalNAcb4(Neu5Aca3) Galb4GlcCer; GM1b, Neu5Aca3Galb3GalNAcb4Galb4GlcCer; GM2, GalNAcb4(Neu5Aca3)Galb4GlcCer; GM3, Neu5Aca3Galb4GlcCer; GM4, N-acetylneuraminylgalactosylceramide; GQ1b, Neu5Aca8Neu5Aca3Galb3GalNAcb4(Neu5Aca8Neu5Aca3)Galb4GlcCer; Gro-3P, glycerol-3-phosphate; GT1a, Neu5Aca8Neu5Aca3Galb3GalNAcb4(Neu5Aca3)Galb4GlcCer; GT1b, Neu5Aca3-Galb3GalNAcb4(Neu5Aca8Neu5Aca3) Galb4GlcCer; GT1c, Galb3GalNAcb4(Neu5Aca8Neu5Aca8Neu5Aca3)Galb4GlcCer; HMG-CoA, 3-hydroxy-3-methylglutaryl-CoA; MAG, myelin-associated glycoprotein; MBP, myelin basic protein; MDR, multidrug resistant protein; MGDG, monogalactosyl-diacylglycerol; MOG, myelin oligodendrocyte glycoprotein; NeuAc, N-acetylneuraminic acid; PLP, proteolipid protein; PNS, peripheral nervous system; SialT, sialyltransferase; SPTLCB, serine palmitoyltransferase long-chain base

1

Introduction

By means of light microscopy, the pathologist Rudolf Virchow (1854) found that the axon of the nerve fibers was surrounded by a substance to which he gave the name ‘‘myelin.’’ A breakthrough in the understanding of myelin was the observation by Ranvier (1878) that myelin forms a covering of the nerve that is periodically interrupted at regular spacings along the nerve. The constrictions in the nerve fiber that separate two internodal regions now carry his name, nodes of Ranvier, and were found in the first half of the twentieth century to allow the saltatory conduction of the nerve impulse (Rosenbluth, 1999). Studies with polarized light, X-ray diffraction, and electron microscopy have shown that, in both the central and peripheral nervous systems (CNS and PNS, respectively), myelin is made up of regular concentric lamellae appearing as alternating dark and less dark lines separated by lipid hydrocarbon chains that appear as unstained zones (> Figure 10‐1). One of the biochemical characteristics that distinguish myelin from other biological membranes is its high lipid-to-protein ratio. Indeed, the lipid/protein ratio of myelin is 2:1, in contrast to the whole brain, which contains more proteins than lipids (> Table 10‐1). Because of its high lipid content, myelin appears white in the macroscopic view. Therefore, highly myelinated regions of the CNS are called ‘‘white matter,’’ in contrast to the poorly myelinated regions which are called ‘‘grey matter’’ (Stegemeyer and Stegemeyer, 2004). During the last 25 years, important data have been gathered concerning the synthesis and function of lipids in the nervous system, although this field has received little attention compared with that of their protein counterparts. The same period has seen the identification and cloning of cDNAs and genes implicated in the biosynthesis of myelin lipids and proteins, which signaled the beginning of the knockout era: transgenic null-mutant animals have been created for almost every enzyme implicated in the biosynthesis of myelin-enriched lipids. The study of these animals shows that the formation of myelin is considerably less sensitive to the alteration of lipid content than the maintenance of myelin.

Functional dynamics of myelin lipids

10

. Figure 10‐1 The structure of the myelin membrane. (a) Ultrastructure of myelin in the mouse optic nerve. Note that myelin (m) appears dark compared to the axon that appears white. (b) Schematic structure of CNS myelin. Note that the extracellular leaflets of adjacent lamellae become closely apposed to each other to form the intraperiod line (IPL), while the cytoplasmic membrane leaflets fuse to form the major dense line (MDL). The two major proteins (MBP and PLP) of the CNS myelin are depicted. One postulated function for PLP is that it acts like glue to keep the adjacent layers of myelin tightly joined together. The lipid bilayers are shown as plasma membrane. CNP, MAG, MOG, and other minor myelin proteins are not shown. Figure adapted from Quarles et al., 2006 (Courtesy of Dr. Simon Ngamli Fewou)

. Table 10‐1 Lipid composition of the CNS myelin and brain of different species Myelin Componenta Total protein Total lipid Cholesterol Cerebroside Sulfatide Total galactolipids Phospholipids Sphingomyelin

Human 30.0 70.0 27.7 22.7 3.8 27.5 43.1 7.9

White matter Bovine 24.7 75.3 28.1 24.0 3.6 29.3 43.0 7.1

Rat 29.5 70.5 27.3 23.7 7.1 31.5 44.0 3.2

Human 39.0 54.9 27.7 19.8 5.4 26.4 45.9 7.7

Bovine 39.5 55.0 23.6 22.5 5.0 28.6 46.3 6.7

Gray matter (Human) 55.3 32.7 22.0 5.4 1.7 7.3 69.5 6.9

Whole Brain (Rat) 56.9 37.0 23.0 14.6 4.8 21.3 57.6 3.8

a Total protein and lipid figure in percentage dry weight. All others are in percentage total lipid weight. For further quantification, we refer the reader to Quarles et al. (2006)

In this chapter, we review current ideas regarding the lipid biosynthesis, its role in the regulation of protein transport, in myelin biogenesis, and its impact on the regulation of oligodendrocyte (OL) physiology. As further background, we refer the reader to our recent chapter on myelin lipids (Taylor et al., 2004).

2

Lipid Biosynthesis

2.1 Sphingolipid Biosynthesis Sphingolipids are a family of lipids that comprise both structural lipids and a series of highly bioactive compounds that participate in the regulation of cell growth, differentiation, diverse cell functions, and

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apoptosis. They are typically found in high amounts in eukaryotic plasma membranes, and their content is particularly high in apical membranes of epithelial cells (Simons and van Meer, 1988) and in CNS and PNS myelin where they constitute about 30% of total lipids (> Table 10‐1). Structurally, they are composed of a sphingoid base, a straight chain amino alcohol of 18–20 carbon atoms, which normally carries a singular long- or very long-chain fatty acid, saturated or unsaturated at C15, bound to the amino group at the C2 carbon (for review see Holthuis et al., 2001) to form ceramide. Sphingolipids are classified as phosphosphingolipids and glycosphingolipids based on the polar head group on the ceramide backbone. The biosynthesis of sphingolipids starts in the endoplasmic reticulum (ER) with the synthesis of the sphingoid base and ceramide. Serine-palmitoyltransferase (SPT; EC 2.3.1.50) catalyzes the rate-limiting step in de novo synthesis of sphingolipids (Merrill and Jones, 1990), the pyridoxal-50 -phosphate-dependent condensation of L-serine and palmitoyl-CoA to 3-ketosphinganine (Weiss and Stoffel, 1997). Mammalian SPT comprises two homologous proteins, SPT long-chain base 1 (SPTLCB1) and SPT long-chain base 2 (SPTLCB2), which are heterodimers of 53- and 63-kDa subunits, respectively, and both of which are required for its full enzymatic activity (Hanada et al., 1997, 1998, 2000; Yasuda et al., 2003). The conversion of 3-ketosphinganine to the sphingoid base, sphinganine, is catalyzed by the 3-ketosphinganine reductase, an enzyme encoded by the follicular lymphoma variant translocation-1 gene (Kihara and Igarashi, 2004). Sphinganine is then N-acylated by ceramide synthase to form dihydroceramide, which is finally desaturated to form ceramide (> Figure 10‐2). In mammals, ceramide synthase protein is encoded by six members of the ceramide synthase (CerS) gene family also called longevityassurance homologue (Lass) gene family (Pewzner-Jung et al., 2006). Overexpression of any CerS protein in cultured cells results in an increase in cellular ceramide, but the ceramide species produced varies. Overproduction of CerS1 protein increased C18:0-ceramide levels preferentially, and overproduction of CerS2 and CerS4 increased levels of C22:0- and C24:0-ceramides. CerS5 and CerS6 produced shorter ceramide species (C14:0- and C16:0-ceramides); however, only CerS-5 was able to incorporate C18:1-CoA (Mizutani et al., 2005, 2006; Pewzner-Jung et al., 2006). In addition to being implicated in the synthesis of very long chain fatty acid ceramide, CerS2 is specifically expressed in OLs and Schwann cells. Moreover the level of CerS2 in the mouse brain is developmentally upregulated, with a maximum expression level at postnatal day 21 (Becker et al., 2007), which correlate with the peak of myelination. These results might suggest a close relationship between CerS2 expression and myelination. In contrast, CerS1 is specifically expressed in brain, in the cortical region (Becker et al., 2007), while CerS5 and -6 are expressed in brain but also in other tissues (Mizutani et al., 2005). The galactosylation of ceramide in the ER lumen produces galactosylceramide (GalCer) and is catalyzed by UDP-galactose:ceramide galactosyltransferase (CGT), a type 1 integral membrane protein (Sprong et al., 1998). Following its biosynthesis, a fraction of GalCer reaches the lumen of Golgi and is sialylated by the action of sialyltransferase to form N-acetylneuraminyl-galactosylceramide (GM4), or used as a substrate by cerebroside sulfotransferase (CST) with 30 -phosphoadenosine-50 -phosphosulfate (PAPS) to synthesize sulfatide (> Figure 10‐2). CST is a Golgi type II membrane protein that also catalyzes the synthesis of seminolipid and sulfated lactosylceramide (Honke et al., 1996, 1997). GalCer and sulfatide comprise 23% and 4% of the total mass of myelin lipids, respectively, and together account for one-third of the lipid content in the myelin sheath (Norton and Cammer, 1984), and more than half of the GalC in myelin exists as a 2-hydroxy fatty acid containing isoform that is unique to myelin (Schaeren-Wiemers et al., 1995).

2.1.1 Ganglioside Synthesis Gangliosides are sialic acid-containing glycosphingolipids that are known to modulate the activity of a number of receptor tyrosine kinases, including the insulin receptor (Allende and Proia, 2002). Our present knowledge on the mechanism of ganglioside biosynthesis comes mainly from the pioneering studies of Roseman, Brady, and coworkers (Kaufman et al., 1966; Roseman, 1970; Fishman et al., 1972; Basu et al., 1973). These investigators demonstrated that the glycosyl chains of gangliosides are formed in a stepwise manner by the sequential addition of individual sugar and sialyl groups to the growing glycolipids. Gangliosides are synthesized in the lumen of Golgi from lactosylceramide (LacCer) by specific glycosyltransferases and sialyltransferases (> Figure 10‐3) (Kaufman et al., 1968; Basu et al., 1973; Keenan et al., 1974;

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. Figure 10‐2 Pathways of sphingolipid biosynthesis. The biosynthesis of sphingolipids starts in the ER with the condensation of palmitoyl CoA and serine. The enzymes involved in the biosynthesis of ceramide are listed in order as follows: (1) serine palmitoyltransferase, (2) 3-ketosphinganine reductase, (3) dihydroceramide synthase, (4) dihydroceramide desaturase. Ceramide can be converted to sphingomyelin upon the action of (5) sphingomyelin synthase. In addition, conversion of ceramide to ceramide 1-phosphate is catalyzed by ceramide kinase (6). The hydrolysis of ceramide by ceramidase (7) yields sphingosine, which can be either transformed to sphingosine 1-phosphate by the action of sphingosine kinase (8) or galactosylated by the UDP-Galactose:ceramide galactosyltransferase (CGT: 10) to form galactosylsphingosine, a highly toxic molecule in oligodendrocytes. Sphingosine can be also synthesized from sphingosine-1P by the action of sphingosine-1P phosphatase (9). Ceramide can be converted to galactosylceramide (GalCer) by the action of CGT (A), which utilizes both hydroxylated (OH) and nonhydroxylated (n-) fatty acid-containing ceramide. For the synthesis of hydroxyl fatty acid GalCer, the hydroxy fatty acid ceramide is first synthesized by the action of fatty acid 2-hydroxylase (B), an ER resident membrane protein (Alderson et al., 2004; Eckhardt et al., 2005). GalCer can be also sialylated in the Golgi by sialyltransferase (11) to form sialylgalactosylceramide or GM4. Finally, sulfatide is synthesized by using GalCer as substrate, a reaction catalyzed by cerebroside sulfotransferase (CST) (C) (Adapted from Taylor et al., 2004)

Lloyd et al., 1998; Maccioni et al., 1999). The biosynthesis of LacCer starts on the cytosolic face of the Golgi by the transfer of glucose from UDP-glucose to ceramide to form glucosylceramide (GlcCer). This reaction is catalyzed by UDP-glucose:ceramide glucosyltransferase, a type-III transmembrane protein (Futerman and Pagano, 1991; Jeckel et al., 1992; Ichikawa et al., 1996; Paul et al., 1996). Part of the GlcCer then reaches the lumen of Golgi via FAPP2, a GlcCer binding protein associated with the trans Golgi via phosphatidylinositol4-phosphate and ARF (D’Angelo et al., 2007; Halter et al., 2007). Although previous studies on fluorescent GlcCer analogues had suggested that GlcCer is translocated across the Golgi membrane by the multidrug transporter ABCB1 (van Helvoort et al., 1996; Nicholson et al., 1999; Lala et al., 2000; Veldman et al., 2002; Eckford and Sharom, 2005), it was later shown that translocation by ABCB1 was specific for the fluorescent analog and that natural GlcCer flips by an independent mechanism possibly in the ER after retrograde

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. Figure 10‐3 Ganglioside biosynthesis: The entry point in the biosynthetic pathway of gangliosides of all series is the conversion of glucosylceramide to lactosylceramide (precursor of gangliosides of 0-series) by GalT1. LacCer can then be converted to GM3, GD3, and GT3, the precursor of gangliosides of a-, b-, and c-series, respectively. The respective reactions are catalyzed by SialT1, -2, and -3; SialT2 and 3 are possibly the same enzyme (see text). CST catalyzes the conversion of LacCer to SM3, the precursor of all sulfated gangliosides. Knockout mice have been generated for the following enzymes: GlcT (A), SialT1 (B), SialT2 (C), and GalNAcT (D) (Adapted from van Echten and Sandhoff, 1993, Kolter et al. (2002) and Nagai et al. (2005))

transport via FAPP2 (Halter et al., 2007). After the FAPP2-dependent transport to the Golgi lumen, a galactose is transferred onto GlcCer, leading to the synthesis of LacCer by galactosyltransferase-1 (GalT1). Sialyltransferase-1 (SialT1) catalyzes the addition of sialic acid (N-acetylneuraminic acid; NeuAc) on the galactose residue of LacCer to generate the monosialoganglioside (GM3), which is the precursor of complex gangliosides. Further sialylation of GM3 gives rise to the disialoganglioside GD3 and the trisialoganglioside GT3. GM3, GD3, and GT3 represent the entry substrates for the biosynthesis of gangliosides of the a-, b-, and c-series pathways, respectively, while direct conversion of LacCer to GA2 followed by subsequent addition of sugar and sialic acid gives rise to GD1c, the end product of gangliosides of the 0-series (> Figure 10‐3). The synthesis of GD3 and GT3 might be performed by a single enzyme since cDNA alignment indicates that SialT2 and SialT3 have identical nucleotide sequence (Nakayama et al., 1996), and transfection with SialT2 gave rise to GT3 synthesis (Daniotti et al., 2002). In contrast to these observations, lipidomic analysis of the brain of mice lacking b1,4-N-acetylgalactosaminyl-transferase (GalNacT), the enzyme that converts LacCer, GM3, GD3, and GT3 into their respective products, shows no accumulation of GT3 (Takamiya et al., 1996). GT3 accumulation should have occurred, because GD3 accumulation in the brain of that mutant mouse should have been converted to GT3 as the only downstream metabolite from GD3 in that mutant animal. Thus, it has been concluded that a sialyltransferase different from SialT2 is required for GT3 synthesis in vivo (Kolter et al., 2002). The glycosylation of GM3, GD3, and GT3 is performed by a few glycosyltransferases of limited specificity. It is known that these glycosyltransferases physically associate as a multiprotein and that the N-terminal domain of each enzyme participates in this

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interaction (Giraudo and Maccioni, 2003) and, therefore, dictates the Golgi compartmentalization of the multienzyme complex (Uliana et al., 2006a). For example, the complex formed by GalT1/SialT1/SialT2 is located in the Golgi stack (Uliana et al., 2006b; Halter et al., 2007), while the GalNacT and GalT2 complex is mostly located in the trans-Golgi network (TGN) (Giraudo et al., 1999, 2001). The glycosylation of LacCer, GM3, GD3, and GT3 leads to the generation of asialo-GM2, GM2, GD2, and GT2 by the action of GalNacT. Subsequently, GalT2 converts the previous products to GA1 (asialo-GM1a), GM1a, GD1b, and GT1c, which are further sialylated by the consecutive action of SialT4 and 5 to generate the complex final products of the ganglioside synthesis pathway (Kaufman et al., 1968; Sandhoff and van Echten, 1993; Yamashiro et al., 1995; Taylor et al., 2004).

2.2 Cholesterol Biosynthesis In addition to GalCer and sulfatide that together account for 30% of myelin lipid, the other most abundant lipid in myelin is cholesterol. > Figure 10‐4 schematically outlines the cholesterol biosynthesis pathway.

. Figure 10‐4 Cholesterol biosynthesis: The condensation of acetyl-CoA and acetoacetyl-CoA to form HMG-CoA is the biosynthetic pathway of cholesterol in animal cells, but the rate-limiting step is the reaction catalyzed by HMG-CoA reductase. Additional enzymes in the cholesterol biosynthetic pathway are as follows: (2) mevalonate-5-phosphotransferase, (3) phosphomevalonate kinase, (4) pyrophosphomevalonate decarboxylase, (5) prenyl transferase, (6) prenyl transferase, (7) squalene synthase, (8) squalene epoxidase, (9) squalene oxidocyclase

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Briefly, cholesterol synthesis is initiated in the cytosol by the condensation of acetyl-CoA and acetoacetylCoA to yield 3-hydroxy-3-methylglutaryl-CoA (HMG-CoA), a reaction catalyzed by HMG-CoA synthase. HMG-CoA is then reduced to mevalonate by the action of HMG-CoA reductase (HMGR), a tetrameric protein consisting of two dimers that localize to the ER and peroxisomes in the mouse brain stem and cerebellum (Reinhart et al., 1987; Istvan et al., 2000; Kovacs et al., 2001). HMGR is among the most highly regulated enzymes (Goldstein and Brown, 1990). Transcription and translation of HMGR increase when the concentration of products of the mevalonate pathway is low. Conversely, when sterol concentrations are high, the intracellular HMGR concentration decreases rapidly (Nakanishi et al., 1988). A third level of regulation is achieved by phosphorylation of S872 (human enzyme) by AMP-activated protein kinase, which decreases HMGR activity (Omkumar et al., 1994). The synthesized mevalonate is then transformed to cholesterol after subsequent reactions that include pyrophosphorylation, farnesylation, oxidization, and thereafter cyclization (Taylor et al., 2004).

2.3 Phospholipid Biosynthesis 2.3.1 Phosphosphingolipid Biosynthesis Phosphosphingolipids are sphingolipids that contain a phosphate group attached to the primary hydroxyl of ceramide. Among the phosphosphingolipids, sphingomyelin (SM) is a key membrane component of higher eukaryotes. It provides a reservoir of messenger signals that mediate cellular processes such as programmed cell death, cellular stress, mitogenesis, and senescence (Andrieu-Abadie and Levade, 2002; Kolesnick, 2002; Bektas and Spiegel, 2004; Futerman and Hannun, 2004). The synthesis of SM is mediated by a phosphatidylcholine:ceramide cholinephosphotransferase (SM synthase), a membrane-associated enzyme that transfers the phosphocholine moiety from phosphatidylcholine (PdtCho) onto the primary hydroxyl group of ceramide and generates SM and diacylglycerol (DAG: Ullman and Radin, 1974; Voelker and Kennedy, 1982) (> Figures 10-2 and > 5-5). In OLs, most of the phosphocholine used for the biosynthesis of SM is provided by PdtCho (Vos et al., 1997). In addition, SM synthases can act in the reverse pathway, generating PdtCho from DAG and SM (van Helvoort et al., 1994), indicating that SM synthases might regulate the pool of cellular ceramide and DAG, two highly active biomolecules that are implicated in the regulation of membrane trafficking and apoptosis (Scurlock and Dawson, 1999; Brose and Rosenmund, 2002; Lee et al., 2004). Hence, the physiological significance of the expression of SM synthases in eukaryotic cells might be beyond the regulation of ceramide and DAG pool. Recent investigations have demonstrated that human, mouse, pig, and C. elegans genomes contain at least two SM synthase genes: SM synthase-1 and -2 (Huitema et al., 2004; Yamaoka et al., 2004). Whereas SM synthase-1 was localized at the cis/medial Golgi by cell fractionation (Futerman et al., 1990; Jeckel et al., 1990), a tagged SM synthase-1 has now been located in the trans-Golgi by immunoelectron microscopy (Halter et al., 2007). SM synthesis was found to occur in the TGN of neuronal cells (Sadeghlar et al., 2000). SM synthase-2 is mostly detected at the plasma membrane (Futerman et al., 1990; van Helvoort et al., 1994; Huitema et al., 2004). This finding might indicate that SM synthase-2 is transported at the myelin assembly site where it catalyzes the synthesis of SM necessary for myelin biogenesis. Moreover, the most important role of SM is its capacity to participate in the formation of lipid rafts, the sorting platform that is involved in the transport of cell membrane components and signal transduction (Simons and van Meer, 1988; Verkade and Simons, 1997). In this point of view, SM synthase-1 located in the lumen of Golgi in OLs might be responsible for the synthesis of SM necessary for the formation of the lipid raft. Therefore, SM might play a critical role in the sorting and transport of myelin components necessary for the biogenesis of the myelin sheath, since the raft is the sorting platform for apical membrane trafficking (Hoekstra et al., 2003; Fullekrug and Simons, 2004). The other phosphosphingolipid present in myelin is ceramide 1-phosphate (C1P), which is synthesized by the transfer of a phosphate group to the primary hydroxyl group of ceramide. This reaction is catalyzed by ceramide kinase (> Figure 10‐2; CERK). CERK activity is detected in almost all mammalian tissues, but the level of activity differs from tissue to tissue. In mouse, the highest CERK activity was found in testis and brain (van Overloop et al., 2006). Moreover, by separating the subcellular organelles using differential

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. Figure 10‐5 Biosynthetic pathway of phospholipids and ether lipids: phospholipids and ether lipids are derived from the enzymatic transformation of dihydroxyacetone phosphate (DHAP). The following enzymes play a role in the transformation of DHAP: (1) glycerophosphate dehydrogenase, (2) sn-glycerol-3-phosphate acyltransferase, (3) 1-acyl glycerol-3-phosphate acyltransferase, (4) dihydroxyacetone phosphate acyltransferase, (5) acyl/alkyl dihydroxyacetone phosphate reductase, (6) alkyl dihydroxyacetone phosphate synthase, (7) phosphatidate phosphohydrolase, (8) phosphatidate cytidyltransferase, (9) phosphatidylinositol synthase, (10) phosphatidylcholine: ceramide choline phosphotransferase, (11) diacylglycerol cholinephosphotransferase, (12) diacylglycerol ethanolaminephosphotransferase, (13) phosphatidylethanolamine N-methyl transferase and phosphatidyl-N-methylethanolamine N-methyl transferase, (14) phosphatidylethanolamine:serine transferase, (15) phosphatidylserine decarboxylase, (16) phosphatidylinositol 4-kinase, (17) phosphatidylinositol-4-phosphate 5-kinase, (18) phosphatidylinositol-4,5-phosphate 3-kinase, (19) phosphatidylinositol-3-kinase, (20) phosphatidylinositol-3phosphate 4-kinase, (21) phosphatidylinositol-3,4 phosphate 5-kinase, (22) CDP-DAG:glycerol-3-phosphate phosphatidyltransferase, (23) phosphatidylglycerol phosphatase, (24) cardiolipin synthase (Adapted from Farooqui et al. (2000), Cooke (2004), and Taylor et al. (2004))

centrifugation techniques, van Overloop et al. (2006) found that the CERK was concentrated in the microsomal fraction. In addition, immunofluorescence analysis using different cell lines has demonstrated that CERK associates with the Golgi and plasma membrane (Carre et al., 2004; van Overloop et al., 2006). Therefore, C1P might be synthesized both at the plasma membrane and in the Golgi. C1P is a highly bioreactive molecule that regulates many cellular processes including cell survival and proliferation and stimulation of DNA synthesis (Gomez-Munoz et al., 1995, 2004, 2005).

2.3.2 Glycerophospholipid Biosynthesis Glycerophospholipids are compounds similar to triglycerides. However, they have a phosphate group and a simple organic molecule in the place of one of the fatty acids. Brain tissue contains high amounts of

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phospholipids. In the adult brain, glycerophospholipids, glycolipids, and cholesterol account for 50–60% of the total membrane mass with proteins accounting for most of the remainder. Within the brain, myelin contains the highest amount of glycerophospholipid (Farooqui et al., 2000). The most abundant glycerophospholipids of the mammalian tissue are phosphatidylcholine (PtdCho), phosphatidylethanolamine (PtdEtn), phosphatidylserine (PtdSer), and phosphatidylinositol (PtdIns). Besides the above phospholipids, cellular membranes contain plasmalogens, a phospholipid containing a vinyl ether linkage. > Figure 10‐5 outlines the biosynthetic pathway of PtdCho, PtdEtn, and PtdSer that have been described in eukaryotic organisms (van Golde et al., 1974; Vance, 1990; Saito et al., 1996; Stone and Vance, 2000). First, dihydroxyacetone-phosphate (DHAP) is reduced to glycerol-3-phosphate (Gro-3P), which is successively acylated to produce phosphatidic acid (PtdOH). Alternatively, DHAP can be directly acylated followed by alkylation to produce the alkyl-DHAP, which is the precursor of ether lipids. Subsequently, PtdOH can be converted to DAG and CDP-DAG. Once formed, DAG is used for the synthesis of PtdCho and PtdEtn via the CDP-choline and CDP-ethanolamine pathway, also known as the Kennedy pathway (Yavin and Zeigler, 1977; Arthur and Page, 1991; Bakovic et al., 2007). In this pathway, choline or ethanolamine is converted in the cytosol into phosphocholine or phosphoethanolamine by choline kinase (CKI) or ethanolamine kinase (EKI), respectively (Kent, 1995). In the second reaction, the phosphocholine or phosphoethanolamine is transferred to a nucleotide diphosphate by the action of phosphocholine or ethanolamine cytidylyltransferase to form CDP-choline or CDP-ethanolamine, respectively. Finally, the phosphocholine or phosphoethanolamine is transferred to the 1,2-diacylglycerol by the action of CDPcholine or CDP-ethanolamine:1,2-diacylglycerol choline or ethanolamine phosphotransferase (Vermeulen et al., 1997). These enzymes are integral membrane proteins that are predominantly located in the ER (Vance, 1996; Ross et al., 1997). PtdCho is also synthesized by successive methylation of PtdEtn by PtdEtn N-methyltransferases. An alternative route to synthesize PtdEtn is the decarboxylation of PtdSer by PtdSer decarboxylase, an enzyme located on the outer surface of the mitochondrial inner membrane (Percy et al., 1983; Zborowski et al., 1983). In cultured Chinese hamster ovary cells (Miller and Kent, 1986; Nishijima et al., 1986) and baby hamster kidney cells (Voelker, 1985), the decarboxylation of PtdSer produces more than 80% of the PtdEtn, even when the culture medium is supplemented with ethanolamine, an obligatory substrate of the CDP-ethanolamine pathway. This suggests that the decarboxylation of PtdSer is the primary source of PtdEtn biosynthesis. In the CNS, PtdSer is synthesized exclusively by base exchange. In general, the base-exchange reaction is catalyzed by PtdSer synthase I and II, ER enzymes that are activated by Ca2+ (Kuge and Nishijima, 1997). The difference between the two PtdSer synthases is at the level of substrate specificity. While PtdSer synthase I can synthesize PtdSer from PtdCho, the synthase II uses PtdEtn as a substrate (Voelker and Frazier, 1986; Kuge et al., 1997). The structural analysis of PtdSer synthase I has demonstrated that the enzyme lacks the typical N-terminal signal for ER targeting, but contains a C-terminal Lys-Lys motif that was proposed to be an ER-retention sequence (Stone et al., 1998). Biochemical investigation of the subcellular localization of these synthases has demonstrated that the activity of both synthases is associated exclusively with the mitochondrial-associated membranes and the ER membrane (Saito et al., 1996; Stone and Vance, 2000). On the other hand, CDP-DAG is directly converted to PtdIns by PtdIns synthase and phosphatidylglycerol phosphate (PtdGroP) by PtdGroP synthase. PtdGroP is subsequently dephosphorylated to phosphatidylglycerol (PtdGro) by PtdGroP phosphatase. PtdGro is finally converted to cardiolipin by cardiolipin synthase.

3

Sorting and Transport of Lipids During Myelin Assembly

Myelin formation during development and myelin maintenance throughout adult life depends not only on a tight regulation of the expression of genes implicated in the synthesis of myelin components but also on unique membrane trafficking machinery for the proper sorting and targeting of specific components to the myelin sheath. Individual myelin components are synthesized in several compartments, sorted, and transported to the sites of myelin synthesis by different mechanisms (Benjamins and Smith, 1984; Morell et al., 1994; van Meer and Holthuis, 2000; Anitei and Pfeiffer, 2006). The difference in lipid composition

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between cellular organelles and between organelles and plasma membrane can not be explained solely by local metabolism, but can be attributed to the sorting and transport mechanism. How cells decide which lipids need to be moved and in which direction is still a mystery. However, it is known that the selectivity in lipid transport is the main mechanism for lipid sorting (Sprong et al., 2001). In eukaryotic cells, the transport of lipids to the plasma membrane is made possible by monomeric and vesicular transport (van Meer and Holthuis, 2000). A monomeric exchange happens when a lipid desorbs from the membrane into the aqueous phase, diffuses across it, and inserts itself into the opposite membrane (Sprong et al., 2001). Proteins may stimulate lipid transport between membranes by bringing membranes together (Ladinsky et al., 1999). Alternatively, lipid transfer proteins might provide a hydrophobic binding site and act as a carrier. On the other hand, the vesicular transport of lipids toward the plasma membrane happens mostly by lateral segregation of lipids from the Golgi membrane and their exclusion from retrograde transport vesicles (van Meer, 1989). Ceramide is synthesized in the ER and translocates to the Golgi compartment for conversion to more complex sphingolipid species. There are at least two known pathways by which ceramide is transported from the ER to Golgi. The first is mediated by vesicles (Funato and Riezman, 2001). In this system, synthesized ceramide from the ER is packed in a cargo vesicle and delivered by fusing the vesicle membrane to the Golgi membrane. These cargo vesicles preferentially target ceramide molecules to the cis-Golgi (Hanada et al., 2003). The second transport pathway (the ATP- and cytosol-dependent pathway) has been described both in yeast (Funato and Riezman, 2001) and in mammalian cell lines (Hanada et al., 2003, 2007; Kawano et al., 2006) and is known to be mediated by ceramide transfer protein (CERT). CERT is a cytoplasmic protein containing a phosphatidylinositol-4-monophosphate-binding domain and a putative domain for catalyzing lipid transfer (START). This protein specifically extracts ceramide from the phospholipid bilayer of the ER membrane and targets it to the SM synthesis site at the Golgi membrane after diffusion through the cytosol. This targeting event is mediated by the PH domain of CERT (Kumagai et al., 2005, 2007). Alternatively, CERT may induce membrane contacts between ER and trans-Golgi (Munro, 2003). Like ceramide, the glycosphingolipid (GSL) GalCer is synthesized in the lumen of ER, and is a substrate for the synthesis of sulfatide in the lumen of Golgi by CST. This indicates that GalCer must be transported to the Golgi for sulfatide synthesis. It has been reported from in vitro experiments that GalCer translocates from the luminal to the cytosolic face of the ER following its synthesis (Burger et al., 1996). In addition, a GSL transfer protein has been described, which is capable of transferring both GlcCer and GalCer from donor to target membranes, in vitro (Sasaki and Demel, 1985; Sasaki, 1990). This suggests that the transport of GalCer from ER to Golgi might be mediated by those proteins. Except GalCer and ceramide that are synthesized in the ER and transported to the plasma membrane both by vesicular and monomeric transport, SM and the complex GSLs are synthesized in the Golgi lumen and have no access to the monomeric transport (Nilsson and Dallner, 1977; Brown et al., 1993; Burger et al., 1996). In this case, the transport toward the plasma membrane from the Golgi lumen happens by incorporation into anterograde vesicles and exclusion from retrograde vesicles (reviewed by van Meer, 1989; Sprong et al., 2001). This means that in the Golgi lumen, sphingolipids are subjected to lateral segregation from other membrane lipids such as PtdCho. In addition, elaborate studies on lipid transport using Madin-Darby Canine Kidney (MDCK) cells have been performed to elucidate the mechanisms by which lipids are transported in polarized cells such as OL.

3.1 Lipid Transport in Polarized Cells The plasma membrane of polarized cells is divided into two specific compartments: the apical and basolateral membrane compartments. These compartments are distinct from each other by their specific protein and lipid composition. For example, the apical domain displays a twofold higher level of GSLs with a significantly lower level of phospholipids. Such a typical lipid composition is found in OLs (Stoffel and Bosio, 1997). To build up such unique compartments, lipids and proteins have to be sorted and transported to the appropriate plasma membrane compartment. The apical transport of GSLs in polarized cells such as MDCK occurs by direct transport from the TGN. In such a transport mechanism, GSLs are first sorted in

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the TGN by association with protein to form a GSL-enriched domain (Brown and Rose, 1992), or GSL raft (Simons and Ikonen, 1997). The budding of these GSL-enriched domains will give rise to vesicles, which will be transported to the apical compartment (Matlin and Simons, 1984; Misek et al., 1984; Pfeiffer et al., 1985; Simons and Wandinger-Ness, 1990; Zegers and Hoekstra, 1997; Chang et al., 2006). In such a transport pathway, the glycosylphosphatidylinositol (GPI)-anchored proteins associate with sphingolipids and cholesterol in the TGN (Lisanti and Rodriguez-Boulan, 1990; Mun˜iz and Riezman, 2000). Alternatively, the apical delivery of lipids can be made by an indirect pathway in polarized cells. The lipid–protein complex formed in the TGN is first transported to the basolateral domain of the plasma membrane where it is endocytosed and transcytosed to the apical surface (Nyasae et al., 2003; Polishchuk et al., 2004). The similarity between the lipid composition of the myelin sheath and the apical and basolateral membrane domain in polarized cells (Stoffel and Bosio, 1997) suggests that apical and basolateral intracellular transport of lipids may also occur in OL.

4

Role of Lipids in the Regulation of Protein Sorting and Transport

4.1 Fatty Acids and the Regulation of Myelin Protein Sorting and Transport Membrane proteins enter the membrane environment through transportation from their site of synthesis. Following synthesis, proteins are subjected to various posttranslational modifications that include acylation. Cysteinyl-palmitoylation is the major, dynamic posttranslational lipid modification of proteins that appears necessary to direct them to cholesterol/sphingolipid-rich microdomains (rafts) in the plasma membrane (Mumby, 1997; Paterson, 2002; Smotrys and Linder, 2004). In most cases, palmitoylation is the signal for membrane attachment of proteins that have been previously myristoylated at an N-terminal glycine residue or prenylated at the C-terminus (Mumby, 1997; Paterson, 2002; Smotrys and Linder, 2004). In CNS myelin, Src-family tyrosine kinase is palmitoylated in this way, but proteolipid protein is acylated at multiple sites (Bizzozero and Good, 1991). Recently, it has been demonstrated that palmitoylation is the sorting determinant of PLP/DM20 for transport to the myelin membrane and that the N-terminal 13 amino acids, which are palmitoylated at 3 cysteine sites, were sufficient to target PLP/DM20 to the myelinlike membrane in vitro (Schneider et al., 2005). Moreover, the fatty acid chain length of sphingolipids in yeast was found to be crucial for membrane delivery of protein cargo. These findings support the idea that lipids are not playing only a structural role by separating the extracellular from the intracellular compartment, but are more deeply implicated in the regulation of cellular physiology such as sorting and transport.

4.2 Sphingolipids/Cholesterol and the Regulation of Intracellular Transport In membranes, sphingolipids appear organized in clusters or domains called rafts. These domains are formed in the TGN by self-association of newly synthesized sphingolipids/cholesterol and proteins (Simons and Ikonen, 2000). By this association, sphingolipid/cholesterol/proteins are packed into vesicles and delivered to the plasma membrane. The raft formation is the process by which most plasma membraneassociated proteins are sorted and transported. Hence, by recruiting proteins to the raft, sphingolipids regulate the transport of proteins that lack plasma membrane-targeting signals. This hypothesis is confirmed by numerous experiments that used different cell lines. Evidence for the role of sphingolipids and cholesterol in intracellular trafficking has been demonstrated in neuronal cells that also display a polarized trafficking mechanism (Ledesma et al., 1998). Neuronal inhibition of sphingolipid synthesis affects the sorting and transport of Thy-1 protein to the axon, implicating sphingolipids directly to the axonal sorting mechanism. Cholesterol, a component of the raft domain has also been suggested to play a role in the intracellular delivery of proteins. By culturing MDCK cell depleted of LDL, the principal source of cholesterol, the trafficking of gD1-DAF (GPI-anchored protein that preferentially associates to rafts in the TGN) was inhibited (Hannan and Edidin, 1996). In OL, proteolipid protein associates with the CHAPSinsoluble membrane fraction after leaving the ER, but before exiting the Golgi, suggesting that myelin lipids

Functional dynamics of myelin lipids

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and proteins assemble in the Golgi complex before transport to the myelin sheath (Kra¨mer-Albers et al., 2006). Moreover, the binding of PLP/DM20 to cholesterol suggests that cholesterol is required for the sorting and transport of PLP/DM20 to the myelin sheath (Simons et al., 2000). In addition, the myelin and lymphocyte protein (MAL) is a tetraspan raft-associated proteolipid predominantly expressed by OLs and Schwann cells. MAL is synthesized in the ER and transported to the plasma membrane most likely by vesicular delivery (Zacchetti et al., 1995; Kim and Pfeiffer, 1999). To date, it is still unclear how MAL transport to the myelin sheath is regulated. However, it is evident that MAL is redistributed in the endosome–lysosome compartment in sulfatide-storing kidney cells (Saravanan et al., 2004). This finding points in the direction of sulfatide as a regulator of the transport of MAL to the myelin sheath.

5

Role of Lipids in the Biogenesis and Maintenance of Myelin

The biogenesis of myelin by Schwann cells or OLs requires the coordinate synthesis, transport, and integration of large quantities of specific proteins and lipids into the organized multilamellar structure (Morell and Ousley, 1994). The dry mass of myelin is characterized by a high proportion of lipid (70–85%), and consequently, a low proportion of protein (15–30%) (> Table 10‐1; Quarles et al., 2006). In comparison to myelin, most biological membranes display a high protein-to-lipid ratio, with identical lipid species. This suggests that there are no myelin-specific lipids; rather, there are myelin-enriched lipids. Galactolipids fall in this category and constitute 27–30% of the total myelin lipids (Norton and Cammer, 1984). Because of the enrichment of galactolipids (especially GalCer and sulfatide) in OLs and myelin of all mammals, it had been speculated that they would be essential for the formation of the myelin sheath, but in fact, that does not appear to be the case. Specifically, mice lacking CGT that do not synthesize GalCer, sulfatide, monogalactosyldiacylglycerol (MGDG), GM4, and seminolipid (> Table 10‐2; Bosio et al., 1996; Coetzee et al., 1996) are surprisingly able to synthesize the myelin membrane, which exhibits the characteristic ultrastructure of compact myelin, including the major dense line and intraperiod line both in the CNS and PNS (> Figure 10‐1). Similarly, mice lacking CST that do not synthesize sulfatide, seminolipid, LacCer sulfate, and sulfated gangliosides (> Table 10‐2) are also able to synthesize a compact myelin membrane (Honke et al., 2002). Nevertheless, careful analysis of myelin from these mice indicates the presence of substantial alteration of the myelin structure at the paranodal junction observed in the CNS (Dupree et al., 1999; Marcus et al., 2000; Ishibashi et al., 2002; Marcus and Popko, 2002; Marcus et al., 2002; Rasband et al.,

. Table 10‐2 Lipids based comparison between CGT- and CST-null mutant mice Galactolipids GalCer Sulfatide (SM4) MDGD GM4 OH-GlcCer GalEAGa Seminolipid LacCer-sulfate (SM3) SM2a SM1a SB1a

Synthesis compartment ER Golgi ER Golgi Golgi ER Golgi Golgi Golgi Golgi Golgi

() means that the lipid is lost (+) means that the lipid is present (0) means that the lipid does not normally exist in myelin a Galactosylalkylacylglycerol

CGT-null     +    + + +

CST-null +  + + 0 +     

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2003) and PNS (Hoshi et al., 2007). The disruption of the paranodes in the CNS of galactolipid mutant mice is probably due to defects in the clustering of nodal, paranodal, and juxtaparanodal proteins such as sodium and potassium channels (Dupree et al., 1999; Ishibashi et al., 2002), neurofascin-155 (Dupree et al., 1999; Schaffer et al., 2004), caspr, and paranodin (Dupree et al., 1999). Further, CGT- or CST-null mice display significant alterations of the myelin sheath and axon in adulthood (Stoffel and Bosio, 1997; Coetzee et al., 1998; Dupree et al., 1998, 2005; Marcus et al., 2006). These findings suggest that GalCer and sulfatide are required for the stabilization and maintenance of myelin. However, to draw such a conclusion it will be important to segregate the roles played by sulfatide and GalCer and the other glycolipids also missing in the mutants. Beside galactolipids, the other abundant lipids of the myelin membrane are cholesterol and phospholipids. Based on weight, the level of cholesterol in myelin is comparable to that of galactolipids, and the total phospholipids are the most abundant (> Table 10‐1). On a molar basis, in contrast, CNS myelin preparations contain more cholesterol than any other lipid classes (Morell, 1984; Morell and Jurevics, 1996). In addition to being the most abundant myelin lipid species, cholesterol is known to be implicated in the regulation of some cellular physiology such as membrane structure, thickness, fluidity (Ohvo-Rekila et al., 2002), and to limit ion leakage through the membrane (Haines, 2001). Together with other lipids, cholesterol participates in the formation and stabilization of lipid microdomains that serve as platforms for protein sorting and signal transduction (Simons and Toomre, 2000; van Meer and Lisman, 2002; van Meer and Vaz, 2005). Moreover, cholesterol defines the biophysical properties of all cell membranes. By compacting phospholipids, it may reduce membrane fluidity, and defines the functional properties of membrane-resident proteins, such as ion channels and transmitter receptors (Burger et al., 2000). Most brain cholesterol is unesterified and primarily localized within the myelin sheath. However, cholesterol seems to be required for myelination. This idea is supported by the fact that the CNS white matter of mice with OL-specific disruption of cholesterol biosynthesis is severely hypomyelinated and the mice develop ataxia and tremor. Examination of OLs by TUNEL experiment shows no signs of cell death, indicating that OL apoptosis was not the reason for hypomyelination (Saher et al., 2005). Moreover, numerous studies using murine (Quan et al., 2003) and human (Thelen et al., 2006) brains have demonstrated that the level of cholesterol rises during development and declines with aging. The increase of cholesterol during development was significantly higher in the brain stem and spinal cord, two regions of the CNS known to contain high amounts of myelin. These data strongly support the hypothesis that cholesterol synthesis is critical for myelin biogenesis. In addition to major lipid component, the myelin membrane also contains minor components such as gangliosides, which represent 0.1–0.7% of total myelin lipid (Suzuki et al., 1967; Ledeen et al., 1980; Ccochran et al., 1982; Quarles et al., 2006). Besides GM4, the monosialoganglioside GM1 represents the most abundant ganglioside species in the myelin membrane. Numerous studies have demonstrated that myelin basic protein could interact directly with gangliosides such as GM1, GM4, and GD1b, at least in vitro (Yohe et al., 1983; Ong and Yu, 1984). In addition, GM1, GT1b, GD1a, and GD1b modulate protein phosphorylation in myelin, and in contrast, completely inhibit the phosphorylation of the 18.5-kDa MBP isoform (Chan, 1987). More importantly, the ganglioside content of myelin increases during the maturation of the myelin sheath and may reflect myelination. To determine if ganglioside accumulation is a crucial factor in myelinogenesis, genetically engineered mice with defects in enzymes catalyzing specific biosynthetic pathways have been generated (> Figure 10‐3). Analysis of these mutant animals has provided strong insights for the role of gangliosides in myelinogenesis. Mice lacking specific ganglioside series develop normal myelin (Horinouchi et al., 1995; Furukawa et al., 2001; Kolter et al., 2002). In contrast, gangliosides are required for growth and myelin maintenance. This affirmation is strongly supported by the results from analysis of mice lacking all or a number of ganglioside series. Mice deficient in ceramide glucosyltransferase (> Figure 10‐3a) begin to die as early as embryonic day 7.5, indicating that GlcCer and higher-order glycolipids are important for development, but it is important to know that accumulation of ceramide in such an animal could also lead to deleterious effects. As shown in > Figure 10‐3, GalNAcT-null (> Figure 10‐3d) animals lacking the major gangliosides GM2, GD2, GM1a, GD1b, GD1a, GT1a, GT1b, and GQ1b, and SialT2-null (> Figure 10‐3c) animals lacking GD3, GD2, GD1b, GT1b, and GQ1b, displayed only a subtle impairment of brain function that included demyelination (Sheikh et al., 1999; Chiavegatto et al., 2000). Double mutant mouse deficient in both

Functional dynamics of myelin lipids

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GalNacT and SialT1 synthesize only LacCer and SM3 as major brain gangliosides. These mice display a striking vacuolar pathology in the white matter of the CNS with axonal degeneration and perturbed axo– glial interaction (Yamashita et al., 2005). Moreover, double mutant mice that lack both GalNacT and SialT2 and express only GM3 as major brain gangliosides display a disrupted paranodal axo–glial junction (Susuki et al, 2007a). Similar paranodal abnormalities were observed in the PNS of SialT2 mutant mice lacking the b-series gangliosides, but with increased levels of GM1 and GD1a (Okada et al., 2002). These findings suggest a role for gangliosides in the maintenance of the CNS myelin.

6

Role of Lipids in the Regulation of Oligodendrocyte Physiology

6.1 Fatty Acids As one of the fattiest tissues in the body, the brain needs fats (together with glucose) for energy, structure, and maintenance of its normal function. In the CNS and PNS cells, as well as in the other cells of living organisms, fatty acids (FAs) are key components of phospholipids and sphingolipids, which are the most abundant components of the cellular membranes and myelin. They are commonly classified as saturated, monounsaturated (MUFA), and polyunsaturated fatty acids (PUFA) (Agostoni and Bruzzese, 1992; Millar and Kunst, 1997). According to the hydrocarbon chain length, FAs are also classified as short-chain, longchain fatty acids (LFAs), and very long chain FAs (VLCFAs). The richest source of saturated and monounsaturated FAs in the brains of most animals is myelin (Bourre and Baumann, 1980; for review, see Poulos, 1995). FAs are known to play a major role in the regulation of myelin thickness, myelin structure, and compaction (Bourre et al., 1978a, b). In addition, the observation of deficiencies in PUFAs in MS patients has led to attempts to influence the disease course by dietary uptake, particularly by increased intake of specific PUFAs of n-3 and n-6 series (Borlak and Welch, 1994; Mayer, 1999). Moreover, dietary supplementation of gamma-linolenic acid (18:3n-6) ameliorates the course of both acute and chronic experimental autoimmune encephalomyelitis (EAE: Harbige et al., 2000), and FAs from n-6 series improve biochemical parameters and cognitive functions in rats with EAE (Yehuda et al., 1997). PUFAs in these cases might influence the disease course by repairing the myelin sheath or stimulating OL progenitor differentiation. This idea is supported by findings that MBP and PLP mRNA levels were reduced in pups nursed by mothers that were fed a fat-free diet, and this effect was reversed by feeding the mother with a corn-oil-based diet rich in PUFAs (DeWille and Farmer, 1992). In addition, supplementing primary cultured OLs with PUFAs results in an increase of CNP, MBP, and PLP expression (van Meeteren et al., 2006). The stimulating effect of PUFAs on the differentiation of OL progenitors might be mediated through the thromboxane receptor, since it has been recently demonstrated that the metabolites of arachidonic acid, eicosanoids (e.g., thromboxane A2 and prostaglandins) are produced in cells following the action of cellular stimuli that activate phospholipases A2 and C, leading to the liberation of membrane-esterified arachidonic acid. Free arachidonic acid is first metabolized by the cyclooxygenases (COX-1 or COX-2) and then by terminal prostaglandin synthases to produce the prostaglandins (PGD2, PGE2, PGF2a, PGI2) and thromboxane A2 (TXA2). Arachidonic acid metabolites have been implicated in apoptosis and neurodegeneration (Brault et al., 2004; Farooqui et al., 2004). On the other hand, one of these eicosanoids, TXA2, has been implicated in the proliferation of OL progenitors and survival of mature OLs through its interaction with TXA2 receptors (Lin et al., 2005; Ramamurthy et al., 2006) present in myelinated fibers of the optic nerve and striatum (Borg et al., 1994). In addition, it is also likely that PUFAs might modulate the expression of OLspecific genes through syntaxin-3, since it is evidenced that PUFAs of n-3 and n-6 series promote membrane growth in PC12 cells in vitro by acting on syntaxin-3 (Darios and Davletov, 2006).

6.2 Sphingolipids and the Regulation of Oligodendrocyte Lineage Progression The normal timing of OL differentiation can be reconstituted in cultures of postnatal rat brain. This requires that OL progenitor cells (OPCs) are stimulated to proliferate by platelet-derived growth factor

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(PDGF: Raff et al., 1988). It also requires the presence of hydrophobic signals such as thyroid hormone (TH) or retinoic acid (RA) (Barres et al., 1994). Clonal analyses in such cultures show that the progeny of an individual OPC stops dividing and differentiates at about the same time, even if separated and cultured in different microwells, indicating that an intrinsic timing mechanism operates in OPCs to limit their proliferation and initiate differentiation after a certain period of time or number of cell divisions (Temple and Raff, 1986; Barres et al., 1994). Among the molecules that can modulate the differentiation of OPCs in vitro and in vivo are lipids, particularly sphingolipids. The development of antibodies against GSLs has been a critical tool to identify specific stages of OL lineage progression both in vitro and in vivo. The early progenitor stage is characterized by the expression of specific antigens such as PDGF alpha-receptor, A2B5 and GD3 ganglioside, and by uni- or bipolar morphology. The next stage is the late progenitor or prooligodendroblast (Pro-OL) stage, which is identified by the expression of an unidentified sulfated antigen called Pro-OL antigen (POA: Gard and Pfeiffer, 1990; Knapp, 1991; Bansal et al., 1992), which reacts with the monoclonal antibodies O4 and A007 (these antibodies also recognizes sulfatide and seminolipids on differentiated OL). At the Pro-OL stage of lineage progression, sulfatide is not synthesized (Bansal and Pfeiffer, 1994a; Bansal et al., 1992). Further, inhibition of sulfation both in vivo (Bansal et al., 1999; Hirahara et al., 2004) and in vitro (Bansal and Pfeiffer, 1994a) completely eliminates immunoreactivity of Pro-OLs with O4/A007, confirming that POA is a sulfated antigen. Cells at this stage are morphologically characterized by additional primary processes and are still proliferative. The decision to stop proliferating and start differentiating is made at the pre-GalCer stage, characterized by immunoreactivity with the monoclonal antibody R-mAb that recognizes both sulfatide and GalCer (Ranscht et al., 1982; Bansal et al., 1989). Although cells at this stage are R-mAb-positive, the immunoreactivity with O1 (a monoclonal antibody that reacts with GalCer but not sulfatide) is still negative. This is a highly transient stage and it is possible that R-mAb could be recognizing an antigen other than sulfatide at this stage. Terminal differentiation of OL starts as cells exit the pre-GalCer stage and is characterized by a change in the morphology and a dramatic increase in secondary processes. Biochemically, this stage of OL lineage progression is characterized by the appearance of GalCer, sulfatide, CNP, and MAG on the OL membrane (Pfeiffer et al., 1993). This stage can be identified by immunostaining with O1, O4, and R-mAb, which specifically binds to the major GSLs, GalCer, and sulfatide (Bansal et al., 1989; Bansal and Pfeiffer, 1992) (> Figure 10‐6). As cells exit from the immature OL stage of the lineage progression, they start to synthesize markers for mature OL such as MBP, PLP, and MOG. Among these proteins, MBP is the only protein that is required for synthesis of the myelin sheath (Peterson and Bray, 1984).

. Figure 10‐6 Schematic representation of the oligodendrocyte developmental pathway. Each stage of the lineage is characterized by a change in morphology, migratory and proliferative capacity, and the expression of specific protein and lipid markers (Adapted from Pfeiffer et al., 1993; Taylor et al., 2004)

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More than a decade ago, it was proposed that GalCer and sulfatide regulate OL lineage progression. The implication of GalCer and sulfatide in the regulation of OL physiology was observed in vitro. When late progenitors were exposed to anti-GalCer/sulfatide (R-mAb) or anti-sulfatide (O4), their terminal differentiation was reversibly blocked (Bansal and Pfeiffer, 1989; Bansal et al., 1999). Since the inhibition of OL lineage progression at the Pro-OL stage was not observed with anti-GalCer (O1) or other anti-lipids antibodies such as anti-human natural killer-1 (HNK-1), or anti-cholesterol, it was concluded that sulfatide is the glycosphingolipid that regulated the terminal differentiation during OL lineage progression (Bansal and Pfeiffer, 1989; Bansal et al., 1999). The generation of mice lacking sulfatide and GalCer together (Bosio et al., 1996; Coetzee et al., 1996) or sulfatide alone (Honke et al., 2002) has been an important contribution in understanding the effect of GalCer and sulfatide in the development of OL in vivo. In the absence of GalCer and sulfatide together (Bansal et al., 1999) or sulfatide alone (Hirahara et al., 2004), the terminal differentiation of OL is enhanced, indicating that sulfatide, but not GalCer, plays a key role in the entry of OL progenitors into terminal differentiation. Unfortunately, the mechanism by which GSLs mediate the regulation of OL development has not yet been definitively identified. However, pioneering studies indicate that exposure of OL cultures to antiGalCer followed by crosslinking of the complex GalCer/anti-GalCer induces the translocation of membrane surface GalCer to the internal MBP domain, disruption of microtubules and microfilaments within the myelin sheath, and influx of extracellular calcium (Dyer and Benjamins, 1988). Further, treatment of mature OLs with O4 or RmAb leads to process retraction and upon crosslinking of the complex O4/ sulfatide or RmAb/GalCer/sulfatide with secondary antibodies results in a dramatic hyperphosphorylation of MAPK; in contrast, crosslinking of the complex O1/GalCer had no effect on the phosphorylation state of MAPK (Bansal and Pfeiffer, 1994b; Stockdale Ngamli-Fewon, and Pfeiffer, unpublished observation). Taken together, these findings indicate that GSLs can act as receptors that mediate signal transduction. Moreover, anti-GalCer and antisulfatide together induce dysmyelination in vitro (Rosenbluth and Moon, 2003) while antisulfatide alone induces demyelination in vivo (Rosenbluth et al., 2003).

6.3 Gangliosides and Oligodendrocyte Physiology Gangliosides are sialic acid-containing GSLs that are known to modulate the activity of a number of receptor tyrosine kinases, including the insulin receptor (Allende and Proia, 2002). For example, the tyrosine kinase activity of the epidermal growth factor receptor can be enhanced or repressed by gangliosides GD1a or GM3, respectively (Bremer et al., 1986; Liu et al., 2004). In addition, the activities of the platelet-derived growth factor receptor and the nerve growth factor receptor TrkA are negatively regulated by overexpression of GM1 (Mitsuda et al., 2002; Nishio et al., 2005). In both cases GM1 appears to act by displacing the PDGF receptor or TrkA from lipid rafts to the nonraft compartment (Allende and Proia, 2002; Pike, 2003; Ikonen and Vainio, 2005). GM1, GD1a, GD1b, and GT1b are the most abundant gangliosides in the adult mammalian nervous system (Yu et al., 1989). The neurobiological roles of the major nervous system gangliosides are not completely defined. However, the gangliosides GT3 and O-acetyl GT3 are surface antigens that are expressed at the early stage of OL lineage progression and immunoreact with the monoclonal antibody A2B5 (> Figure 10‐6; Farrer and Quarles, 1999). Together with GD3 and GD1a, A2B5-labeled gangliosides are the most abundant gangliosides expressed in early progenitor cells as assessed by double immunostaining with NG2 in the human brain (Marconi et al., 2005). As OLs differentiate, these ganglioside epitopes disappear from the membrane surface and the immature stage shows no immunoreactivity. Marconi et al. (2005) also showed that GD2 is preferentially expressed by mature OLs in the human adult brain. In the peripheral nervous system (PNS), the most abundant gangliosides are GM3, GD3, and sialosylneolactotetraosylceramide [NeuAc(a2-3 or a2-6)Gal(b1-4)GlcNAc(b1-3)Gal(b1-4)Glc(b1-1)Cer] also known as sialosylparagloboside. At the early stage of Schwann cell development GM3 and GD3 with 50 and 18 mol%, respectively, are the most abundant gangliosides of the PNS, but the amount of these lipids decreases as Schwann cells mature and myelinate (Chou et al., 1982). In contrast, the amount of sialosylparagloboside did not change with PNS development. The particularity of the PNS gangliosides is the presence of VLCFAs in the sialosylparaglobosides

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compared with the CNS gangliosides, which did not contain VLCFAs. Another important difference between PNS and CNS gangliosides is the presence of GlcNAc in the neolactosyl series in the PNS instead of GalNAc in the gangliosyl series of the CNS (Chou et al., 1982; Ogawa-Goto and Abe, 1998). The implication of gangliosides in the regulation of cellular physiology has been studied mostly in nonglial cells. However, it has been shown that exogenous GM3 enhances differentiation of OLs (Yim et al., 1994), indicative of its role in OL differentiation. Numerous studies have reported the involvement of gangliosides in the long-term stabilization of axon and myelin contacts through the interaction of GD1a/ GT1b with MAG both in the CNS and PNS (Sheikh et al., 1999; Pan et al., 2005). In vitro studies have demonstrated that gangliosides are functional nerve cell ligands for MAG (Vyas et al., 2002), and that binding of MAG to gangliosides leads to signal transduction by inducing the translocation of p75NTR to lipid rafts (Fujitani et al., 2005). Moreover, the ganglioside GM1, which is not the binding partner of the myelin MAG protein, is implicated in the stabilization of the paranodal axo–glial junctions and ion channel clusters in myelinated nerve fibers both in the CNS and PNS (Susuki et al., 2007a, b). These findings suggest a functional role for gangliosides in the development and maintenance of the CNS.

7

Conclusion

During the last two decades, lipids have attracted widespread attention due to the appreciation that this class of molecules has a major impact on various biological processes. Lipids are the major components in the cell membrane of all organisms and are synthesized at different membrane compartments and then transported to the plasma membrane. Together with membrane-associated proteins, lipids build up the plasma membrane and act as barriers that separate the extracellular from the intracellular compartment. In addition to their structural role at the plasma membrane, lipids are being assigned a broad range of new functions, including regulation of cell growth and differentiation, signal transduction, regulation of intracellular trafficking, and apoptosis. Future avenues of research are likely to be directed toward a better understanding of these new functions of lipids. Lipids have been originally described as cornerstones in the field of neurochemistry and myelin biology. Gene-targeting studies have shown that most of the lipids present in the myelin sheath are needed for myelin stabilization and maintenance, although these are not critical for initial myelin formation, with the exception of cholesterol. In this chapter, we emphasized areas of particular promise in myelin lipidomics that include analysis of mechanisms by which lipids regulate myelin biogenesis, protein sorting, transport, and OL physiology. For further background, we refer the reader to our recent comprehensive chapter on myelin lipids (Taylor et al., 2004).

Acknowledgments We would like to thank Dr. Martin R. Schiller (UConn Health Center) for useful suggestions during the writing and correction of this chapter. We also acknowledge the support of the National Institutes of Health through the grant NS10861.

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Section 3

Function of Neural Lipids

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The Phosphoinositides

G. D’Angelo . M. Vicinanza . A. Di Campli . M. A. De Matteis

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270

2 2.1 2.2 2.3

The The The The

Phoshoinositide Kinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 273 Phoshoinositide 3-Kinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 273 Phosphoinositide 4-Kinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 274 Phosphatidylinositol Monophosphate Kinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 276

3 3.1 3.2 3.3

The The The The

Phosphoinositide Phosphatases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 278 Phosphoinositide 3-Phosphatases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 278 Phosphoinositide 4-Phosphatases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279 Inositol 5-Phosphatases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 280

4

Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 282

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_11, # Springer ScienceþBusiness Media, LLC 2009

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Abstract: Phosphatidylinositol (PtdIns) is a membrane phospholipid that comprises the polar myoinositol hexahydroxycyclohexane headgroup attached via a phosphoester bond to sn-1,2-diacylgycerol 3-phosphate. The phosphoinositides are derivatives of PtdIns in which one or more of the OH groups on the inositol ring have undergone esterification with a phosphate group. In many cell lines and tissues, the phosphoinositides represent up to 15% of the total cellular phospholipids, and they show remarkable differences in concentrations among their diverse species (ranging from around 10% of total phospholipids for PtdIns, to trace amounts of PtdIns(3,4,5)P3). In the central nervous system, the phosphoinositides account for less than 4% of the total phospholipids (less than 1% dry weight in gray matter). Nevertheless, the phosphoinositides have emerged as key regulators of a plethora of biological functions, including synaptic transmission. The importance of this class of lipids is underlined by the finding that genetic impairments in phosphoinositide metabolism produce serious health disorders that often involve the nervous system.

1

Introduction

Phosphatidylinositol (PtdIns) is the precursor for all of the phosphoinositides, and it comprises a D-myo-inositol 1-phosphate molecule that in mammals is generally linked to 1-stearoyl, 2-arachidonoyl diacylglycerol, through the phosphate group. This particular fatty-acid composition of PtdIns is probably related to the correct insertion of the phosphoinositides into the lipid bilayer, to allow sufficient exposure of the polar inositol headgroup for its interactions with cytosolic proteins. Reversible phosphorylation of the inositol ring in positions D-3, D-4, and D-5 leads to the production of seven different phosphoinositide species (see > Figure 11-1). PtdIns itself is however the most abundant of these inositol lipids in mammalian cells under basal conditions, and it usually constitutes 10% of the total membrane phospholipids. Of the monophosphorylated phosphoinositides in cells, >90% is found as PtdIns(4)P, while PtdIns(3)P and PtdIns(5)P each make up from 2 to 5%. Similarly, PtdIns(4,5)P2 is the most abundant of the bis-phosphorylated phosphoinositides (99%). The actual cellular levels of the single tris-phosphorylated phosphoinositide, PtdIns(3,4,5)P3, can vary remarkably in response to various stimuli, although its maximal levels of upregulation remain 500-fold lower than those of PtdIns(4,5)P2. When expressed as the time spent by a molecule of myo-inositol as part of the phosphoinositides pool, the half-life has been estimated as about 7 h (Chikhale et al., 2001); however, the interconversions among the different phosphorylated species of the phosphoinositides occur with much faster kinetics. This last feature renders the phosphoinositdes susceptible to rapid changes in their relative concentrations upon stimulation or inhibition of their metabolizing enzymes. The phosphoinositides are concentrated in the cytoplasmic leaflet of cellular membranes, where they are substrates for different enzymes, including the phosphoinositide kinases (PIKs) and phosphatases (> Figure 11-1), and several phospholipases. An interesting new concept has arisen from evidence that has indicated that each of the seven phosphoinositide species has a particular subcellular distribution, with a predominant localization to a specific subset of membranes (De Matteis and Godi, 2004) (> Figure 11-1 and > 11-2). The common phosphoinositide precursor, PtdIns, is synthesized in the reaction: CDP

diacylglycerol þ myo

inositol CMP þ phosphatidyl

1D

myo

inositol

This reaction is catalyzed by the enzyme CDP-diacylglycerol–inositol 3-phosphatidyltransferase (EC 2.7.8.11), which is otherwise known as PtdIns synthase (PIS). PIS is mainly localized in the endoplasmic reticulum (ER) in both mammalian (Antonsson, 1997) and yeast (Gardocki et al., 2005) cells. Once produced, PtdIns can enter the secretory pathway as a membrane constituent or it can be mobilized via specific lipid-transfer proteins that are collectively known as the phosphoinositide-transfer proteins (PITPs) (Cockcroft, 2007). Indeed, the phosphorylation of PtdIns has been reported to occur mostly in compartments different from the ER (Loijens et al., 1996; Gehrmann and Heilmeyer, 1998; Cockcroft and De Matteis, 2001).

The phosphoinositides

11

. Figure 11-1 The phosphorylation/dephosphorylation cycles of phosphoinositides. The table lists the steps catalyzed by phosphoinositide kinases (dark grey arrows) and phosphoinositide phosphatases (light grey arrows) from yeast and higher eukaryotes, along with their intracellular locations. Abbreviations: GC,Golgi complex; PM, plasma membrane; ER, endoplasmic reticulum; N, nucleus; E, endosomes; LE, late endosomes; Ly, lysosomes; SV, synaptic vesicles; CCV, clathrin-coated vesicles; Mi, mitochondria; ND, not determined

The level of any given phosphoinositide in a cellular subcompartment is mainly the result of the combined actions of the specific kinases and phosphatases. Indeed, the PIKs and the phosphoinositide phosphatases have been localized to almost all intracellular membrane compartments, including the plasma membrane, the nucleus, secretory granules, endosomes, the ER, and the Golgi complex (> Figure 11-1). Relatively high local concentrations of different phosphoinositides can produce specialized membrane– cytosol interfaces, to which proteins with specific affinities for specific phosphinositides can be recruited. In this framework, the discovery of a growing number of protein modules with relatively high affinities for different specific phosphinositide species has provided clues that have allowed us to at least begin to

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. Figure 11-2 ‘‘‘‘P’’hosphoinositide-localization-map’’ on the endocytic and exocytic pathways. During endocytosis, PtdIns (4,5)P2 is required for invagination of coated pits (CP) at the plasma membrane, after which the local level of PtdIns(4,5)P2 decreases because of the activity of a 5-phosphatase. At steady-state, PtdIns(3)P is present almost exclusively in early endosomes (EE) and internal vesicles of multivesicular bodies (MVB), and it is required in Golgi-to-vacuole transport in yeast and for multiple endocytic steps, including docking, fusion, and motility of the EE in mammals. PtdIns(3)P functions in concert with Rab5 in recruitment of FYVE-domain-containing proteins. The conversion of PtdIns(3)P into PtdIns(3,5)P2 occurs at the MVBs/late endosome (MVB/LE) owing to PIK-FYVE (Fab1p in yeast), and is required for protein sorting at the MVBs and for controlling the size of the vacuole/lysosome (Ly). The initial localized production of PtdIns(4,5)P2 at the site of phagocytosis, is followed by a decrease through the action of PLC and type I PI(3)K, which converts PtdIns(4,5)P2 into PtdIns(3,4,5)P3. Later, PtdIns(3)P, produced by type III PI(3)K (Vps34), is required for phagosomal maturation. Ph, phagosome; Ph-ly, phagolysosome. For regulated exocytosis, PtdIns(4)P is generated in the secretory granules (SG), whereas PtdIns(4,5)P2 is generated at the plasma membrane through the Arf6- and calcium-dependent activities of PIP (5)K, which is required for the docking and fusion steps. The phosphoinositide/phosphatidylcholine balance maintained by the PITP Sec14p is necessary for the structure and function of the Golgi complex and considerable evidence indicates that in yeast and mammals the relevant phosphoiositide in the Golgi complex is PtdIns (4)P. Defects in yeast and mammalian Golgi-localized PI(4)Ks impair constitutive transport from the Golgi complex to late endosomes and to the plasma membrane, and perturb Golgi architecture

The phosphoinositides

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decipher the biological functions of the phosphinositides themselves. In this chapter, we will focus on the description of the enzymes that are involved in phosphoinositide metabolism, with particular regard to their intracellular distributions.

2

The Phoshoinositide Kinases

Many of the PIKs have been conserved during evolution from unicellular yeast to mammals. Historically, the PIKs were initially described as enzymatic activities that transfer a phosphate group to a precise position on the inositol ring of PtdIns, or of its phosphorylated derivatives. Studies on the purified enzymes then led to their classification into three main families: the PtdIns 3-kinases (PI3Ks), PtdIns 4-kinases (PI4Ks), and PtdInsP 5-kinases (PIP5Ks). More recently, the genes encoding several of these enzymes have been identified, and the specific features of each one have been investigated or are under active investigation.

2.1 The Phoshoinositide 3-Kinases In the mid 1980s, a PIK activity was found to be associated with polioma middle T antigen (Macara et al., 1984; Sugimoto et al., 1984; Whitman et al., 1985) and with phosphorylated proteins from PDGFstimulated fibroblasts (Kaplan et al., 1987). This activity could phosphorylate inositol phospholipids at position D-3 of their inositol head group. Over the last two decades, these PI3Ks have emerged as a complex family of enzymes that can catalyze the phosphorylation of the different inositol lipids on position D-3. On the basis of their substrate specificities, and structural and functional homologies, the PI3Ks have been subdivided into three main classes: I, II, and III (Fruman et al., 1998). The class I PI3Ks are further subdivided into subclasses IA and IB: class IA PI3Ks are heterodimeric enzymes consisting of a catalytic subunit (p110a, p110b, and p110g) and a regulatory subunit (p85a, p85b, and p55g), while class 1B PI3K comprises a p110 catalytic subunit and a p101 regulatory subunit. Recently, A homologue of the p101 regulatory subunit has also been described, and is known as p84 or p84PIKAP (Suire et al., 2005; Voigt et al., 2006). At present, there is no evidence that within the IA and IB PI3K subclasses there are any significant preferences for combinations of specific catalytic/ regulatory subunit pairs, thus increasing the complexity of this family of kinases. In vivo, PtdIns(4,5)P2 is the preferred substrate of class I PI3Ks, and hence the primary product here is PtdIns(3,4,5)P3. However, these PI3Ks can also phosphorylate PtdIns and PtdIns (4)P in vivo. The class I PI3Ks have been a major focus of attention since they are generally coupled to receptor activation at the plasma membrane by extracellular stimuli. Thus they have been implicated in a wide range of cellular processes, including cell-cycle progression, and cell growth, motility, adhesion, and survival. As indicated above, upon activation, type I PI3Ks produce PtdIns(3,4,5)P3, and after some delay, PtdIns(3,4)P2. Evidence from many laboratories have arrived at the concept that a subset of pleckstrinhomology (PH) domains selectively bind PtdIns(3,4,5)P3 and PtdIns(3,4)P2 with relatively high affinities, and several proteins that contain these types of PH domain have been demonstrated to be involved in class I PI3K-dependent cellular function. The resulting model thus ascribes specific roles in the deciphering of the changes in the concentrations of PtdIns(3,4,5)P3 and PtdIns(3,4)P2 during cellular events to their various PH-domain-containing effectors. However, in many cases, how PtdIns(3,4,5)P3/ PtdIns(3,4)P2 binding to their effector proteins modulates their downstream activities is not clear. Nevertheless, a common theme among the PtdIns(3,4,5)P3/ PtdIns(3,4)P2 effectors is that they have sufficient affinity for these phosphinositides to be relocated to the plasma membrane from the cytosol after the activation of the class I PI3Ks. This recruitment brings the PtdIns(3,4,5)P3/ PtdIns(3,4)P2 effectors in close proximity to their own substrates and other binding partners, thus leading to topological activation. At the same time, it is becoming obvious that in some cases the binding of PH domains to PtdIns(3,4,5)P3/ PtdIns(3,4)P2 can instead remove intramolecular inhibition that is mediated by the PH domains, and as a consequence, activate an effector (Hawkins et al., 2006). It is, indeed, possible that both of these mechanisms operate concomitantly in most cases.

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The class II PI3Ks are larger proteins that have catalytic domains that are 45–50% identical to those of the class I PI3Ks. The class II PI3Ks have a C-terminal region that has homology with the C2 domain of the classical protein kinase C isoforms that mediate Ca2+/ lipid binding. Although they preferentially phosphorylate PtdIns and PtdIns(4)P in vitro, the in vivo substrates and functions of the class II PI3Ks are less clear. There are three members of these class II PI3Ks (PI3K-C2a, PI3K-C2b, and PI3K-C2g), and they can be activated by several stimuli, although, again, the physiological consequences of their activation remain to be determined in many cases. Nevertheless, they have been seen to have roles in migration of cancer cells (Maffucci et al., 2005), cytoskeleton organization (Katso et al., 2006), neurosecretory granule exocytosis (Meunier et al., 2005), smooth-muscle-cell contraction (Wang et al., 2006), and insulin signaling (Falasca et al., 2007). Moreover, PI3K-C2a can bind clathrin, and it is stimulated by this interaction, with its overexpression inducing the redistribution of the mannose 6-phosphate receptor from the trans-Golgi network (TGN) to the cell periphery and inhibiting endocytosis (Gaidarov et al., 2001; Gaidarov et al., 2005). Thus, PI3K-C2a might control clathrin-dependent sorting events at the TGN through a localized generation of PtdIns(3)P at sites of clathrin-coated bud formation. Accordingly, PI3K-C2a has been seen to localize in spots in the perinuclear area (Gaidarov et al., 2001), and also in the nucleus in resting cells (Falasca et al., 2007), while it translocates to the plasma membrane upon cell stimulation (Falasca et al., 2007). Similarly, PI3K-C2b has been seen in the nucleus, and it translocates to the plasma membrane upon cell stimulation (Crljen et al., 2002). The class III PI3Ks are highly related to the yeast Vps34 gene product, and as with this yeast enzyme, they use PtdIns as the specific substrate for the production of PtdIns(3)P. Different trafficking processes around transport to the yeast vacuole depend on this Vps34-mediated production of PtdIns(3)P. Indeed, mutations in the Vps34 protein result in the missorting of vacuolar proteins, changes in vacuole morphology, and defects in the endocytic pathway (Stack et al., 1995; Takegawa et al., 1995; Kihara et al., 2001). In mammalian cells, inhibition of the Vps34 orthologe by microinjection of specific antibodies or by gene silencing prevents the formation of internal vesicles in endosomes, and thus of multivesicular bodies (MVBs), without impinging on vesicle fusion to lysosomes (Futter et al., 2001; Johnson et al., 2006). Knockdown of hVps34 reduces the amount of PtdIns(3)P associated with late endosomes, with the PtdIns(3)P indeed enriched in the endocytic/ lysosomal compartment (Gillooly et al., 2000), where it is involved in the recruitment of different proteins that have domains that are known to bind to PtdIns(3)P with high affinities and specificities: the FYVE and PX domains. Among these proteins, the best characterized are the early endosomal GTPase Rab5 effectors, which regulates endocytic membrane fusion (Simonsen et al., 1998; Zerial and McBride, 2001), EEA1 and rabenosyn-5 (Nielsen et al., 2000; Christoforidis and Zerial, 2001). Other proteins that localize to the endocytic compartment through FYVE domains include Smad anchor for receptor activation (SARA), which mediates TGFb signaling (Tsukazaki et al., 1998; Miura et al., 2000; Hayes et al., 2002), and hepatocyte-growth-factor-regulated tyrosine kinase substrate (Hrs), which is a homologue of yeast Vps27p and is involved in endosome/ lysosome trafficking and MVB formation (Odorizzi et al., 1998; Lloyd et al., 2002; Bache et al., 2003).

2.2 The Phosphoinositide 4-Kinases The mammalian genome contains four different genes encoding for PI4Ks: PI4KIIa, PI4KIIb, PI4KIIIa, and PI4KIIIb. The PI4Ks catalyze phosphorylation of PtdIns on the D-4 position of the inositol ring, hence leading to the production of PtdIns(4)P, which is the major precursor in the synthesis of the other phosphoinositides, including PtdIns(4,5)P2, PtdIns(3,4)P2, and PtdIns(3,4,5)P3. The PI4Ks have been classified into types II and III based on biochemical differences of the purified enzymes. The type II PI4Ks strongly associate with membranes, due to S-palmitoylation in a stretch of cysteines that has been conserved throughout evolution from yeast to higher eukaryotes. These can be distinguished from the type III PI4Ks by virtue of their lower Km for ATP and PtdIns, their insensitivity to wortmannin, and their sensitivity to inhibition by adenosine (Pike, 1992; Gehrmann and Heilmeyer, 1998). PI4K type II activity was initially isolated from plasma membranes, and also in association with the epidermal growth factor (EGF) receptor in A431 cells (Balla and Balla, 2006). In later immunocytochemical

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studies, PI4KIIa and PI4KIIb were localized to intracellular membranes, and mostly to the TGN and endosomes(Balla and Balla, 2006). Nevertheless, a fraction of PI4KIIa is present on plasma membranes under resting conditions, and PI4KIIb has been seen to be recruited onto plasma membranes in a Rac-dependent manner after growth-factor stimulation (Balla and Balla, 2006). Together with the finding that about 50% of PtdIns(4,5)P2 is produced via wortmannin-insensitive PI4Ks, this suggests that type II PI4Ks have a substantial role in sustaining the production of PtdIns(4,5)P2 at the plasma membrane. On the other hand, the predominant localization of type II PI4Ks on endomembranes has stimulated investigation of their roles in the production of PtdIns(4)P in TGN and endosomal membranes. Knock-down of PI4KIIa interferes with the PtdIns(4)P-dependent recruitment of the clathrin adaptor proteins AP-1 and Golgilocalized, g-ear containing, ARF-binding proteins (GGAs) to the Golgi complex (Wang et al., 2003; Wang et al., 2007). Interestingly, knockdown of PI4KIIa has been shown to reduce the association of PtdIns(4)P biosensors (e.g. the PH domains of FAPP1 and OSBP1) to the Golgi complex, while a complete cytosolic redistribution was seen only by combining PI4KIIa knock-down and wortmannin administration (Balla et al., 2005). Thus, it can be concluded that types II and III PI4Ks both contribute to the establishment of the Golgi complex pool of PtdIns(4)P. The yeast homologue of type II PI4Ks has been identified as the protein Lsb6p (Strahl and Thorner, 2007), which has been implicated in endosome mobility in yeast. However, its involvement in this process is independent of its catalytic activity and it appears rely mainly on the property of this protein to bind Las17p, a protein promoting actin filament polymerization (Strahl and Thorner, 2007). PI4KIIIa is mainly localized to ER membranes in mammals (Wong et al., 1997), although the overexpressed enzyme has also been found in the Golgi area (Nakagawa et al., 1996) and in the nucleus (Heilmeyer et al., 2003). PI4KIIIa interacts with the ER/Golgi localized PITP RdgBaI/Nir2, which presumably supplies the PI4KIIIa with its substrate (Aikawa et al., 1999). Despite its predominant localization to the early aspects of the secretory pathway, the role of PI4KIIIa in the ER remains to be clarified. Some evidence actually indicates a role for PI4KIIIa in PtdIns(4)P production at the plasma membrane (Balla et al., 2005). These data are supported by the intracellular localization of the yeast orthologe of PI4KIIIa, Stt4p. Stt4p is, indeed, mainly present at the plasma membrane, where its product, PtdIns(4)P, is metabolized to PtdIns(4,5)P2 by the yeast PtdIns(4)P 5-kinase Mss4p (Audhya and Emr, 2002). Accordingly, in an Stt4 temperature-sensitive mutant cell line, the levels of both PtdIns(4)P and PtdIns(4,5)P2 were decreased by 50% at the nonpermissive temperature (Audhya et al., 2000). The presence of Stt4p in the ER is not prominent in yeast; nevertheless, a functional connection between Stt4p and the ER-localized PtdIns(4)P phosphatase Sac1p has been proposed (Foti et al., 2001). Moreover, Sac1-null cells have an 8- to 12-fold increase in intracellular PtdIns(4)P, which can be restored to wildtype levels by inactivation of Stt4, but not of the PI4KII and PI4KIIIb orthologes Lsb6 and Pik1, respectively (Foti et al., 2001; Tahirovic et al., 2005). More recently, it has been shown that Sac1p relocalises to the Golgi complex under starving conditions, where it eliminates a PtdIns(4)P pool produced by Pik1p (Faulhammer et al., 2007). Although mammalian PI4KIIIb can also be found in the nucleus (de Graaf et al., 2002), it is primarily localized to the membranes of the Golgi complex (Wong et al., 1997), to where it is recruited by the GTP-bound form of the small GTPase ARF1 (Godi et al., 1999). It also interacts with, and is regulated by, the Ca2+-binding protein NCS-1(Zhao et al., 2001) and the 14-3-3 proteins (Hausser et al., 2006), and in the latter case, the interaction is regulated by a stimulatory phosphorylation of PI4KIIIb by protein kinase D (PKD) (Hausser et al., 2006). PI4KIIIb can, in turn, act on the small GTPase Rab11 and recruit it to the Golgi complex (de Graaf et al., 2004). The importance of this interaction network involving PI4KIIIb is highlighted by its conservation through evolution. Indeed, the yeast orthologe of PI4KIIIb, Pik1p, has been shown to interact with yeast Arf1p (Walch-Solimena and Novick, 1999), and with the yeast orthologes of NCS-1 Frq1p (Hendricks et al., 1999) and Rab11 Ypt31p (Sciorra et al., 2005). As all of these proteins are involved in some aspects of membrane trafficking at the Golgi complex, it has become clear that PI4KIIIb, and thus PtdIns(4)P, has an active role in organizing the function of the Golgi complex (De Matteis and D’Angelo, 2007). Several studies have, indeed, shown independently that Pik1p supplies the pool of PtdIns(4)P that is needed for Golgi-to-plasma-membrane membrane trafficking in yeast (Hama et al., 1999; Walch-Solimena and Novick, 1999; Audhya et al., 2000).

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The first evidence that the PI4Ks have roles in the secretory pathway in mammalian cells arose from the observation that expression of a dominant-negative, dead-kinase form of PI4KIIIb (PI4KD656A) induces alterations in the organization of the Golgi complex (Godi et al., 1999). PI4KIIIb, however, is responsible for only a fraction of the PtdIns(4)P generated at the Golgi complex. Indeed the Golgi complex possesses two PI4K activities: a basal type II activity due to PI4KIIa, and an ARF-induced type III activity, due to PI4KIIIb (Godi et al., 1999; Wang et al., 2003). The maintenance of the correct balance of PtdIns(4)P at the Golgi complex appears to be crucial for Golgi function, since the overexpression of a functional enzyme (wild-type PI4KIIIb) decreases the rate of TGN-to-cell-surface delivery of both influenza hemagglutinin (HA) and the temperature-sensitive variant of the G protein of vesicular stomatitis virus (VSVG) in MDCK cells. At the same time, the expression of the kinase-dead PI4KIIIb inhibits TGN-to-plasma-membrane transport of VSVG, while it stimulates TGN-toplasma-membrane delivery of the apical marker HA, possibly because of a defective incorporation of the apical cargo into membrane rafts (Bruns et al., 2002). As PtdIns(4)P is the natural precursor of PtdIns(4,5)P2, some of the effects seen when PtdIns(4)P metabolism is perturbed can be ascribed to altered levels of PtdIns(4,5)P2. Nonetheless, it has become clear over the last few years that PtdIns(4)P has its own function and effectors at the Golgi complex. Proteins that can bind PtdIns(4)P can be divided into two functional groups: those involved in coat assembly, and those involved in lipid transport and metabolism. In the latter, four proteins (FAPP1, FAPP2, OSBP1 and CERT) share a common PH domain that binds preferentially to PtdIns(4)P, which at least for OSBP1 and FAPP1 has been shown to bind ARF1(De Matteis and D’Angelo, 2007). These proteins are thus localized to the trans side of the Golgi complex through their PH domains, while both OSBP1 and CERT contain an ER-binding motif (the FFAT domain) in their sequences, which results in them being localized at the interface between the ER and the Golgi complex. All of the proteins belonging to this class share a common domain organization, with their PH domain at the N-terminus, and, for all but FAPP1, a second lipid-binding/ transferring domain in their C-terminal half (D’Angelo et al., 2007). CERT and FAPP2 have indeed been demonstrated to act as lipid-transfer proteins, with the substrates of ceramide and glucosylceramide, respectively (Hanada et al., 2003; D’Angelo et al., 2007). Through their lipid-transfer activities, CERT and FAPP2 sustain sphingomyelin synthesis (Hanada et al., 2003) and glycosphingolipids synthesis (D’Angelo et al., 2007; Halter et al., 2007), respectively, at the Golgi complex. As the binding of these proteins to the Golgi complex is regulated by PtdIns(4)P, phosphoinositide metabolism and, in particular, PtdIns(4)P production appear to be master regulators of sphingolipid metabolism (Toth et al., 2006; D’Angelo et al., 2007). Moreover, both CERT and FAPP2 have roles in vesicular trafficking between the TGN and the plasma membrane (Godi et al., 2004; Vieira et al., 2005; Fugmann et al., 2007), suggesting a link between sphingolipid metabolism and membrane trafficking. The PtdIns(4)P effectors include several proteins that are involved in coat assembly in vesicle budding. Indeed, EpsinR, the GGAs and AP-1, which participate in clathrin coat assembly at the TGN, have been shown to use PtdIns(4)P as a membrane receptor. Thus, the number and the nature of these PtdIns(4)P effectors at the Golgi complex explains the important morphological and functional effects that can be produced by PI4K malfunction on this organelle.

2.3 The Phosphatidylinositol Monophosphate Kinases The main product of the PtdIns monophosphate kinases (PIPKs) is PtdIns(4,5)P2, and although it only comprises about 1% of the phospholipids in the cytoplasmic leaflet of the plasma membrane, PtdIns(4,5)P2 is a key molecule in an astonishing number of cellular functions. PtdIns(4,5)P2 is the source of the second messenger Ins(1,4,5)P3, which induces the release of Ca2+ from intracellular stores, and diacylglycerol, which acts in combination with Ca2+ to activate protein kinase C (Michell, 1975; Berridge and Irvine, 1984, 1989). PtdIns(4,5)P2 is also the precursor of PtdIns(3,4,5)P3, which, as indicated above, acts as a second messenger through its contribution to the membrane recruitment and activation of many proteins. PtdIns (4,5)P2 is, per se, the binding site at the plasma membrane for many proteins containing PH domains and other phoshoinositide-interacting domains (McLaughlin and Murray, 2005). It also activates a number of

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ion channels at the plasma membrane, participates in phagocytosis, exocytosis, clathrin-mediated endocytosis, and synaptic vesicle trafficking. In addition, PtdIns(4,5)P2 is bound by many proteins that are involved in the regulation of the actin cytoskeleton, and it regulates cytoskeleton–plasma membrane adhesion (McLaughlin and Murray, 2005). Two classes of PIPKs that can phosphorylate the monophosphorylated phosphoinositides to produce PtdIns(4,5)P2 have been defined based on their sensitivity to phosphatidic acid (PA). The type I PIPKs are stimulated by PA, while the type II PIPKs are not (Moritz et al., 1992; Jenkins et al., 1994). The genes encoding these two families of enzymes have been cloned, and the gene products have been shown to selectively phosphorylate different positions on the inositol ring of the monophosphorylated phosphoinositides. The type I enzymes phosphorylate PtdIns(4)P in the D-5 position, to make PtdIns(4,5)P2, and are thus PIP5Ks, whereas the type II PIPKs phosphorylate PtdIns(5)P to make PtdIns(4,5)P2, and are consequently PIP4Ks (Rameh et al., 1997). Furthermore, broad in-vitro substrate specificities of these PIPKs have been reported: type I PIP5K can produce PtdIns(3,5)P2 and PtdIns(3,4,5)P3 from PtdIns(3)P and PtdIns(3,4)P2, respectively, and type II PIP4K can produce PtdIns(3,4)P2 by phosphorylating PtdIns(3) P (Fruman et al., 1998; Tolias et al., 1998; Anderson et al., 1999). Three genes (a, b, and g) of the mammalian type I PIPKs are indexed in databases, and the type Ig isoform exists as three splice variants, each of which has distinct functions, and possibly, subcellular distributions (Clarke et al., 2007). As with the type I PIPKs, the type II PIPKs are present as three different isoforms (a, b, and g) although with no currently known splice variants. A comparison of the primary sequences between the type I and type II PIPKs reveals sequence identities of only 28–33%, whereas across the isoforms within each subtype there is high homology (66–78% identical). These type I and type II PIPKs appear to be functionally nonredundant, even though they synthesize the same product, PtdIns(4,5) P2, and their quantitative contributions to the production of PtdIns(4,5)P2 are quite different. Pulselabeling experiments have suggested that the phosphorylation of PtdIns(4)P in position D-5 represents the major route for PtdIns(4,5)P2 synthesis (Stephens et al., 1991; Whiteford et al., 1997). Moreover, the limited amount of PtdIns(5)P in cells (about 2% of the total monophosphorylayed phosphoinositides) means that it is indeed an unlikely candidate for the supply of large quantities of PtdIns(4,5)P2. Nonetheless, type I and type II PIPKs localize to different intracellular compartments (type I, plasma membrane and nucleus; type II, cytosol, ER, nucleus and actin cytoskeleton, and not plasma membrane), leaving open the possibility that they produce distinct pools of PtdIns(4,5)P2. The type I PIPKs have been implicated in the regulation of secretion, endocytosis, and the actin cytoskeleton, and their activities can be regulated by small GTP-binding proteins, such as Rho, Rac, and ARF, which interact directly with type I PIPKs, but not with type II (Honda et al., 1999; Tolias and Carpenter, 2000; Aikawa and Martin, 2003). The first evidence of a role for PtdIns(4,5)P2 in exocytosis came from the identification of type I PIP5K as a required co-factor in the ATP-dependent priming step that precedes Ca2+-triggered secretion of dense-core vesicles (Hay et al., 1995). More recently, an important advance demonstrated that ARF6 has a role in dense-core vesicle exocytosis through the control of the activity of PIP5K, and hence synthesis of the required plasma-membrane pool of PtdIns(4,5)P2. The physiological functions of the type II PIPKs are not yet well defined. A specific type II isoform, type II PIPKb, interacts with the p55 subunit of the tumor necrosis factor-a (TNFa receptor, and may thus have a role in TNFa-mediated signaling (Castellino et al., 1997). Although none of the known PIP5K isoforms have been visualized in the Golgi complex, there is evidence that indicates that PtdIns(4,5)P2 has a role in ER-to-Golgi transport and in the formation/release of post-Golgi transport carriers. The former was deduced by the demonstration that the PtdIns(4,5)P2binding PH domains, such as those of the b-spectrins, inhibit ER-to-Golgi transport of VSVG in permeabilized NRK cells, potentially by inhibiting the association of spectrin with the Golgi complex and/or with pre-Golgi transport intermediates (De Matteis and Morrow, 1998; Godi et al., 1998). Moreover, by inhibiting PtdIns(4,5)P2 production with primary alcohols (which inhibit the synthesis of PA, an activator of PIP5K(Sweeney et al., 2002; Siddhanta et al., 2003)), it has been shown that PtdIns(4,5)P2 synthesis is required for the release of transport intermediates from the TGN and for maintaining the structural integrity and function of the Golgi complex, both in growth-hormone-secreting rat pituitary (GH3) cells and in isolated Golgi membranes (Siddhanta et al., 2000). In contrast, an increase in PtdIns(4,5)P2 levels

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(induced by PIP5Ka transfection) triggers the formation of actin comets on vesicular structures, some of which derive from the Golgi complex (Rozelle et al., 2000; Stamnes, 2002). Thus PtdIns(4,5)P2 might be required both for the formation of post-Golgi transport carriers and for their actin-dependent movement. PIKfyve is the mammalian homologe of the yeast lipid kinase Fab1p(Sbrissa et al., 1999), and it can phosphorylate PtdIns and PtdIns(3)P in the D-5 position of inositol, generating PtdIns(5)P and PtdIns (3,5)P2, respectively. PIKfyve has been localized to the late endocytic compartment due to the binding of its FYVE domain to PtdIns(3)P, which is enriched in this compartment. Recent studies using the expression of a kinase-dead PIKfyve point mutant and microinjection of specific antibodies have demonstrated a role for PIKfyve in controlling fluid-phase endocytosis through the regulation of MVB morphogenesis (Ikonomov et al., 2002; Sbrissa et al., 2002; Ikonomov et al., 2003). Similarly, in yeast, Fab1p has been implicated in vacuole homeostasis. Indeed, in fab1-null cells, a remarkable enlargement of the average vacuolar area has been seen, along with defects in vacuolar protein sorting and vacuole acidification (Strahl and Thorner, 2007).

3

The Phosphoinositide Phosphatases

The phosphoinositide phosphatases are divided into three major categories based on their hydrolysis of the 3-, 4- and 5-phosphorylated phosphoinositides.

3.1 The Phosphoinositide 3-Phosphatases The phosphoinositide 3-phosphatases include the tumor suppressor phosphatase and tensin homologue on chromosome ten (PTEN), its related protein transmembrane phosphatase with tensin homology (TPIP) and myotubularin (MTM), and the myotubularin-related proteins (MTMRs) (Maehama et al., 2001; Wishart et al., 2001). PTEN is a dual specificity (inositol lipid and tyrosine) phosphatase that is mutated in many sporadic human tumors (Downes et al., 2007). PTEN can use diverse substrates in vitro, ranging from phosphotyrosyls to the inositol phosphates and inositol phospholipids (Downes et al., 2007). The in vivo substrate specificity of PTEN appears to be determined by its weak binding to membranes via its C2 and N-terminal PtdIns(4,5)P2-binding motif. Thus, PTEN should cycle between a membrane-associated active state and a nonmembrane-associated inactive state (Downes et al., 2007). The net result of this is that PTEN uses the inositol phospholipids as preferred substrates in vivo, and PtdIns(3,4,5)P3 in particular; it can dephosphorylate all of the 3-phosphorylated phosphoinositides in vitro. The PTEN protein contains a phosphatase domain, a region with homology to tensin, and a PDZ protein-interaction domain. The PTEN protein phosphatase activity has been shown to be involved in many cellular functions, including cell-cycle progression, apoptosis, and cell contact and migration. On the other hand, it is generally accepted that many of the effects of PTEN inactivation in cancer depend on its lipid phosphatase activity that antagonizes PI3K signaling. Recently, use of the PTEN catalytic core motif to screen the Dictyostelium genome using the BLAST programmes identified a new protein in the Dictyostelium genome that has been named phospholipid-inositol phosphatase (PLIP). PLIP has a preference for PtdIns5P as substrate (Merlot et al., 2003) and has been localized to the Golgi complex, where it is thought to be involved in membrane trafficking. Other PTEN-related proteins have also been shown to be localized to the Golgi complex: transmembrane phosphatase with tensin homology (TPTE)(Guipponi et al., 2001; Walker et al., 2001), which is specifically expressed in testis, and TPTE- and PTEN-homologous inositol-lipid phosphatase (TPIP; Walker et al., 2001). However, the functions of TPTE and TPIP at the Golgi complex remain to be determined. The MTM phosphatases are members of the protein tyrosine phosphatase superfamily. They were initially thought to be protein phosphatases, but it was then demonstrated that recombinant myotubularin (MTM1) can remove the phosphate in position D-3 from PtdIns(3)P (Robinson and Dixon, 2006). Subsequently, several MTMRs were shown to hydrolyze PtdIns(3)P and also PtdIns(3,5)P2 (Robinson and Dixon, 2006). The MTMs are conserved amongst eukaryotes, with the human and zebrafish genomes containing the same 14 family members, suggesting that most vertebrates possess the

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same complement of MTMs (Robinson and Dixon, 2006). Metazoan MTMs have been grouped into six subclasses, with each single member expressed in invertebrates corresponding to each subclass (Robinson and Dixon, 2006). Strikingly, nearly half of the MTMs in metazoa are catalytically inactive, while Saccharmyces cerevisae appears to lack inactive family members. The roles of the catalytically inactive MTMRs are not known, but they have been proposed to function as adapters for the active forms (Kim et al., 2003; Nandurkar et al., 2003). The structural hallmarks of the MTMs are a PH-GRAM (pleckstrin homology, glucosyltransferases, Rab like GTPase activators, and MTMs) domain and a large protein tyrosine phosphatase domain. The different subclasses of the MTMs contain additional conserved protein domains, such as the FYVE, DENN (differentially expressed in normal vs. neoplastic), and PH domains. Overexpression studies have contributed to the validation of the function of the MTMs as inositol phosphatases in vivo, both via actual measurements of the phosphoinositides and via the monitoring of the abnormal endocytic compartment (Robinson and Dixon, 2006). Moreover, the overexpression of MTM1 leads to the release of the endosomal antigen EEA1 from endosomes (Robinson and Dixon, 2006). A critical and still unsolved issue relates to the localization of the single MTMs with regard to their known substrates. PtdIns(3)P has been reported to be enriched on early endosomes and on the internal vesicles of MVBs, while PtdIns(3,5)P2 appears to have functions at the late endosomes and lysosomes (Robinson and Dixon, 2006). There are indications that different MTMs localize to the plasma membrane, late endosomes, the Golgi complex, and the ER. Only recently was MTM1 shown to localize both on early and late endosomes under basal conditions, via its binding the PI3K complex VPS15/VPS34 and 3-phosphorylated phosphoinositides (Cao et al., 2007). Mutations in MTM1 result in the X-linked human disease known as myotubular myopathy, which is characterized by hypotonia and respiratory insufficiency. This disease affects about 1 in 50,000 new-born males, most of which die within the first month of life due to respiratory failure, although some do survive for several years. On the other hand, mutations in the MTMRs proteins MTMR2 and MTMR13/SBF2 (Kim et al., 2002) cause severe demyelinating neuropathies, such as CharcotMarie-Tooth disease types 4B1 and 4B2.

3.2 The Phosphoinositide 4-Phosphatases In the context of a database search among human phosphatase genes (Ungewickell et al., 2005) that use PtdIns(4,5)P2 as their major substrate in vitro, two phosphoinositide 4-phosphatases were cloned recently: types I and II. Accordingly, the levels of PtsIns(4,5)P2 in a cell line expressing the type I enzyme were reduced by 20%. While the physiological roles of the phosphoinositide 4-phosphatases remain under investigation, they have been shown to localize to the endosomes/lysosomes (Coronas et al., 2007). Furthermore, a phosphatase activity is associated with the Sac phosphatase domain in yeast Sac1p and Inp53p, and in the mammalian Sac1-3 and synaptojanins. These have phosphatase activities that are mainly directed toward PtdIns(4)P, PtdIns(3)P, and PtdIns(3,5)P2 in vitro. However, yeast Sac1 mutants have very high levels of PtdIns(4)P, thus indicating that in vivo Sac1p has a preference for PtdIns(4)P versus these other two substrates. Yeast strains with mutations in the Sac1 gene show an array of phenotypes, including inositol auxotrophy (Whitters et al., 1993), secretory defects in chitin deposition, disorganization of the actin cytoskeleton, and impairment of ATP uptake and protein translocation to the ER (Mayinger et al., 1995). Moreover, mutations in Sac1 can bypass the essential requirement for Sec14 (responsible for the major yeast PITP) in protein transport from the Golgi complex to the plasma membrane. The human homologue of Sac1 has been cloned, and it behaves as the yeast isoform in terms of its substrate specificity and its localization to the ER and Golgi complex (Rohde et al., 2003). Moreover, it has been shown that hSac1 interacts with the coatomer protein-I (COPI) complex; mutation of a putative COPI-binding motif (KXKXX) abolishes this interaction and results in the accumulation of hSac1 in the Golgi complex (Rohde et al., 2003). Recently, it was also shown that yeast Sac1p translocates from the ER to the Golgi complex under cell-starvation conditions, with the induced shutdown of cell proliferation (Faulhammer et al., 2007). After translocation, Sac1p eliminates the Pik1p-generated pool of PtdIns(4)P. In addition, the Pik1p/Frq1p complex (see above) is released from the Golgi complex under nutrient

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depletion, suggesting the existence of a synergistic loop between the PI4Ks and phosphoinositide 4-phosphatases that are responsible for the growth-dependent control of the Golgi phosphoinositides (Faulhammer et al., 2007).

3.3 The Inositol 5-Phosphatases The inositol 5-phosphatases comprise a large family of phosphatases that are defined by their catalytic domain of approximately 350 amino acids, which has homology with the apurinic/apyrimidic family of endonucleases (Astle et al., 2007). Inositol 5-phosphatases remove the phosphate in position D-5 from different membrane phosphoinositides, as well as from soluble inositol phosphates (Astle et al., 2007), and they have been classified into four types according to their substrate specificities. The type I 5-phosphatases can hydrolyze only water-soluble substrates (i.e., the inositol phosphates). The type II 5-phosphatases can use the lipid substrates, although not necessarily exclusively. These type II 5-phosphatases comprise a heterogeneous group of proteins that can be further divided into four subgroups based on their sequence conservation: the GAP-domain-containing 5-phosphatases, the Sac-1-homology-domain-containing 5-phosphatases, the proline-rich-domain-containing 5-phosphatases, and the skeletal-muscle and kidney-enriched inositol 5-phosphatase (SKIP). The type III 5-phosphatases SHIP1 and SHIP2 have SH2 domains and can hydrolyze the phosphate at the D-5 position of the phosphoinositides and inositol phosphates that also have a phosphate group at the D-3 position. Finally, the type IV 5-phosphatases hydrolyze only lipid substrates, such as PtdIns(4,5)P2 and PtdIns(3,4,5)P3. The only member of the type I family of 5-phosphatases, 5-phosphatase I (also known as the 43-kDa 5-phosphatase), can hydrolyze the soluble inositol phosphates Ins(1,4,5)P3 and Ins(1,3,4,5)P4 and can thus regulate intracellular calcium signaling. It has a CAAX motif in its C-terminal portion that indicates its localization to the plasma membrane, the site of Ins(1,4,5)P3 production (De Smedt et al., 1996). Consistent with this, ATP-induced cytosolic Ca2+ oscillations are abrogated in cells overexpressing 5-phosphatase I (De Smedt et al., 1997), while cells underexpressing this 5-phosphatase shows spontaneous Ca2+ oscillations in the absence of agonists, and increased sensitivity to agonists, together with increased basal levels of Ins(1,4,5)P3 (Speed et al., 1999). These increased levels of Ins(1,4,5)P3 correlate with a transformed cellular phenotype and tumor formation in nude mice (Speed et al., 1996). The GAP domain containing type II 5-phosphatases include OCRL and 5-phosphatase II. OCRL is the causative gene of the recessive X-linked inherited disorder in humans (Lowe, 2005) that is known as Lowe’s oculocerebrorenal syndrome, which is characterized by renal failure, growth and mental retardation, and cataracts (Lowe, 2005). The OCRL protein is ubiquitously expressed and has a central 5-phosphatase domain and a C-terminal RhoGap-like domain. OCRL can hydrolyze PtdIns(4,5)P2, Ins(1,4,5)P3, PtdIns (3,4,5)P3, and Ins(1,3,4,5)P4, with a preference for PtdIns(4,5)P2 (Schmid et al., 2004). OCRL has been mainly localized to endosomal structures and the TGN, where it is believed to regulate vesicular trafficking between the TGN and endosomes (Ungewickell et al., 2004). Moreover, OCRL binds the clathrin heavy chain, the a-adaptin subunit of AP-2, and APPL1(Ungewickell et al., 2004; Erdmann et al., 2007), and the knock-down of OCRL alters the intracellular distribution of AP-1 and the mannose 6-phosphate receptor (Choudhury et al., 2005). All of this has indicated a role for OCRL in the organization of the TGNendosomal/-lysosomal interface. Of note, the attempts to reproduce Lowe’s oculocerebrorenal syndrome in mice by knocking-out the OCRL gene have been unsuccessful, suggesting the presence of a compensation mechanism (Janne et al., 1998). As an ubiquitously expressed, 75-kDa protein, 5-phosphatase II shows catalytic activity against PtdIns (4,5)P2, Ins(1,4,5)P3, PtdIns(3,4,5)P3, and Ins(1,3,4,5)P4. It is encoded by the Inpp5b gene, and has a high degree of homology with OCRL. Of particular interest, INPP5B knock-out mice are healthy, while the combined knock-out of OCRL and INPP5B has resulted in an embryonically lethal phenotype, further supporting overlapping functions for these two phosphatases (Janne et al., 1998). On the other hand, in cultured cells, OCRL and 5-phosphatase II do not completely overlap in their subcellular distributions, with 5-phosphatase II also being present in earlier stations of the secretory pathway, including the ER-Golgi intermediate compartment (Williams et al., 2007). 5-Phosphatase II interacts with the GTP bound form of

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Rab5, and this interaction produces an increase in the 5-phosphatase activity toward PtdIns(3,4,5)P3, thus priming the Rab5-stimulated production of PtdIns3P. Moreover, the 5-phosphatase II knock-down promotes Rab5-dependent receptor-mediated endocytosis (Shin et al., 2005). Synaptojanins-1 and -2, hSac2 and hSac3/Fig4 are 5-phosphatases that have a Sac-1 homology domain. Synaptojanin-1 was initially isolated through its binding to the SH3 domain of Grb2 (McPherson et al., 1994), and then it was later characterized as the major presynaptic 5-phosphatase in rat brain (McPherson et al., 1996). The substrate specificity of synaptojanin-1 is reported as 5-phosphate hydrolysis of PtdIns(4,5)P2, Ins(1,4,5)P3, PtdIns(3,4,5)P3, and Ins(1,3,4,5)P4 (McPherson et al., 1996). The enzymatic activity of the synaptojanins is not, however, restricted to this 5-phosphatase activity; indeed, the presence of the Sac1 homology domain on the N-terminal portion of the synaptojanins renders them active in hydrolyzing PtdIns(3)P, PtdIns(4)P, and PtdIns(3,5)P2 (Guo et al., 1999). Synaptojanin-1 forms molecular complexes with a number of proteins that are involved in synaptic vesicle recycling, including dynamin, amphiphysins I and II, endophilin, syndapin, Eps15, AP-2, and Dap160/intersectin (Astle et al., 2007). Synaptojanin-1 knock-down and depletion studies have highlighted a role for this phosphatase in clathrinmediated endocytosis, which appears to be tightly regulated by the local and dynamic levels of PtdIns(4,5) P2. Synaptojanin-2 shows good sequence homology with synaptojanin-1, and contains an N-terminal Sac1 homology domain and a central 5-phosphatase domain. Nevertheless, synaptojanins-1 and -2 diverge at the C-terminus, with synaptojanin-1 containing a proline-rich domain and an AP-2 binding domain that are absent in synaptojanin-2. Synaptojanin-2 has also been shown to be involved in endocytosis. Knock-down of synaptojanin-2 results in endocytic defects, with a decrease in the numbers and sizes of vesicles in all stages of endocytosis (Rusk et al., 2003). Moreover, synaptojanin-2 interacts with the GTP-bound form of Rac1, and localizes to the plasma membrane when co-transfected with dominant-positive Rac1, and also under EGF stimulation (Malecz et al., 2000; Nemoto et al., 2001). hSac2 is a Golgi-localized, nonconventional 5-phosphatase, as it lacks the 5-phosphatase domain that characterizes this family of enzymes. The 5-phosphatase activity of hSac2 resides in its Sac domain, which has a Km of 14.3 mM against PtdIns(4,5)P2 (Minagawa et al., 2001), which is comparable to that of other type II 5-phosphatases. Similar to hSac2, hSac3/Fig4 has also been shown to have a 5-phosphatase activity that is associated with its Sac domain, which uses PtdIns(3,5)P2 as substrate, thus producing PtdIns(3)P. In yeast, Fig4 has been shown to interact with the Fab1 activator Vac14, suggesting the existence of an enzymatic loop for the regulation of PtdIns(3,5)P2/PtdIns(3)P turnover (Duex et al., 2006). Recently, it has been shown that in mammals Sac3/Fig4 is the causative gene for neurodegeneration in the pale-tremor mouse and in patients with Charcot-Marie-Tooth disorder type 4J (Chow et al., 2007). Proline-rich inositol polyphosphate 5-phosphatase (PIPP) belongs to the proline-rich domain containing type II 5-phosphatases. PIPP is a 107-kDa 5-phosphatase that is active in hydrolyzing PtdIns(4,5)P2, Ins (1,4,5)P3, PtdIns(3,4,5)P3, and Ins(1,3,4,5)P4 (Mochizuki and Takenawa, 1999). It contains proline-rich domains at its N and C termini, with a central 5-phosphatase domain and a recently described SKICH domain, which mediates its constitutive association with the plasma membrane (Gurung et al., 2003). In neuronal cells, PIPP has been shown to negatively regulate neurite outgrowth after NGF stimulation (Ooms et al., 2006), while in differentiating rat pheocromocytoma cells, PIPP is enriched in the neuronal growth cone. The knock-down of PIPP in these cells produces hyper-elongated neurites after NGF stimulation (Ooms et al., 2006). SKIP is a 51-kDa 5-phosphatase that is ubiquitously expressed and is active on PtdIns(4,5)P2, Ins(1,4,5)P3, PtdIns(3,4,5)P3, and Ins(1,3,4,5)P4. In nonstimulated cells, it localizes in the perinuclear area, co-localizing with ER markers, while upon stimulation it translocates to plasma membrane ruffles (Gurung et al., 2003). This translocation to the plasma membrane is mediated by a C-terminal domain known as SKICH. SKIP overexpression decreases actin stress fibers, but does not affect the extent of membrane ruffling upon stimulation (Ijuin et al., 2000). SKIP is also one of the genes implicated in a contiguous gene syndrome, Miller–Dieker syndrome, a severe form of lissencephaly that is caused by defects in neuronal migration (Kato and Dobyns, 2003). The type III 5-phosphatases SHIP1 and SHIP2 contain an N-terminal SH2 domain, a central 5-phosphatase domain, and a C-terminal proline-rich domain. Moreover, the C-terminal region of

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SHIP2 has a number of protein-interaction motifs: two WW binding motifs, an NPXY motif, and a sterile alpha motif (SAM) domain. SHIP1 has a significant role in regulation of the immune system and in cell polarity, and it is required for normal cell motility and chemotaxis (Nishio et al., 2007); SHIP2 has a role in insulin signaling and obesity regulation. These two phosphatases show preferential activities against inositol phospholipids and inositol phosphates with a phosphate in position D-3. SHIP2 localizes to the cytosol in resting cells, while it translocates to plasma-membrane ruffles during cell attachment and under growthfactor stimulation. Moreover, SHIP2 forms complexes with proteins involved in actin dynamics (Dyson et al., 2001; Prasad et al., 2001). The 72-kDa 5-phosphatase, which is also known as pharbin (in rat) or type IV 5-phosphatase, is the most active phosphatase for hydrolysis of PtdIns(3,4,5)P3 to PtdIns(3,4)P2, although it shows no activity against the soluble inositol phosphates. The over-expression of this peri-Golgi-localized 72-kDa 5-phosphatase results in a rapid depletion of both cellular PtdIns(3,4,5)P3 and PtdIns(4,5)P2 (Kisseleva et al., 2002). Recently it was reported that 72-kDa 5-phosphatase inhibits FcgR-mediated phagocytosis, affecting pseudopod extension and phagosome closure, possibly via control of the amplitude and duration of PtdIns(3,4,5)P3 levels at the phagocytic cup (Horan et al., 2007).

4

Concluding Remarks

Eukaryotic cells are composed of a system of sealed organelles that are formed from internal membranes, and that are connected by an intense flux of material. Nevertheless, these subcellular organelles maintain their identities by regulating their composition, in terms of their resident proteins and lipids. The phosphoinositides have major roles in this maintenance of subcellular organelle identity, through their actions as short-lived topological determinants that can recruit effector proteins from the cytosolic environment to specialized membrane interfaces (> Figure 11-2). Indeed, it has been proposed that together with GTPases, the phosphoinositides provide a combinatorial code of spatio-temporal signals that regulate the subcellular anatomy (Behnia and Munro, 2005). Within this framework, the extreme plasticity of phosphoinositide metabolism also explains why they have evolved as the controllers of the activation state of cells. More in general, the inositol phospholipids and phosphates appear to act as universal molecular switches in the spatial and temporal regulation of a wide variety of intracellular phenomena.

Acknowledgments We thank C. P. Berrie for editorial assistance; and E. Fontana for artwork. The authors acknowledge the support of Telethon and AIRC.

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Lipid Mediators and Modulators of Neural Function: Lysophosphatidate and Lysolipids

D. N. Brindley . A. U. Bra¨uer

1

General Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 290

2 2.1 2.2 2.3 2.4

Signaling by Extracellular Lysophosphatidate and Lysophosphatidylcholine . . . . . . . . . . . . . . . . . . . 290 LPC Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 290 Signaling Through LPA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 290 Lysophosphatidate Receptor Expression During Brain Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 291 Lysophosphatidate Receptor KO Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 293

3 3.1 3.2 3.3 3.4 3.5

Metabolism of Extracellular Lysophosphatidylcholine and Lysophosphatidate . . . . . . . . . . . . . . . . . 293 Synthesis of Extracellular Lysophosphatidylcholine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 293 Role of Autotaxin and the Synthesis of Extracellular Lysophosphatidate . . . . . . . . . . . . . . . . . . . . . . . . . 293 Autotaxin Expression in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 294 Autotaxin Expression During Neurological Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 294 Autotaxin Knockout Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 295

4

Role of Secretory Phospholipase A2 in the Synthesis of Extracellular Lysophosphatidate . . . . . . . 295

5 5.1 5.2 5.3 5.4 5.5

Function of Lipid Phosphate Phosphatases in Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 295 Extracellular Dephosphorylation of LPA by Lipid Phosphate Phosphatases . . . . . . . . . . . . . . . . . . . . . . 295 Intracellular Functions of the Lipid Phosphates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 296 Intracellular Functions of the Lipid Phosphate Phosphatases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 296 Lipid Phosphate Phosphatase Expression in the Central Nervous System . . . . . . . . . . . . . . . . . . . . . . . . 298 Lipid Phosphate Phosphatases and Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 299

6

Importance of Phosphatidate Phosphatases-1 (Lipin) Activities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 299

7 7.1 7.2 7.3 7.4 7.5

Neurobiological Effects of LPA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 300 Tau Hyperphosphorylation by LPA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 300 LPA in Myelination Process . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 300 LPA During Brain Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 301 Effects of LPA on Brain Tumors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 301 Effects of LPA on Epileptic Seizure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 301

8 8.1 8.2 8.3

Plasticity-Related-Genes (Lipid Phosphate Phosphatase-Related Proteins) . . . . . . . . . . . . . . . . . . . . . 302 PRG Expression in the Adult Central Nervous System and During Development . . . . . . . . . . . . . . . 302 PRG Response to LPA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 302 PRG Expression After Brain Damage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 303

9

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 303

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_12, # Springer ScienceþBusiness Media, LLC 2009

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List of Abbreviations: ATX, autotaxin or lysophospholipase D; CNS, central nervous system; C1P, ceramide 1-phosphate; C20:4, arachidonic acid; DAG, diacylglycerol; EAE, experimental autoimmune encephalomyelitis; EPMR, epilepsy with mental retardation; LCAT, lecithin:cholesterol acyltransferase, LPA, lysophosphatidate; LPC, lysophosphatidylcholine; MAG, monoacylglycerol; MS, multiple sclerosis; PA, phosphatidate; PLD, phospholipase D; PRG, plasticity related genes; S1P, sphingosine 1-phosphate; 2-AG, sn-2-monoarachidonoylglycerol

1

General Introduction

There is an increasing body of knowledge that demonstrates important roles for lysophosphatidate (LPA) and other biologically active lipids in controlling cell signaling including regulation of cell division, death, and movement. These processes are important aspects of brain development, repair, and the normal functioning of the brain. We now know that LPA activates at least five G-protein coupled receptors. The signal that cells receive depends upon the G-proteins that couple to the receptors and the relative expression of various receptors on different cells. Additionally, the role of LPA in controlling signal transduction is regulated by enzymes that are responsible for its turnover and also for controlling signaling events downstream of receptor activation. This review will, therefore, discuss how synthesis of extracellular LPA from lysophosphatidylcholine (LPC) and degradation of LPA to monoacylglycerol (MAG) are regulated to modulate signal transduction in various cells. The discussion will also emphasize, where possible, how these processes control the responses of the brain during its development, normal functioning, and repair.

2

Signaling by Extracellular Lysophosphatidate and Lysophosphatidylcholine

2.1 LPC Receptors LPC was reported to signal through G-protein coupled receptors such as G2A (G2 accumulation) and GPR4 (G protein-coupled receptor-4) (Kabarowski et al., 2001; Zhu et al., 2001; Rikitake et al., 2002; Lin and Ye, 2003; Radu et al., 2004; Kim et al., 2005) (> Table 12-1). However, some of this work has been retracted (Zhu et al., 2001; Witte et al., 2005). G2A and GPR4 are now thought to be proton-sensing receptors (Murakami et al., 2004; Tomura et al., 2005). Antagonism of these receptors by high mM concentrations of lysolipids was thought to result from nonspecific effects (Seuwen et al., 2006). Thus some of the reported effects of LPC could be mediated indirectly after the metabolism of LPC to other bioactive lipids.

2.2 Signaling Through LPA Receptors LPA is present in biological fluids at mM concentrations and it is a potent growth factor. LPA mediates signals through five G-protein-coupled receptors (> Table 12-1, > Figure 12-1) (Guo et al., 1996; Fukushima et al., 1998; Goetzl and An, 1998; Bandoh et al., 1999; Lynch, 2002; Noguchi et al., 2003; Lee et al., 2006). Coupling through Gai decreases cAMP concentrations and causes activations of phosphatidylinositol 3-kinase and Ras; G12/13 stimulates phospholipase D (PLD) and Rho leading to stress fiber formation; Gq activates phospholipase C, Ca2+ transients and protein kinase C isoforms (English et al., 2001; English et al., 2002) (> Figure 12-1). Activated LPA receptors also transactivate receptor tyrosine kinases (Daub et al., 1996; Luttrell et al., 1997; Herrlich et al., 1998; Prenzel et al., 1999; Goppelt-Struebe et al., 2000; English et al., 2002; Wu and Cunnick, 2002; Yart et al., 2002; Wang et al., 2003) through the release of bg-subunits, activation of Src, matrix metalloproteinases, or PLD2 (Wang et al., 2003). Recent work shows that part of the action of LPA in promoting cell growth is through its receptor-mediated effects in decreasing the nuclear localization and cellular abundance of the tumor promoter, p53 (Murph et al., 2007).

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. Table 12-1 Ligands, tissue distribution and G-protein coupling of LPA and LPC receptors Receptor designation LPA1 (Edg-2, vzg-1) (Hecht et al., 1996; Contos and Chun, 1998) LPA2 (Edg-4) (Contos et al., 2000) LPA3 (EDG-7) (Contos et al., 2000; Contos and Chun, 2001) LPA4 (p2y9/GPR23) (Noguchi et al., 2003; Lee et al., 2007) LPA5 (GPR92) (Kotarsky et al., 2006; Lee et al., 2006) G2A (Bolick et al., 2007; Meyer zu Heringdorf and Jakobs, 2007) GPR4 (Lum et al., 2003; Meyer zu Heringdorf and Jakobs, 2007)

High affinity ligand LPA

Organ distribution in adult mice CV, GI, CNS and PNS, Gonadal tissue, lung heart, spleen, moderate levels in kidney, thymus, stomach and muscle Testicular tissue, kidney, low levels in brain, heart, lung spleen, thymus, and stomach Kidney, testis, lung and in low levels in small intestine, heart, spleen, thymus, stomach

Expression during brain development Embryonic expression in the ventricular zone during cortico neurogenesis and in the white matter in postnatal stages High level of expression during embryonic expression

G-protein coupling Gi, G12/13, Gq/11

High expression around birth

Gi, Gq/11

LPA

Heart, skin, thymus, bone marrow

Embryonic brain, embryonic stem cells

Gq/11, G12/13

LPA

Small intestine, Moderate levels in skin, spleen, stomach, thymus, lung and liver

Embryonic brain, embryonic stem cells, dorsal root ganglia (DRG)

Gq/11, G12/13

LPC and/or SPC, proton SPC and/or LPC, proton

Predominantly in lymphoid tissue, Endothelium

Not detected

Gi, Gs, Gq; G 12/13

Endothelium

Not detected

Gi

LPA > PA LPA

Gi, Gq/11, G12/13

CV, cerebrovascular tissue; GI, gastrointestinal tissue; AEC, aortic endothelial cells; CNS, central nervous system tissue; EC, endothelial cells; LPC, lysophosphatidylcholine, lymph, lymphocytes; LPA, lysophosphatidic acid; PA, phosphatidate; S1P, sphingosine 1-phosphate; SPC, sphingosylphosphorylcholine

This LPA effect involves activation of phosphatidylinositol 3-kinase, which increases the proteosomal degradation of p53 and thereby decreases p53-mediated transcriptional regulation.

2.3 Lysophosphatidate Receptor Expression During Brain Development LPA1 (vzg-1/edg-2) is strongly expressed during embryonic cerebral cortex development, where it is enriched in the ventricular zone (Hecht et al., 1996). The expression of LPA1 receptor declines shortly before birth. At some point during postnatal life up to adulthood, LPA1 expression reemerges in a different cellular brain region, namely in and around white matter tracts. In situ hybridization analysis has shown LPA1 mRNA expression in oligodendrocytes, which are the myelinating glia cells in the brain (Weiner et al., 1998), as well as in Schwann cells, the peripheral nervous system’s myelinating cells (Weiner and Chun, 1999).

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. Figure 12-1 LPA mediated signaling through a specific class of G-protein coupled receptors. LPA induces diverse cellular responses including proliferation, retraction, cell survival, migration, proliferation, differentiation by binding to multiple G-protein-coupled receptors. SRF, serum response factor; Rock, Rho-associated kinase; PI3K, phosphoinositide 3-kinase; Akt, protein kinase B; cAMP, cyclic adenosine monophosphate; p42/p44 MAPK, p42/p44 mitogen-activation protein kinase; PLC, protein kinase C; IP3, inositol 1,4,5-triphosphate; DAG, diacylglycerol; AC, adenylyl cyclase

LPA2 (edg-4) is heavily expressed in embryonic mouse brain, whereas in adult mice LPA2 is only detectable in testes and kidney (Contos et al., 2000). In situ hybridization studies suggest mRNA expression in postmitotic neuronal regions, such as cortical plate and ventricular zone of embryonic mice (Fukushima et al., 2001). LPA3 (edg-7) exhibits the highest mRNA expression levels around birth in mouse CNS, whereas only a low level of transcripts have been found during embryonic development and in adult brain (Contos et al., 2000). In adult mice, LPA3 mRNA expression is detected by RT-PCR in kidney, testes, lung, and to a lesser extent in small intestine, heart, spleen, thymus, and stomach (Contos and Chun, 2001). LPA4 (p2y9/GPR23) mRNA is expressed in adult mouse heart, skin, and thymus, as well as in embryonic brain and embryonic stem cells as analysed by northern blotting (Lee et al., 2007).

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LPA5 (GPR92) is mainly detected by northern blot analyses in adult mouse small intestine, spleen, lung, skin, thymus, and stomach, as. LPA5 mRNA is also expressed strongly in embryonic brain, embryonic liver, and embryonic stem cells (Lee et al., 2006).

2.4 Lysophosphatidate Receptor KO Mice LPA1( / ) mice show 50% lethality in the neonatal phase due to impaired suckling, attributable to defective olfaction (Contos et al., 2000). Furthermore, a loss of LPA responsiveness in embryonic cerebral cortical neuroblasts has been shown (Claesson-Welch, 1994). Interestingly, no specific abnormalities were detected in histological sections of brains (Contos et al., 2000). Apart from these effects, the surviving mice show decreased body size, craniofacial dysmorphism and increased apoptosis in the peripheral nerves. Additionally, a low incidence of frontal hematoma in the perinatal pups was observed (Contos et al., 2000). LPA2( / ) mice displayed no obvious phenotypic abnormalities (Contos et al., 2002). LPA1( / )/ LPA2( / ) double knockout mice developed without any apparent additional phenotypes compared with LPA1( / ) (Contos et al., 2002). LPA3( / ) mice have a significantly reduced litter size, which could be attributed to delayed implantation and altered embryo spacing. This is associated with a dramatic decrease in the prostaglandins and prostaglandin-endoperoxide synthase 2 expression levels. Exogenous administration of prostaglandins rescued the delayed implantation but did not prevent defects in embryo spacing, suggesting that prostaglandins have a role in implantation downstream of LPA3 signaling (Ye et al., 2005). Results for LPA4 and LPA5 knockout mice have not yet been reported.

3

Metabolism of Extracellular Lysophosphatidylcholine and Lysophosphatidate

3.1 Synthesis of Extracellular Lysophosphatidylcholine LPC is a major plasma lipid that is bound to albumin (Brindley, 1993) and it is the major precursor of extracellar LPA. Physiological concentrations of LPC are different between tissues and body fluids, in general, the concentration range from 5 to 180 mM. There are two major routes for the production of extracellular LPC. The first is through the action of lecithin:cholesterol acyltransferase (LCAT), which is found in circulating high-density lipoproteins. LCAT transfers the fatty acid from postion-2 of phosphatidylcholine to cholesterol to produce cholesterol ester and LPC. This LPC contains mainly saturated fatty acids that are normally preferentially located at position-1 of phosphatidylcholine. However, a large proportion of circulating LPC is polyunsaturated (Brindley, 1993) and this indicates another route for the production of extracellular LPC. At least part of the polyunsaturated LPC is derived from secretion by the liver. However, it is also possible that other organs and cell types could be involved in producing circulating LPC. Hepatocytes secrete LPC that contains a large proportion of arachidonate (C20:4) (Mangiapane and Brindley, 1986; Graham et al., 1988a, b; 1991; Brindley, 1993). It was originally postulated that the production of LPC by the liver might represent a novel transport system for delivering choline and polyunsaturated fatty acids to the brain (Brindley, 1993). Although, this is still probably the case, we now know that LPC is a major precursor of extracellular LPA, which activates important G-protein coupled receptors that control signal transduction in the brain.

3.2 Role of Autotaxin and the Synthesis of Extracellular Lysophosphatidate A major route for the synthesis of extracellular LPA is through the actions of a secreted lysophospholipase D (> Figure 12-2) that is also known as autotaxin (ATX) (Saulnier-Blache, 2004). This enzyme is important in

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. Figure 12-2 Metabolism of extra-cellular arachidonoyl-(C20:4)lysophosphatidylcholine to bioactive lipids. LPA, 2-AG, and eicosanoids activate their respective receptors as indicated by the dashed arrows. DAG = diacylglycerol: MAG = monoacylglycerol

cancer since it is produced by tumor cells and it is implicated in tumor progression, metastasis, and angiogenesis (Tokumura et al., 2000; Nam et al., 2001; Moolenaar, 2002; Umezu-Goto et al., 2002). These actions are probably caused mainly by the production of LPA, which then provides a powerful signal for stimulating cell migration and division through its receptors. These latter actions of LPA are also involved in wound repair and development of tissues. ATX plays an important physiological role in these processes. CNS expresses high amounts of an isoform of ATX that is associated almost exclusively with oligodendrocytes, which are involved in myelin formation (Dennis et al., 2005). ATX can also convert sphingosylphosphorylcholine to sphingosine 1-phosphate (S1P) (Clair et al., 2003), although the physiological importance of this in S1P production through ATX is not firmly established. S1P is a sphingolipid that is analogous to LPA. S1P also activates a family of G-protein-coupled receptors and it is especially important for the regulation of angiogenesis and immune responses. The activity of ATX appears to be partly regulated by product inhibition through LPA and S1P (van Meeteren et al., 2005). In addition to its catalytic activity, ATX decreases adhesions of oligodendrocytes to the extracellular matrix through its C-terminal region. This action facilitates morphological remodeling (Dennis et al., 2005). It was, therefore, suggested that ATX is a matrixcellular protein that participates in the regulation of myelination by a novel signaling pathway leading to changes in integrin-dependent focal adhesion assembly and consequently oligodendrocyte interactions with the extracellular matrix (Fox et al., 2004).

3.3 Autotaxin Expression in the Brain ATX is expressed in the proliferating subventricular and choroid plexus epithelium during embryonic development. After birth, ATX is found mainly in white matter areas in CNS. In adult brain, ATX is expressed in leptomeningeal cells, oligodendrocyte precursor cells, choriod plexus, ciliary, iris pigment, retinal pigment epithelial cells, and cells of the CNS vasculature (Narita et al., 1994; Fuss et al., 1997; Sato et al., 2005; Savaskan et al., 2007).

3.4 Autotaxin Expression During Neurological Diseases ATX protein levels are elevated by about 85% in the cerebrospinal fluid of patients at time of initial diagnosis of relapsing-remitting type multiple sclerosis (MS), the most common form of MS (Hammack et al., 2004). People with this type of MS show episodes of acute decline in neurologic function, followed by partial or complete recovery periods free of disease progression. In contrast, in experimental autoimmune encephalomyelitis (EAE) mice, ATX expression levels are decreased in the brain and spinal cord at the onset of clinical symptoms, i.e., prior to signs of demyelination (Fuss et al., 1997). The downregulation of ATX expression in CNS at early stages of EAE has been discussed as an impairment of oligodendrocyte function

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before detectable damage to the myelin sheath occurs (Fuss et al., 1997). This difference in results between the two studies suggests that the function of ATX depends on the stage/episode of the disease. A detailed analysis of ATX expression in the course of the disease would contribute greatly to a better understanding of the function of ATX in MS. ATX is massively upregulated at the lesion site and in the denervated area following acute traumatic brain injury in the mouse (Savaskan et al., 2007). This lesion-induced increase is restricted to the site of incision and to the area of axonal degeneration, whereas nonaffected brain regions did not show any altered expression. Double immunocytochemistry revealed that many ATX-positive cells were also positive for the astrocyte marker, glial fibrillary acidic protein. These results emphasize the importance of the role of ATX and LPA in repair processes in the brain.

3.5 Autotaxin Knockout Mice ATX-deficient mice die at embryonic day 9.5 with profound vascular defects in the yolk sac and embryo. At E8.5, ATX-deficient embryos showed allantois malformation, neuronal tube defects and asymmetric headfolds. The onset of these abnormalities coincided with increased expression of ATX and LPA receptors that occurs in normal embryos. ATX heterozygous mice appear healthy but show half of the normal ATX activity and plasma LPA levels (Tanaka et al., 2006; van Meeteren et al., 2006).

4

Role of Secretory Phospholipase A2 in the Synthesis of Extracellular Lysophosphatidate

A second pathway for the production of extracellular LPA is through the action of secretory phospholipase A2 (sPLA2). The LPA synthesized by this route should be mainly saturated and it is particularly important in inflammatory conditions. sPLA2 hydrolyzed phospholipids including phosphatidate in membrane microvesicles that are shed from cells during inflammation (Fourcade et al., 1995). The authors concluded that upon loss of phospholipid asymmetry, cell-derived microvesicles provide a preferential substrate for sPLA2. Sphingomyelin hydrolysis, which is provoked by various cytokines, regulates sPLA2 activity and facilitates LPA production.

5

Function of Lipid Phosphate Phosphatases in Signaling

5.1 Extracellular Dephosphorylation of LPA by Lipid Phosphate Phosphatases Extracellular LPA is degraded by a family of LPPs (> Figure 12-2). These enzymes are also able to dephosphorylate wide variety of other lipid phosphates including extracellular sphingosine 1-phosphate (S1P). There are three different LPP isoforms and a splice variant. The LPPs belong to a large phosphatase family that includes Wunen proteins in Drosophila (Brindley and Waggoner, 1998; Brindley, 2004; Sigal et al., 2005). LPPs possess six transmembrane domains, three conserved active site domains and a glycosylation site that is on an external hydrophilic loop between the first and second catalytic sites (Brindley, 2004). The active site of the LPPs is on the outer surface of plasma membranes, or the luminal surface of internal membranes (Zhang et al., 2000; Brindley, 2004). The ecto-phosphatase activity of the LPPs can theoretically control the concentrations of SIP and LPA outside the cell and counteract the effect of increased ATX activity. LPP1 over-expression increased ectoLPP activity and attenuated the activation of ERK, PLD, Ca2+ transients, and cell division by exogenous LPA (Jasinska et al., 1999; Pilquil et al., 2001b). In other work, gonadotropin-releasing hormone (GnRH) increased ecto-LPP expression in ovarian cancer cells and this explained the anti-proliferative effects of GnRH (Imai et al., 2000). LPP3 over-expression decreases growth, survival, and tumorigenesis of ovarian

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cancer cells by increasing exogenous LPA degradation (Tanyi et al., 2003). Exogenous LPA increases ectoLPP1 activity in platelets and this decreases further LPA accumulation and LPA-induced shape changes and aggregation (Smyth et al., 2003). Ecto-LPP activities also regulate extracellular LPA accumulation and proliferation of pre-adipocytes (Simon et al., 2002). This combined work establishes a role for the ecto-LPP activities in regulating cell signaling by extracellular lipid phosphates. A further dimension for the role of the ecto-activities of the LPPs is through the product that is formed. Dephosphorylation of LPA yields monoacylglycerols (MAG). These compounds are not bioactive except for 2-arachidonoylglycerol (2-AG), which is an endogenous activator of cannabinoid (CB1 and CB2) receptors (> Figure 12-2) (Hillard, 2000; Sugiura and Waku, 2000; Gokoh et al., 2005). The role of LPPs in producing 2-AG may be physiologically important since a significant proportion of the circulating LPC contains arachidonate (Brindley, 1993). Its metabolism by autotaxin would then yield archidonoyl-LPA (> Figure 12-2). Therefore, ecto-LPP activities in the brain could contribute to the production of an endocannabinoid, in addition to other pathways that have been described which lead to the activation of cannabinoid receptors in the brain (Song and Zhong, 2000; Sugiura and Waku, 2000; Freund et al., 2003; Kishimoto et al., 2003, 2005; Basavarajappa, 2007). Also, 2-AG can be metabolized to arachidonate that is a precursor for formation of eicosanoids, which are also important regulators of cell activation (> Figure 12-2). The other consequence of LPP action on extracellular LPA, S1P, phosphatidate (PA), and ceramide 1-phosphate (C1P) is that the products, MAG, sphingosine, diacylglycerol (DAG) and ceramide, respectively, can far more readily enter the cell than the equivalent phosphate esters (Sciorra and Morris, 2002; Sigal et al., 2005; Zhao et al., 2007). Sphingosine, DAG, and ceramide are themselves bioactive and they can, therefore, regulate cell signaling. In addition, they are also rapidly rephosphorylated inside the cell by specific kinases. In the case of human lung endothelial cells, increased expression of LPP1 degrades S1P and results in increased uptake of sphingosine that is converted to internal S1P by sphingosine kinase (Zhao et al., 2007). This generation of internal lipid phosphates subsequent to ecto-activity of LPPs generates important intracellular signaling molecules as illustrated in > Table 12-2. Intracellular lipid phosphates are also generated directly in response to activation of cell-surface receptors.

5.2 Intracellular Functions of the Lipid Phosphates Activation of G-protein coupled receptors (e.g., LPA receptors) and receptor tyrosine kinases stimulates phospholipase D activities producing PA (> Figures 12-1 and > 12-3). PA activates a variety of intracellular signaling targets (> Table 12-2) including ERK, mTOR, and sphingosine kinase-1 thus increasing S1P formation. S1P activates ERK and it stimulates cell division and protects against apoptosis (> Figure 12-3). S1P also stimulates the mobilization of intracellular Ca2+. PA can also be metabolized to LPA (> Figure 12-3). Intracellular LPA can activate nuclear LPA1 receptors that regulate pro-inflammatory gene expression (Gobeil et al., 2003; Moughal et al., 2004). Intracellular polyunsaturated LPA was also claimed to stimulate PPARg receptors (McIntyre et al., 2003; Siess and Tigyi, 2004; Zhang et al., 2004). This conclusion has been questioned since LPA did not stimulate the transcription of PPARg-sensitive genes in pre-adipocytes (Simon et al., 2005). The LPPs can also degrade C1P, which is involved in inflammation. C1P production activates PLA2, to release C20:4 (Pettus et al., 2003a, 2004) and S1P activates COX-2 thus coordinately leading to eicosanoid production (Pettus et al., 2003b, 2005).

5.3 Intracellular Functions of the Lipid Phosphate Phosphatases LPPs are also expressed on internal membranes (Brindley, 2004; Sigal et al., 2005; Morris et al., 2006). Intracellular LPPs can control intracellular concentrations of PA, LPA, C1P, and S1P (> Figure 12-3) and thereby signaling targets as listed in > Table 12-2. The Pyne Group (Alderton et al., 2001; Long et al., 2005) showed that LPPs control ERK activation by extracellular thrombin as well as LPA, S1P, PA. These effects correlated with decreases in intracellular PA and could not be explained by ecto-LPP activity. The effects of

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. Table 12-2 Possible signaling lipids that could be regulated by LPP activity Phosphatidate (PA) (1) Stimulates NADPH oxidase and H2O2 production in neutrophils (2) Binds to and stimulates protein kinase C-z (3) Stimulates phosphatidylinositol-4-phosphate kinase (4) Stimulates phospholipase C-g (5) Increases Ras-GTP, Raf and ERK activation (6) Increases cell division through activation of mTOR (7) Increases sphingosine kinase-1 activity (8) Increases stress fiber formation (9) Binds to and inhibits protein phosphatase-1 (10) Relative LPA and PA in membranes control their curvature, vesicle budding and transport

References Regier et al. (2000) Limatola et al. (1994) Moritz et al. (1992) Jones and Carpenter (1993) Ghosh et al. (1996), Rizzo et al. (1999) Fang et al. (2001), Kam and Exton (2004), Avila-Flores et al. (2005) Delon et al. (2004) Ha and Exton (1993), Cross et al. (1996) Jones and Hannun (2002) Ktistakis et al. (1995), Ktistakis et al. (1996), Abousalham et al. (1997), Jones et al. (1999), Schmidt et al. (1999), Weigert et al. (1999), Huijbregts et al. (2000), Zimmerberg (2000), Abousalham et al. (2002)

Diacylglycerol (DAG) (1) Stimulation of classical and novel protein kinase Cs (2) Activation of RasGRP (guanine nucleotide releasing proteins) Lysophosphatidate (1) Activates PPAR-g receptors (2) Activates nuclear receptor, LPA1 Sphingosine 1-phosphate (1) Mobilizes intracellular Ca2+ independently of IP3 (2) Increases actin stress fiber formation (3) Increases ERK activity and cell division (4) Protects against apoptosis (5) Increases COX-2 activity and eicosanoid synthesis Ceramide 1-phosphate (C1P) (1) Involved in synaptic vesicle movement and neutrophil phagocytosis (2) External C1P stimulates fibroblast division (3) Binds and activates cytosolic phospholipase A2, thereby increasing arachidonate and prostaglandin E2 production (4) Blocks the activation of apoptosis in macrophages by inhibiting acidic sphingomyelinase activity

References Newton (1997) Stone (2006) References McIntyre et al. (2003), Zhang et al. (2004) Gobeil et al. (2003), Moughal et al. (2004)

References Pyne and Pyne (2000) Pyne and Pyne (2000), Spiegel and Milstien (2003) Pyne and Pyne (2000), Hanna et al. (2001), Spiegel and Milstien (2003) Pyne and Pyne (2000), Spiegel and Milstien (2003) Pettus et al. (2003b), Pettus et al. (2005) References Bajjalieh et al. (1989), Kolesnick and Hemer (1990), Shingal et al. (1993), Hinkovska-Galcheva et al. (1998) Go´mez-Mun˜oz et al. (1995) Pettus et al. (2003a, 2004)

Go´mez-Mun˜oz et al. (2004)

Increased lipin activity should increased DAG production through phospholipase D and decrease PA accumulation. The lower PA concentrations should decrease LPA formation, the activation of sphingosine kinase and thus the production of sphingosine 1-phosphate. These actions could regulate signaling cascades that are stimulated by the bioactive lipids as shown above

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. Figure 12-3 Metabolic interrelations among intracellular bioactive lipids formed following activation of phospholipase D1, or D2. Phosphatidate also activates sphingosine 1-phosphate (S1P) formation

LPP2 and LPP3 on intracellular PA and S1P concentrations, respectively, control cell survival (Long et al., 2005). Increased LPP1 activity also attenuates Ca2+-transients and production of inflammatory cytokine, IL-8, downstream of LPA receptor activation (Zhao et al., 2005). HEK 293 cells that over-express LPP3 exhibited greater DAG formation following stimulation of phospholipase D and consequent PA formation (see > Figure 12-3). PLD2 and LPP3 are both present in caveolin-1-enriched microdomains (Sciorra and Morris, 1999). It was postulated that chronic increases in DAG concentrations following overexpression of LPP1 decreased expression of some PKCs and thereby ERK activation. This could decrease cell division (Long et al., 2005) and PDGF-induced cell migration (Long et al., 2006). Overexpression of LPP1 also blocks PLD activation and PA formation by LPA and PDGF (Pilquil et al., 2001a, b). This indicates that LPP1 alters cell signaling upstream of PLD activation in addition to converting PA to DAG (> Figure 12-3). The stimulation of cell migration by LPA requires activation of PLD2 and increased expression of LPP1 attenuates this stimulation and consequent migration (Pilquil et al., 2006). Individual LPPs regulate cell activation in different ways. A major effect of LPP2 is to control entry into S-phase of the cell cycle (Morris et al., 2006). Increasing LPP2 activity increases entry into S-phase, whereas decreasing the activity has opposite effect. All of the LPPs are expressed in the brain (Sigal et al., 2005). Although, the LPPs are normally thought to metabolize PA formed after PLD action (Brindley, 2004), our recent work showed that a major effect of LPP1 in decreasing PA accumulation is upstream and not downstream of PLD activation (Pilquil et al., 2006). The active sites of LPPs are outside the cell, or on the luminal surface of ER, or Golgi membranes (Brindley, 2004). By contrast, PLDs should produce PA on the cytosolic surface of membranes. Thus there is a barrier to the hydrolysis of PA formed on the cytosol surface of membranes unless the PA is efficiently transported to the active sites of LPPs. A second possible route for the metabolism of PA is through the actions of the type-1 phosphatidate phosphatases (PAP1) as discussed in Sect. 12.5.

5.4 Lipid Phosphate Phosphatase Expression in the Central Nervous System mRNA for LPP1 is highly expressed in human aorta, bladder, uterus, kidney, mammary gland, spinal cord, small intestine, adrenal gland and thyroid gland. However, mRNA expression in the brain is relatively low (Kai et al., 1997). LPP2 RNA expression is more restrictive with a 1.4 kb size detected highly in pancreas and

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a smaller amount in brain. There was also strong expression in placenta, but the RNA was somewhat smaller (1.3 kb) (Hooks et al., 1998). mRNAs for LPP3 is expressed more ubiquitously in human tissues with particularly high expression in heart and skeleltal muscle (Kai et al., 1997). Suzuki et al. showed that LPP3 mRNAs are expressed in developing and mature rat brains (Suzuki et al., 1999). Dominant expression was found in the ventricular germinal zone without significant expression in the intermediate, mantle, and marginal zones throughout the embryonic brain and spinal cord. The dominant expression in the ventricular germinal zone was maintained at P0 and P7, but it was markedly decreased at later postnatal stages. Persistently high expression was detected in Bergmann glial cells of the cerebellar Purkinje cell layer throughout postnatal development.

5.5 Lipid Phosphate Phosphatases and Development Evidence that LPPs play a decisive role in cell migration processes in vivo has been provided by developmental studies on the Drosophila embryo. Drosophila express two Wunen proteins homologous to LPP3. Wunen-1 and -2 negatively regulate the migration of primordial germ cells (Zhang et al., 1997; StarzGaiano et al., 2001; Burnett and Howard, 2003). Introduction of mouse LPP1 has no effect on an endogenous Drosophila germ-cell-specific factor in vivo, whereas human LPP3 causes aberrant migration and germ cell death (Burnett and Howard, 2003). It was proposed that Drosophila germ-cell migration and survival are controlled by competition for hydrolysis of a lipid phosphate between germ cells and soma (Renault et al., 2004). These results demonstrate that individual LPPs have distinct functions and that they participate in a signaling system for cell migration that may be conserved between flies and human beings. This conclusion is supported by work with mouse embryos from LPP3 knockout mice, which failed to form a chorio-allantoic placenta and yolk sac vasculature. Some embryos showed a shortening of the anteriorposterior axis (Escalante-Alcalde et al., 2003) similar to that in axin deficiency, a critical regulator of Wnt signaling. It was proposed (Escalante-Alcalde et al., 2003) that LPP3 could normally function as a Wnt signaling antagonists.

6

Importance of Phosphatidate Phosphatases-1 (Lipin) Activities

PAP1 enzymes were originally differentiated from the LPPs since they require Mg2+ for activity and they are inhibited by N-ethylmaleimide (Jamal et al., 1991). The LPPs are integral membrane proteins, whereas PAP1 activity is translocated onto membranes in response to an accumulation of fatty acids, acyl-CoA esters and phosphatidate in these membranes (Brindley, 1988). This translocation causes PAP1 to become metabolically active at the site of PA synthesis. The translocation represents a feed-forward signal for increasing DAG production for the synthesis of triacylglycerols, phosphatidylcholine and phosphatidylethanolamine (Brindley, 2004). Consequently, a major function for PAP1 in the brain is to facilitate phospholipid synthesis for membrane turnover and myelination. However, PAP1 is ideally located in the cytosol to translocate and metabolize PA that is formed on the cytosolic surface of membranes after PLD activation (> Figure 12-3). In support of this function, PAP1 activities have been shown to be involved in the regulation of cell signaling. PAP1 co-immunoprecipitates with EGF receptors, which on activation appear to transfer the PAP1 activity to protein kinase-C-e (PKC-e), a DAG-dependent PKC (Jiang et al., 1996). PAP1 activity is also involved in cyclo-oxygenase expression and eicosanoid formation when WISH cells are activated through protein kinase C (Johnson et al., 1999) and when macrophages are stimulated with lipopolysaccharide (Grkovich et al., 2006). The activity of PAP1 helps to preserve cell integrity and it protects against apoptosis (Fuentes et al., 2003). These latter events are associated with PLD activation. Attempts to purify and identify the structure of PAP1 failed and so it was very difficult to explore the role of its activity in regulating glycerolipid synthesis and cell activation. This situation changed with the finding in February 2006 that yeast PAP1 (Smp2, Pah1) is the orthologue of mammalian lipins (Han et al., 2006). Phosphorylation of Pah1p regulates PA production in yeast. This controls the activity

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of Opi1p, a transcriptional repressor of phospholipid biosynthetic genes (O’Hara et al., 2006). Genetic analysis suggests that Pah1p regulates transcription of these genes through both Opi1p-dependent and independent mechanisms (Carman and Han, 2006). The transcriptional activity depends upon the catalytic activity of Pah1 (Han et al., 2007). It was already known that the mammalian lipins regulate triacylglycerol synthesis, adipose tissue development and gene transcription. The fatty liver dystrophy (fld) mouse has a null mutation in the lipin (Lpin1) gene (Peterfy et al., 2001). These mice are characterized by absence of adipose tissue, insulin resistance, and susceptibility to atherosclerosis (Reue et al., 2000; Reue and Donkor, 2006). White and brown adipose tissue and skeletal muscle from fld mice have no PAP1 indicating that lipin-1 is responsible for all the PAP1 activities (Donkor et al., 2007). The expression of two lipin-1 slice variants in adipocytes is essential for not only triaclylgycerol synthesis by also PPARg expression and adipocyte differentiation (Phan et al., 2004, 2005). PAP1 activity in liver is not significantly different in fld mice compared to controls (Donkor et al., 2007; Harris et al., 2007) because the liver also expresses lipin-2 and -3. All of the mammalian lipins display PAP1 activity, but to different extents (Donkor et al., 2007). The mammalian lipins specifically degrade PA and they are inactive against LPA, C1P, and S1P (Donkor et al., 2007). In liver, PAP1 activity is involved in promoting triacylglycerol and phospholipid synthesis in starvation and diabetes (Brindley, 1988). In this respect, lipin-1 expression is promoted by PPARg coactivator 1a (PGC-1a) and lipin-1 activates a subset of PGC-1a target pathways including b-oxidation and it inhibits FA synthesis (Finck et al., 2006). There is a physical interaction of lipin-1 with PPARa and PGC-1a which amplifies the PGC-1a/PPARg-mediated control of hepatic metabolism in starvation. This interaction does not depend upon the catalytic activity of lipin-1. In adipose tissue, lipin-1 expression controls gene transcription through PPARg and this explains why fld mice, which do not express lipin-1, fail to develop mature adipocytes. These results again illustrate the role of PAP1 (lipins) in controlling both glycerolipid synthesis and gene transcription. The functions of lipins in the brain have not been determined, but the brains of mice and human beings express relatively high levels of lipin-2 as determined by RT-PCR. It is, therefore, likely that lipin-2 in the brain could regulate phospholipid synthesis, signaling by PA and also gene transcription.

7

Neurobiological Effects of LPA

7.1 Tau Hyperphosphorylation by LPA Growth cone collapse and neurite retraction are crucial processes during nervous system development and neurodegeneration and are less understood than the regulation of neurite extension. The bioactive lipid, LPA, is one of the molecules which induces neurite retraction. This happens through the activation of specific G-protein (Gaq, Gai, Ga13, Ga12)-coupled receptors (LPA1–5) and the activity modulation of the small GTP-binding protein, Rho. This GTPase regulates many cellular processes, such as actomyosinbased contraction, actin and tubulin cytoskeleton organization, cell shape, and cell division. Several years ago, the group of Wandosell described an increase in site-specific Alzheimer’s disease-like Tau phosphorylation during LPA-induced neurite retraction. The Tau phosphorylation was mediated by glycogen synthase kinase (GSK-3) (Sayas et al., 1999, 2002a, b). Hyperphosphorylated Tau is the main constituent of paired helical filaments (PHFs) in neurofibrillary tangles. The same group also showed that the activation of GSK-3 by LPA is not downstream of the Gai and Gaq pathways, but downstream of Ga12 or Ga13. Interestingly, GSK-3 activation by Ga13 is RhoA-mediated, whereas its activation by Ga12 is Rho-independent.

7.2 LPA in Myelination Process A role for LPA in myelination and/or neurodegeneration is suggested by the finding that LPA promotes survival and cell–cell adhesion of myelinated Schwann cells of the peripheral nervous system (Weiner and Chun, 1999; Weiner et al., 2001).

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7.3 LPA During Brain Development During brain development, LPA regulates cortical neurogenesis and is involved in the process of migration and cortical layer formation in vertrebrates (Hecht et al., 1996; Fukushima et al., 2000; Kingsbury et al., 2003; Fukushima, 2004). In these studies, Chun’s group proved with brain slice cultures that postmitotic neurons in the CNS act as an LPA-generating source and that LPA can thereby regulate neuronal migration and differentiation in the neocortex. In fact, LPA can cause a gyrification of the lissencephalon rodent cortex (Kingsbury et al., 2003).

7.4 Effects of LPA on Brain Tumors Tumors of the CNS arise from several different cellular lineages that include glia, such as astrocytes and oligodendrocytes. The World Health Organisation defines three grades of astrocytic tumours: astrocytoma (WHO grade II), anaplastic astrocytoma (WHO grade III) and glioblastoma multiforme (GBM, WHO grade IV). Constitiuting 60% of all glial tumours, GBM is the most common and malignant. Median survival in patients with GBM is about 15 months. GBM cells are highly motile and invade the normal brain parenchyma diffusely (Burger et al., 1983), secreting several factors that result in autocrine motility signaling. Immunohistochemistry studies demonstrate that ATX is found in all astrocytic tumors but not in normal astrocytes (Hoelzinger et al., 2005; Kishi et al., 2006) and it stimulates tumor motility. The LPA1 receptor, to which LPA produced by ATX binds, is predominantly expressed in GBM cells and tissues. The blood-brain-barrier is disrupted in GBM tissue, which leads to direct exposure of tumor cells to plasma components. In humans, LPC concentration is extremely high in plasma (100–300 mM) (Kishimoto et al., 2002), but low in the brain, such as in cerebrospinal fluids (several mM) (Mulder et al., 2003). This suggests that if BBB is disrupted in GBM the tumor is exposed to LPC derived from plasma. LPC is then converted to LPA by ATX expressed by glioblastomas, and consequently, LPA1 is activated, leading to the enhanced cell motility and high invasiveness of GBM.

7.5 Effects of LPA on Epileptic Seizure The nervous system is thought to respond to recurring epileptic seizures by becoming increasing hyperexcitable, which alters dendritic growth. The primary evidence of this is the frequently observed decrease in dendritic branches and spines in epilepsy (Multani et al., 1994; Nishimura et al., 2007). Several studies have shown alterations in lipid metabolism or lipid signaling after seizure (Bazan et al., 1986, 2002; Katsura et al., 2000; Kulagina et al., 2004). Bazan et al. (1986) and Dahlin et al. (2007) demonstrated that seizures increase diacylglycerol (DAG) and un-esterified fatty acid levels in the rat neocortex. Katsura et al. (2000) later showed that, together with a transient lipid increase, there is a transient release/reuptake of Ca2+ and a transient change in levels of cAMP/cGMP. Epileptic seizure induced in rats also increases secretory phospholipase A2 activity, especially in the hippocampus and cortex (Yegin et al., 2002). Additionally, significant decreases have been observed in phosphatidylethanolamine content together with increases in LPC in hippocampal synaptosomes. PLA2 represents a growing superfamily of enzymes that catalyze the hydrolysis of glycerophospholipids at sn-2 position (Bonventre, 1999). Therefore, PLA2 can modulate the synthesis of bioactive lipids, which is regulated through several cell mechanisms in signaling cascades. Strong evidence exists that phospholipids play numerous roles in epilepsy, which was also supported by clinical studies in which plasma phospholipid fatty acid levels changed in children with epilepsy after ketogenic diet enriched with n-3 fatty acids (Dahlin et al., 2007). Vining found that 50–67% of epileptic children treated with such a diet achieve a >50% seizure reduction. Of 17 fatty acids analyzed, linoleic acid and eicosapentaenoic acid were increased, whereas arachidonic acid and 5,8,11 eicosatrienoic acid were decreased (Vining, 1999). Despite these results, no correlation between changes in fatty acid levels and seizure controls has been found to date and the exact nature of the link remains unclear.

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Epilepsy with mental retardation (EPMR) belongs to a group of inherited neurodegenerative disorders, the neuronal ceroid lipofuscinoses. EPMR is characterized by normal early development, onset of seizures by age 5–10 and progressive mental deterioration (Herva et al., 2000). EMPR has two main stages of disease – progressive and advanced. The CLN8 gene, which underlies EPMR, encodes a novel transmembrane protein thought to play a role in lipid synthesis, transport or sensing. In the progressive stage, EPMR patients exhibit reduced levels of ceramide (galactosyl- and lactosylceramide) and sulfatide. However, these patients also show an increase in molecular species containing polyunsaturated acyl-chains, particularly phosphatidylserine and phosphatidylethanolamine. In the advanced stage, saturated, and monounsaturated species were overrepresented among phosphatidylserine, phosphatidylethanolamine and phosphatidylinositol (Hermansson et al., 2005). It is speculated that these changes in brain sphingo- and glycerophospholipid molecular profiles may result in altered membrane stability, lipid peroxidation, vesicular trafficking, or neurotransmission.

8

Plasticity-Related-Genes (Lipid Phosphate Phosphatase-Related Proteins)

Plasticity-related genes (PRGs/LPRs) are a defined novel subclass of the LPP superfamily, comprising five brain- and vertebrate-specific membrane proteins, which interfere with lipid phosphate signaling and thereby promote neurite growth (Brauer et al., 2003; Savaskan et al., 2004). The first gene, named PRG-1, was identified by screening of a cDNA library of lesioned hippocampus as part of a study searching for genes involved in axon guidance processes. Four other genes were found by in silico analyses to have high homology to PRG-1. Sequence and hydropathy analysis reveals that PRGs show homology to LPP1. The PRGs, like the LPPs, contain six putative transmembrane regions and the C- and the N-terminus are located intracellularly (Brindley, 2004). However, PRGs differ in their catalytic motif sequences in that a few of their conserved amino acids differ to those of LPPs (Brindley, 2004; Sigal et al., 2005). In the case of PRG-1 and PRG-2 alone, C-terminus is particularly long, consisting of around 400 amino acids, but is, like all the PRGs and LPPs, still predicted to be located intracellularly (Brindley, 2004).

8.1 PRG Expression in the Adult Central Nervous System and During Development PRG-1 mRNAs exhibit dominant expression in late mouse embryonic stages, specifically in the hippocampus region (E18). After birth, mRNA expression increases dramatically in several neuronal brain areas, such as the neocortex, hippocampus ,and cerebellum. The transcript increase is detectable until postnatal day p20 and then it decreases to a stable level in adulthood (Brauer et al., 2003). PRG-3 mRNAs are expressed during mouse embryonic development as early as E14 in the dorsal cortex, cingulated cortex and anlage of the hippocampus. At E16, PRG-3 mRNAs are strongly detectable in the cortical plate of the cortex and moderately expressed in the marginal zone and intermediate zone. This pattern is unchanged until birth. After birth, PRG-3 is expressed in all cortical layers. In the adult, brain tissue expresses PRG-3 mRNA homogenously (Savaskan et al., 2004; Wang and Molnar, 2005).

8.2 PRG Response to LPA Functional studies of PRG-1 demonstrated that its overexpression attenuates LPA-induced neurite collapse in neuronal cells (Brauer et al., 2003). Point mutation in PRG-1 conserved catalytic domain 2, his252 to lysine, led to an abolishment of this protective effect. Further tests showed an increase in MAG in PRG-1 overexpressing cells, whereas the point mutated PRG-1 version showed no alteration in LPA-MAG levels (Brauer et al., 2003). Nevertheless, McDermott et al. could not find ecto-LPA phosphatase activity in cells overexpressing PRG-1 (McDermott et al., 2004). On the basis of incomplete conservation of the catalytic

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motif in PRG-1 compared to LPP1, they suggested that PRG-1 must have alternate biological activity that is responsible for its effect on the attenuation of LPA-induced neurite retraction. Resolving this discrepancy between findings and thereby clarifying the function of protein and phospholipid metabolism in the brain requires further investigation. In contrast, PRG-3 does not attenuate LPA-induced neurite collapse (Savaskan et al., 2004). Interestingly, PRG-3 overexpression in neuronal cells leads to an increase in neurite length, as well as neurite numbers, and a ‘‘spreading-like’’ cell body (Savaskan et al., 2004).

8.3 PRG Expression After Brain Damage Entorhinal lesion in the adult brain leads to anterograde degeneration of perforant path fibers in their main hippocampal termination zones. Subsequently, remaining fibers sprout and form new synapses on the denervated dendrites. PRG-1 expression shows upregulation, particularly in the deafferented zone. The expression occurs on outgrowing axons and their growth cones, replacing those lost (Brauer et al., 2003). These results suggest that PRG-1 could be involved in processes of regeneration in the adult CNS. For example, increased ATX overexpression around brain lesions (Savaskan et al., 2007) converts plasmaderived LPC to LPA. To control axonal outgrowth, only PRG-1 overexpressing axons are able to sprout into the deafferented zone to form new synapses because of the effects of PRG-1 in modulating LPA activity. The kainic acid model of chemically-induced generalized seizures is an attractive model for evaluating activity-dependent neuronal gene expression changes. PRG-3 mRNA expression is downregulated in the hippocampus shortly after kainic acid-induced seizure (Savaskan et al., 2004). Kainic acid exerts its principal action on limbic structures, producing a state of convulsive seizures. Within the hippocampus, kainic acid-induced seizures initiate a process of events that ends in the death of certain neurons, particularly those that reside in the CA3 and CA1 subfields. However, the PRG-3 level recovers five days after seizure, which contradicts the simple explanation that PRG-3 downregulation correlates with cell death.

9

Conclusions

The CNS is highly organized by a broad variety of brain cells. The number of cells generated in the developing nervous system is probably regulated at several levels. In comparison with other organs in mammalians, CNS contains the largest diversity of lipid classes and lipid molecular species. Increasing evidence indicates that bioactive lipids as well as enzymes and proteins, which regulate lipid metabolism participate in the regulation of brain development, brain function and dysfunction. Information on the complexity, composition, and kinetic turnover of bioactive lipids can reveal substantial insights into neuronal plasticity and gene function. Expanding our understanding of lipid regulation in neuronal diseases should identify pathological alterations that are diagnostic of disease onset, progression, or severity. These new insights into the regulation of bioactive lipids could resolve our understanding of how the complex interactions of the nervous system are regulated.

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Witte ON, Kabarowski JH, Xu Y, Le LQ, Zhu K. 2005. Retraction. Science 307: 206. Wu J, Cunnick JM. 2002. Trans-regulation of epidermal growth factor receptor by lysophosphatidic acid and G protein-coupled receptors. Biochim Biophys Acta 1582: 100-106. Yart A, Chap H, Raynal P. 2002. Phosphoinositide 3-kinases in lysophosphatidic acid signaling: Regulation and cross-talk with the Ras/mitogen-activated protein kinase pathway. Biochim Biophys Acta 1582: 107-111. Ye X, Hama K, Contos JJ, Anliker B, Inoue A, et al. 2005. LPA3-mediated lysophosphatidic acid signalling in embryo implantation and spacing. Nature 435: 104-108. Yegin A, Akbas SH, Ozben T, Korgun DK. 2002. Secretory phospholipase A2 and phospholipids in neural membranes in an experimental epilepsy model. Acta Neurol Scand 106: 258-262. Zhang C, Baker DL, Yasuda S, Makarova N, Balazs L, et al. 2004. Lysophosphatidic acid induces neointima formation through PPARgamma activation. J Exp Med 199: 763-774. Zhang QX, Pilquil CS, Dewald J, Berthiaume LG, Brindley DN. 2000. Identification of structurally important domains of lipid phosphate phosphatase-1: Implications for its sites of action. Biochem J 345(Pt 2): 181-184. Zhang N, Zhang J, Purcell KJ, Cheng Y, Howard K. 1997. The Drosophila protein Wunen repels migrating germ cells. Nature 385: 64-67. Zhao Y, Kalari SK, Usatyuk PV, Gorshkova I, He D, et al. 2007. Intracellular generation of sphingosine 1-phosphate in human lung endothelial cells: Role of lipid phosphate phosphatase-1 and sphingosine kinase 1. J Biol Chem 282: 14165-14177. Zhao Y, Usatyuk PV, Cummings R, Saatian B, He D, et al. 2005. Lipid phosphate phosphatase-1 regulates lysophosphatidic acid-induced calcium release, NF-kappaB activation and interleukin-8 secretion in human bronchial epithelial cells. Biochem J 385: 493-502. Zhu K, Baudhuin LM, Hong G, Williams FS, Cristina KL, et al. 2001. Sphingosylphosphorylcholine and lysophosphatidylcholine are ligands for the G protein-coupled receptor GPR4. J Biol Chem 276: 41325-41335. Zimmerberg J. 2000. Are the curves in all the right places? Traffic 1: 366-368.

13

Metabolism and Functions of Platelet-Activating Factor (PAF) in the Nervous Tissue

G. Goracci . M. L. Balestrieri . V. Nardicchi

1

Historical Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 313

2

Methods for PAF Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 315

3

Levels of PAF in Neural Cells and Mammalian Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 316

4 4.1 4.2 4.3 4.4

Metabolism of PAF: General Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 317 Biosynthesis of PAF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 317 De Novo Synthesis of PAF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 318 Remodeling Pathway of PAF Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 319 PAF Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 321

5 5.1 5.2 5.3 5.4

PAF Metabolism in Nervous Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 321 De Novo Synthesis of PAF in Nervous Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 321 PAF Biosynthesis by the Remodeling Pathway in Nervous Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 322 Degradation of PAF in the Nervous Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 322 Regulation of PAF Metabolism in Neural Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 323

6

PAF Receptors and Cell Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 325

7

PAF Receptors Agonists and Antagonists . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 326

8 8.1 8.2 8.3 8.4 8.5

Functions of PAF in the Nervous Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 327 Synaptic Transmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 327 LTP and Memory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 327 Proliferation and Differentiation of Neural Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 329 Gene Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 330 Chemotaxis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331

9 9.1 9.2 9.3 9.4 9.5 9.6 9.7 9.8

Role of PAF in Brain Dysfunctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331 Neuroinflammation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331 Glutamate Neurotoxicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 332 Seizures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 334 Brain Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 334 Oxidative Stress and Aging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 335 Hypoxia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 336 Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 336 Parkinson’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 336

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_13, # Springer ScienceþBusiness Media, LLC 2009

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9.9 9.10 9.11 9.12

Miller–Dieker Lissencephaly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Multiple Sclerosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . HIV Infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Prion Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

337 337 337 338

10

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 338

Metabolism and functions of platelet-activating factor (PAF) in the nervous tissue

13

Abstract: Platelet Activating Factor (PAF, 1-alkyl-2-acetyl-sn-glycero-3-phosphocholine) was first identified as a lipid mediator of inflammation and immunological response. This compound is present at very low concentration in normal mammalian brain where it is synthesized by two distinct pathways. The de novo pathway utilizes 1-alkyl-2-acetyl-sn-glycerol and CDP-choline as substrates of a DTT-insensitive phosphocholine transferase (PAF-PCT). The remodeling pathway requires the production of 1-alkyl-2lyso-sn-glycero-3-phosphocholine (lysoPAF) produced by the hydrolysis of 1-alkyl-2-(long-chain)acylsn-glycero-3-phosphocholine (alkylacylGPC) by the action of phospholipases A2. Alternatively, lysoPAF can be produced by transacylation from alkylacyl-GPC to 1-alk-1’-enyl-2-lyso-sn-glycero-phosphoethanolamine (lysoPlsEtn) produced by a phospholipase A2 as well. LysoPAF is acetylated to PAF by lysoPAF acetyltransferase (lysoPAF-AcT). The relative contribution of the two pathways to PAF synthesis depends on several factors including the concentration of substrates, energy availability, Ca2+ concentration, and phosphorylation state of key enzymes. PAF is transformed into the inactive lysoPAF by PAF acetylhydrolases (PAF-AH). Three intracellular isoforms of PAF-AH have been identified in mammalian brain. In neural cells, PAF is not stored and can be released into the extracellular space. Neural cells possess plasma membrane and intracellular receptors. Plasma membrane receptor (PAFR) has been pharmacologically characterized and cloned. This receptor is expressed in almost all brain areas. Cellular responses to the activation of PAFR are mediated by G-proteins and lead to a complex intracellular signaling. A number of natural and synthetic antagonists of PAFR and intracellular binding sites have been identified. In the brain, PAF participates im physiological mechanisms such as synaptic transmission, long-term potentiation, memory formation, proliferation and differentiation of neural cells, regulation of gene expression, and chemotaxis. Increased levels of PAF, due to upregulation of biosynthetic pathways or to downregulation of PAF-AH, have been observed in pathological conditions such as neuroinflammation, brain ischemia, and neurodegenerative diseases. Relatively high concentration of PAF causes neuronal death by apoptosis, which is linked to the neurotoxic effects of the execessive glutamate release, causing overloading of post-synaptic Ca2+ and the consequent activation of Ca2+-dependent enzymes including PLA2s. List of Abbreviations: acylPAF, 1-acyl-2-(short-chain) acyl-GPC; acyl-DHAP, 1-acyl-DHAP; alkenylacylGPE, 1-alk-1’enyl-2-acyl-GPE; alkylacetylG, 1-alkyl-2-acetyl-sn-glycerol; alkylacylGPC, 1-alkyl-2-(long-chain) acyl-GPC; alkylGP, 1-alkyl-2-lyso-sn-glycero-3-phosphate; AlkylGP-AcT, alkylGP acetyltransferase; DHAP, dihydroxyacetone phosphate; GPC, glycerophosphocholine; GPE, glycerophosphoethanolamine; lysoPAF, 1-alkyl-2-lyso-GPC; lysoPAF-AcT, acetyl-CoA:lyso-PAF acetyltransferase; lysoPC, 1-acyl-2-lysoGPC; mc-PAF, 1-O-alkyl-2-N-methylcarbamyl-GPC; PAF, platelet activating factor; PAF-AH, PAF acetyl hydrolase; PAF-PCT, DTT-insensitive phosphocholinetransferase; PAFR, PAF plasma membrane receptor; PFPC, 1-hexadecanoyl-2-formyl-glycerophosphocholine; PlsEtn, ethanolamine plasmalogens; PlsEtnPLA2, plasmalogen-selective PLA2

1

Historical Background

The term ‘‘platelet-activating factor’’ (PAF) was coined by Benveniste et al. when they reported that a factor able to induce platelet aggregation was released by IgE-stimulated basophils (Benveniste et al., 1972). Different studies then indicated that this ‘‘factor’’ possesses lipophilic properties (Benveniste et al., 1977; Pinckard et al., 1979). Finally, its structure became evident when it was demonstrated that the chemical and biological properties of the semisynthetic compound 1-alkyl-2-acetyl-sn-glycero-3-phosphocholine (> Figure 13‐1) and PAF were identical (Benveniste et al., 1979; Blank et al., 1979; Demopoulos et al., 1979). A few months later, it was clearly demonstrated that the native PAF isolated from rabbit basophils stimulated with Ig-E had the same structure (Hanahan et al., 1980). The discovery of the chemical structure of PAF led to classify this molecule as a ‘‘lipid mediator,’’ a term previously used mainly for eicosanoids. Immediately after the identification of the chemical structure of PAF, there was an enormous intensification of the efforts for developing procedures for its detection in biological samples and for understanding its role in physiological and pathological conditions. The biochemical pathways for PAF synthesis and degradation were also demonstrated thanks to the basic knowledge of the

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. Figure 13‐1 1-hexadecyl-2-acetyl-sn-glycero-3-phosphorylcholine (C16-PAF)

biochemistry of ether lipids developed by a few groups interested in this peculiar area and, particularly, to the important contributions from Fred Snyder’s laboratory in Oak Ridge (Snyder, 1995b). The presence of a long-chain alkyl group at C-1 in PAF molecule suggested a metabolic relationship with 1-alkyl-2-(long-chain)acyl-sn-glycero-3-phosphocholine (alkylacylGPC), which was known to be present in many cell membranes at relatively low concentrations, with respect to 1,2-diacylGPC, but its functional implications had received little attention until the discovery of PAF structural properties. Alkylacyl-GPC belongs to a class of glycerophospholipids commonly indicated as ‘‘ether lipids,’’ which also includes ‘‘plasmalogens’’ (1-alk-1’enyl-2-acyl-sn-glycero-3-phosphoethanolamine or choline; alkenylacyl-GPE or -GPC). The occurrence of ether-linked phospholipids in mammalian tissues has been extensively reviewed (Sugiura and Waku, 1987). In most tissues, the predominant alkyl chain at C-1 of alkylacyl-GPC is 16:0, but also 18:0 and 18:1 can be found although to a lower extent. In the same phospholipid class, fatty acids at C-2 are highly unsaturated with a relatively large content of arachidonic acid (20:4 o-6) (Nakagawa and Horrocks, 1986). The crucial step for the synthesis of ether lipids is the formation of the ether bond. Pioneering studies demonstrated its formation from acyl-dihydroxyacetonephosphate (acyl-DHAP) and long-chain fatty alcohols by the action of alkyldihydroxyacetone synthase producing alkyl-DHAP, which was then reduced to 1-alkyl-2-lyso-sn-glycero-3-phosphate (alkylGP) by NADPH:alkyl-DHAP oxidoreductase (Hajra and Bishop, 1982; Snyder et al., 1985). Alternatively, 1-alkylglycerols could be be directly phosphorylated to alkylGP by a microsomal phosphotranspherase (Chae et al., 1973). Once alkylGP is formed, it can be linked to the formation of several ether lipids and some of them are the direct precursors of PAF.

Metabolism and functions of platelet-activating factor (PAF) in the nervous tissue

2

13

Methods for PAF Assay

PAF is a potent lipid mediator because it induces cellular responses at picomolar concentrations (Demopoulos et al., 1979). The determination of PAF levels represents a significant technical challenge due to the presence of catabolic enzymes (PAF-acetylhydrolases) in cells and plasma. Before the advent of radioimmunoassay (RIA) techniques, quantitation of PAF relied on bioassays based on the interaction of this lipid mediator with washed rabbit platelets. The resultant aggregation or the release of radiolabeled serotonin from these cells was determined with high sensitivity (Hanahan and Weintraub, 1985, Bossant et al., 1989) due to the potency of PAF in inducing biological effects (Bossant et al., 1989, 1990; Henson, 1990; Marathe et al., 1999). Samples assayed for PAF content by platelet aggregation are suspended in 0.1% bovine serum albumin in saline solution and added to platelet suspension. The subsequent change in light transmission is monitored by an aggregometer (Bossant et al., 1989). Quantitation of PAF by release of endogenous platelet serotonin is based on the measurement of serotonin released from washed rabbit platelets by electrochemical detection after its isolation by highperformance liquid chromatography (HPLC) (Bossant et al., 1989). Alternatively, platelets are preloaded with radiolabeled serotonin and the PAF levels in the sample are determined on the basis of the radioactivity of serotonin present in the supernatant after stimulation (Henson, 1990). RIA techniques overcome the limitations of PAF bioassays, which are not sufficiently accurate because the conditions used for platelet preparation affect reproducibility. RIA assay is suitable for PAF quantitation in the range 0.1–30 pmol (Karasawa et al., 1991). Cross-reaction with lysoPAF, lysophosphatidylcholine, and long-chain phosphatidylcholines was less than 0.025%, whereas 1-palmitoyl-2-acetyl-sn-glycero-3phosphocholine cross-reacted slightly (6.25%). Commercially available RIA for PAF assay utilizes the technique of scintillation proximity assay, which eliminates the need to separate antibody bound from free ligands, which is common to heterogeneous methods (GE Healthcare). This assay allows the detection of PAF in vitro within a range of 20–1,280 pg/tube. Radiolabeling of PAF with radioactive precursors for detecting its production in cellular systems offers relatively straightforward and sensitive tools for detecting PAF formation, but they do not allow measurement of its concentration in cells and tissues (Wykle et al., 1988). Indeed, when radiolabeled acetate is employed, 1-acyl-2-acetyl-GPC (acylPAF) and 1-O-alk-1’-enyl-2-acetyl-sn-glycero-3-phosphoethanolamine are also often labeled (Wykle et al., 1988). Chromatographic isolation of PAF is often required before bioassay to avoid interference by compounds having PAF-like biological activity. The most sensitive, accurate, and specific PAF assays are those based on mass spectrometry (MS) (Wykle et al., 1988). The removal of the phosphocholine head group and its replacement with a nonpolar group result in the formation of a volatile derivative suitable to be analyzed by GC-MS and allows detection of as little as 50 fg of PAF (Satouchi et al., 1983; Ramesha and Pickett, 1986; Yamada et al., 1988; Satsangi et al., 1989; Weintraub et al., 1990; Balazy et al., 1991; Haroldsen et al., 1991). This procedure is very sensitive although has the disadvantages of being time consuming, expensive, and susceptible to losses and artifacts. PAF assay based on HPLC coupled to MS or tandem MS (LC-MS or LC-MS/MS) overcomes these problems (Kim et al., 1987; Savu et al., 1996; Harrison et al., 1999). Harrison et al. (1999) developed a PAF assay using negative-ion and normal-phase liquid chromatography/ tandem mass spectrometry (LC/MS/MS) as an alternative technique to the GC/MS and positive-ion LC/MS/MS procedures. Based on this method, the positive ion [M + H]+ derived from PAF and generated by electrospray ionization is abundant, but the potential presence of isobaric 1-octadecanoyl-2-lyso-glycerophosphocholine and 1-hexadecanoyl-2-formyl-glycerophosphocholine (PFPC) in biological samples limits the use of the most abundant collision-induced decomposition (CID) transition (formation of the phosphocholine ion, m/z 524←184). With negative-ion LC/MS/MS, however, stearoyl-lysoGPC and PFPC decompose to the carboxylate anions at m/z 59, 283, and 255, respectively, permitting discrimination of these isobaric molecules even without chromatographic separation. This approach allows detection of 500 pg of PAF with a good signal-to-noise ratio. Another report has achieved better sensitivity, on the order of a few picograms (Watanabe et al., 2003). Several positive-ion LC-MS and LC-MS/MS assays for PAF have been described using either normalphase or reversed-phase HPLC (Oda et al., 1995; Savu et al., 1996), but all these positive-ion methods are

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susceptible to interference from isobaric compounds. Interference from isobaric compounds does not allow detection of acylPAF in human umbilical vein endothelial cells (Oda et al., 1995). Specifically, reversedphase HPLC fails to resolve PAF from lysoPC, and normal-phase HPLC fails to resolve PAF from PFPC and related formylated lipids (Oda et al., 1995). Thin-layer chromatography (TLC) or silica solid-phase extraction of the lipid extract before reversed-phase LC-MS or LC-MS/MS removes some of the interfering lysoPC but can also introduce significant losses of PAF particularly when small samples are analyzed. A highly sensitive and specific LC-MS/MS PAF assay has been described by Owen et al. (2005b, 2007). This method can readily detect as little as 1 pg (1.9 fmol) of PAF and gives a linear response up to 1,000 pg of PAF. Positive-ion and negative-ion electrospray ionization were used with nitrogen as nebulizing and drying gas. PAF was detected in positive-ion mode as the transitions m/z 527←185 (d3-16:0 PAF), m/z 524 ←184 (unlabeled 16:0 PAF), m/z 550←184 (18:1 PAF), and m/z 552←184 (18:0 PAF). In negativeion mode, PAF was detected as m/z _ 511← _62 (d3-16:0 PAF), m/z _ 508← _59 (unlabeled 16:0 PAF), m/z _ 534← _59 (18:1PAF), and m/z _536← _59 (18:0 PAF) (Owen et al., 2005b). This method overcomes the artifacts from isobaric lipids, which limit the usefulness of other existing LC-MS/MS assays for PAF. Further developments of this method and recommendations for avoiding potential pitfalls have been described in detail by Owen et al. (2007). PAF quantitation performed by positive ionization detects as little as 1 pg injected into the column (signal/noise = 6), whereas negative ionization requires 100 pg (signal/ noise = 3.5) or 5 pg using different instrumentation (Kita et al., 2005). Limitations of this method, which offers high sensitivity (detection limit = 1 pg) selectivity and does not need sample derivatization, are the expensive instrumentation and the requirement of performing two HPLC runs per sample. Moreover, the levels of glycerophospholipid family members were measured by combining nano-flowrate HPLC with positive-ion mode ESI-MS and generating a two-dimensional profile. This procedure allowed identification and quantitation of PAF family members in the lipid extract of 70,000 PC12 cells (Whitehead et al., 2007).

3

Levels of PAF in Neural Cells and Mammalian Brain

Neural cells of mamalian brain produce PAF whose concentration varies according to brain areas and to physiological and pathological conditions. A selection of the available data is summarized in > Table 13‐1. PAF was first detected in the lipid extract of bovine brain using conventional GLC-MS (Tokumura et al., 1987). Other molecular species of ether phospholipids with an acetyl group at C-2 but differing in the length and unsaturation at the alkyl chain were also present in the same extract. The total amount of PAF molecular species was 40 pmoles/g of tissue. In addition, other molecular species of 1,2-diacyl-GPC with a short chain (2–4 carbons) at sn-2 position of glycerol were also found. These phospholipids are generally indicated as acylPAF (Tokumura et al., 1989, 1992). PAF was also found in rat brain (0.25  0.15 pmoles/g wet tissue) by bioassay, which measures the release of [3H] serotonin from rabbit platelets (Kumar et al., 1988). The molecule(s) were very likely of neural origin because their concentration increased noticeably following chemical- or electrical-induced convulsions, and similar results were obtained by treatment of perfused isolated rat brains with bicuculline (Kumar et al., 1988). The levels of brain PAF also increase as a consequence of ischemia (Braquet et al., 1988, 1989b; Domingo et al., 1994; Nishida and Markey, 1996). Particularly, PAF levels in ischemic rat brain increased noticeably (8,977.3  1,113 pg/g wet weight) as compared to sham-operated control (997.7  77 pg/g wet weight) (Domingo et al., 1994). In rat, PAF concentration was higher in the brain stem than in the cortex (12.8 vs. 7.0 pmoles/g tissue), and it was undetectable in normal spinal cord but increased after ischemia (Feuerstein and Yue 1989). Changes of PAF concentration were also analyzed in different brain areas during reperfusion after ischemia in gerbils by electron-capture negative chemical ionization GC/MS method (Nishida and Markey, 1996). In hippocampus, the level of PAF was 2.9 ng/g tissue in sham-operated controls, which increased to 6.0 ng/g tissue after 1 h of reperfusion. Unstimulated chick retina possesses measurable amounts of PAF (2.3 pmol/mg protein), which increase after stimulation with Ca2+ ionophore, acetylcholine, or dopamine (Bussolino et al., 1988). Human fetal neural cells in culture produce PAF and its levels increase after stimulation with acetylcholine, but atropine

Metabolism and functions of platelet-activating factor (PAF) in the nervous tissue

13

. Table 13‐1 PAF levels in mammalian brain PAF levels

Source Bovine (brain) Rat (brain)

Method GLC-MS Bioassay

Normal (pmol/g tissue) 40 0.38  10 3

Treatment (pmol/g tissue)

Rat (brain)

RIA

1.11

Chick (retina)

Bioassay

Rat (brain stem)

GLC-MS

2.3 (pmol/mg protein) 12.8

Rat (brain cortex)

GLC-MS

7

Rat (spinal cord)

Bioassay

nd

5.1 (ischemia)

Gerbil (brain) Gerbil (cortex)

RIA GC-MS

1.9 4.9

17.14 (Ischemia) 6.66 (HIV infection)

Gerbil (hippocampus)

GC-MS

4.79

7.43 (HIV infection)

20.4  10 3 (chemoconvulsion) 3.4  10 3 (electroconvulsion) 5.35 (ischemia) 5.93 (12-h reperfusion) 1.07 (7-day reperfusion) 11.3 (acetylcholine)

References Tokumura et al. (1989) Kumar et al. (1988)

Zhang et al. (2007)

Bussolino et al. (1986) Feuerstein and Yue (1989) Feuerstein and Yue (1989) Feuerstein and Yue (1989) Domingo et al. (1994) Nishida and Markey (1996) Nishida and Markey (1996)

reduces this effect (Sogos et al., 1990). The stimulation of cells with the neurotransmitter did not release PAF in the culture medium, whereas rat cerebellar granule cells released almost half of the produced lipid mediator (Yue et al., 1990). A sensitive procedure has allowed the evaluation of the changes in the levels of PAF family members in pheochromocytoma cells (PC12) induced to differentiate neuron-like cells with NGF (Whitehead et al., 2007). Differentiation caused a marked increase of C16:0 PAF, whereas C18:0 PAF was no longer detectable. Also, the levels of the corresponding lyso-derivatives increased in differentiated PC12 cells, and levels of C16:0 lysoPAF were particularly elevated.

4

Metabolism of PAF: General Aspects

4.1 Biosynthesis of PAF PAF biosynthesis occurs via two main pathways, the remodeling and the de novo pathways, and its degradation is carried out by specific intra- and extracellular acetylhydrolases (PAF-AHs), which are a subfamily of phospholipases A2 that removes the sn-2 acetyl group. The enzymes of the remodeling and de novo pathways have a relatively broad substrate specificity that provides a basis for heterogeneity in the molecular species of PAF produced by a given cell or tissue in response to specific stimuli. To date, several reviews have described, with considerable details, the properties and regulation of the enzymes of both PAF biosynthetic pathways (Prescott et al., 1990; Snyder, 1995a; Francescangeli et al., 2000; Arai, 2002; Farooqui and Horrocks, 2006). Particularly, findings on the role of

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the CoA-independent transacylase in the generation of lysoPAF (Uemura et al., 1991), the role of the PAFdependent transacetylation of the PAF-AH in the biosynthesis of PAF analogs (Arai, 2002; Karasawa et al., 2003), and, more recently, the cloning of acetyl-CoA:lysoPAF acetyltransferase (lysoPAF-AcT) markedly expanded the field of PAF metabolism.

4.2 De Novo Synthesis of PAF The de novo pathway is considered of physiological importance because it maintains the resting state levels of PAF in various tissues. This pathway is mainly responsible for PAF biosynthesis induced by neurotransmitters (Bussolino et al., 1986) and fatty acids (Vallari et al., 1990a, b; Venable et al., 1993), and by PMA and 1-oleyl-2-acetyl-sn-glycerol (Nieto et al., 1988a; Camussi et al., 1989). The 1-O-alkyl ether bond of 1-O-alkyl 2-lyso-sn-glycero-3-P (alkylGP), which represents the branch point of the initial step in the de novo synthesis of PAF, is formed via a unique reaction that converts acylDHAP into alkyl-DHAP by means of an alkyl-DHAP synthase (2.1.5.26) (> Figure 13‐2). This reaction, which involves the substitution of the acyl moiety of acyl-DHAP with a long-chain fatty alcohol, is a ratelimiting step in the de novo synthesis of PAF (Brown and Snyder, 1992). It has also been suggested that this enzyme is regulated by phosphorylation–dephosphorylation and that the phosphorylation is probably catalyzed by a protein kinase C (Heller et al., 1991; Baker and Chang, 1994). The ketone group of the alkyl-DHAP is then reduced by a NADPH-dependent oxidoreductase to form alkylGP, which can be converted through the de novo reaction steps to either PAF or the membrane PAF precursor in the remodeling route (alkylacylGPC).

. Figure 13‐2 Pathways for PAF biosynthesis

Metabolism and functions of platelet-activating factor (PAF) in the nervous tissue

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Three sequential steps, catalyzed by intracellular membrane associated enzymes, are involved in the conversion of alkylGP into PAF: (I) the acetylation by the acetyl CoA:1-alkyl-2-lyso-sn-glycero-3phosphate acetyltransferase (Lee et al., 1986; Snyder et al., 1992b); (II) the dephosphorylation by the 1-alkyl-2-acetyl-sn-glycero-3-phosphate phosphohydrolase (Lee et al., 1988; Snyder and Lee, 1992) producing 1-alkyl-2-acetyl-sn-glycerol (alkylacetyl-G); and (III) the transfer of phosphorylcholine from CDPcholine by a DTT-insensitive phosphocholinetransferase (PAF-PCT) (EC 2.7.8.2) to alkylacetyl-G (Renooij and Snyder, 1981; Francescangeli and Goracci, 1989; Lee and Snyder, 1992). The CDP-choline required in the final step is formed by CTP:phosphocholine cytidylyltransferase (EC 2.7.7.15), a rate-controlling enzyme in the production of PAF via the de novo pathway (Blank et al., 1988; Vallari et al., 1990a, b).

4.3 Remodeling Pathway of PAF Biosynthesis PAF biosynthesis via the remodeling pathway requires the formation of lysoPAF from membrane-bound alkylacylGPC, and its molecular species with arachidonic acid at C-2 (alkyl-20:4-GPC) is very likely the preferred substrate (Nieto et al., 1991; Uemura et al., 1991; Venable et al., 1991). The formation of lysoPAF occurs either by direct deacylation of membrane alkylacylGPC by a phospholipase A2 (PLA2), or by a transacylation reaction, alkyl-20:4-GPC being the acyl donor and 1-alk-1’-enyl-2-lyso-sn-glycero-3phosphoethanolamine (lysoPlsEtn) the acceptor (> Figure 13‐3) (Nieto et al., 1991; Uemura et al., 1991; Snyder et al., 1992a; Colard et al., 1993). In any case, the production of lysoPAF for PAF synthesis requires the hydrolysis of membrane-bound alkylacylGPC or of alkenylacylGPE (ethanolamine plasmalogen, PlsEtn) by phospholipases A2 (PLA2s). On the basis of their structural and catalytical properties, PLA2s constitute a superfamily (Balsinde et al., 2002; Brown et al., 2003; Schaloske and Dennis, 2006). Commonly, PLA2s are grouped in three families: Ca2+-dependent cytosolic PLA2 (cPLA2); Ca2+-independent cytosolic PLA2 (iPLA2); and secretory PLA2 (sPLA2). Four isoforms of cPLA2, included in group IV, have been identified and indicated as a, b, g, and d (Six and Dennis, 2000). Splice variants of iPLA2 have been reported to occur in various tissues (Larsson et al., 1998). sPLA2 isoforms are characterized by a low molecular weight (14–16 kDa) and a variable number of disulfide bridges (Murakami et al., 1995). Enzymes belonging to these three families are potentially involved in the generation of lysoPAF and consequently in PAF biosynthesis. Another enzyme, plasmalogen-selective PLA2 (Farooqui et al., 1995b), should also be considered for providing the substrate for the production of lysoPAF by the transacylase reaction. The variety of phospholipases A2 coexisting in any single cell population makes the establishment of their relative contribution to the production of lysoPAF for PAF synthesis quite difficult. On the basis of several experimental observations, it is conceivable that more than one phospholipase A2 is involved in PAF biosynthesis depending on the cell types and on their response to specific stimuli. The acetylation of lysoPAF (Figure 13‐3) occurs via acetyl-CoA:lysoPAF acetyltransferase (lysoPAFAcT) (EC 2.3.1.67). This membrane-bound enzyme, originally described by Wykle and coworkers (Wykle et al., 1980), catalyzes the transfer of the acetyl moiety from acetyl-CoA to the free hydroxyl group at the sn-2 position of lysoPAF. Since its first report, lysoPAF-AcT activity has been detected in several mammalian cell types and tissues (Wykle et al., 1980; Holland et al., 1992; Tonks et al., 2003; Fragopoulou et al., 2005; Shindou et al., 2005; Servillo et al., 2006). Cloning of this enzyme has been successfully achieved recently by Takao Shimizu’s group (University of Tokyo) (Shindou et al., 2007). The enzyme turned out to be a 60-kDa microsomal protein with three putative membrane-spanning domains. Surprisingly, this enzyme can utilize both acetyl-CoA and arachidonoyl-CoA as substrates for the synthesis of PAF or alkyl-20:4-GPC from lysoPAF, respectively. Under resting conditions, the enzyme prefers arachidonoyl-CoA and contributes to membrane biogenesis. Upon acute inflammatory stimulation with lipopolysaccharides, the activated enzyme utilizes acetyl-CoA more efficiently and produces PAF (Shindou et al., 2007). In the remodeling pathway, lysoPAF-AcT is also capable of acetylating lysoplasmalogen and 1-acyl-2lyso-GPC to form the corresponding plasmalogen and acyl analogs of PAF (acylPAF), respectively (Tessner and Wykle, 1987).

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. Figure 13‐3 Pathways of PAF degradation

Metabolism and functions of platelet-activating factor (PAF) in the nervous tissue

13

Stress-induced PAF synthesis via modulation of lysoPAF-AcT (Balestrieri et al., 2003, 2004; Corl et al., 2003; Owen et al., 2005a; Tosaki et al., 2007) is regulated positively by p38 MAPKand negatively by ERK-1/2 (Sakamoto et al., 2002; Owen et al., 2005a). However, MAPK-dependent regulation of lysoPAF-AcT may vary with agonists and cell types. Indeed, other reports indicate that PAF synthesis evoked by various agonists in neutrophyls and other cell types is ERK-1/2 dependent (Bernatchez et al., 2001; Baker et al., 2002; Russo et al., 2003).

4.4 PAF Degradation PAF degradation requires the removal of the acetyl group at the sn-2 position of the glycerol backbone catalyzed by PAF-acetylhydrolase (PAF-AH) (3.1.1.47), which is a Ca2+-independent enzyme belonging to group VII phospholipase A2 (Figure 13‐3) (Arai, 2002). To date, three types of PAF-AH isoforms have been identified in mammals: namely, the intracellular type I and II and the plasma type. Type I PAF-AH is a G-protein-like complex of two catalytic subunits (a1 and a2) and a regulatory b subunit. Type II PAF-AH is a single polypeptide and shows significant sequence homology with plasma PAF-AH. Type II PAF-AH is myristoylated at the N-terminus and, like other N-myristoylated proteins, is distributed in both cytosol and membranes. Plasma PAF-AH, which is also known as lipoprotein-associated phospholipase A2 (Lp-PLA2) due to its characteristics of association with plasma lipoproteins, shares 41% sequence identity with intracellular Type II PAF-AH, whereas both enzymes show less structural similarity to Type I PAF-AH. Type II PAF-AH, as well as plasma PAF-AH, may play roles as scavengers of oxidized phospholipids which are thought to be involved in diverse pathological processes including disorganization of membrane structure and PAF-like pro-inflammatory actions.

5

PAF Metabolism in Nervous Tissue

5.1 De Novo Synthesis of PAF in Nervous Tissue Similar to other cell types, neural cells can synthesize PAF by both the de novo and the remodeling pathways in mammals (Francescangeli and Goracci, 1989; Goracci and Francescangeli, 1991; Baker, 1995). The de novo pathway originates from acyl-DHAP that it is transformed into alkyl-DHAP by alkyl-DHAP synthase (EC 2.5.1.26) present in brain microsomes and using long-chain fatty alcohols as substrates (Hajra, 1970). Alkyl-DHAP is then reduced to AlkylGP by a NADPH-dependent oxidoreductase (EC 1.1.1.101) that is present in microsomes and peroxisomes (LaBelle and Hajra, 1972a, b). As mentioned above, AlkylGP is a common substrate for the synthesis of alkylacetylGP or alkylacylGP and, consequently, represents a branch point for the production of PAF by both biosynthetic pathways. In the cerebral cortex of immature rabbits, the highest specific activity of alkyl-GP acetyltransferase (alkylGP-AcT) was found in the rough microsomal fraction (Baker and Chang, 1993, 1996). The enzyme is also present in the nuclear fraction prepared from the same source and its specific activity is threefold higher than that of the microsomal enzyme (Baker and Chang, 1997, 1998). AlkylacylGP-AcT should possess essential sulfhydryl groups because its specific activity increases in the presence of reducing agents and it is greatly reduced by N-ethylmaleimide. In rat brain, as well as in other tissues, alkyacetylGP is then dephosphorylated (Lee et al., 1988), and the product, alkylacetylG, is converted to PAF by a phosphocholinetransferase (PAF-PCT), which has the highest specific activity in the microsomal fraction (Francescangeli and Goracci, 1989). Rat-brain PAF-PCTshares some properties with the enzyme synthesizing alkylacylGPC or diacylGPC (Binaglia et al., 1973; Radominska-Pyrek et al., 1977) because it requires Mg2+ and is inhibited by Ca2+ (Francescangeli et al., 2000). However, the enzyme is not inhibited by DTT similar to that described in other tissues or cell preparations (Snyder, 1995b). Thus, PAF-PCT should be an enzyme distinct from that synthesizing other cholinephosphoglycerides. A 75% reduction of the rate of PAF formation by PAF-PCT was observed with CMP in the incubation system at the same concentration as CDP-choline (1 mM) (Francescangeli and Goracci, 1989), indicating

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the possibility that PAF-PCT catalyzes a reversible reaction as reported for DTT-sensitive phosphocholinetransferase (Goracci et al., 1981). This hypothesis was confirmed by the CMP-dependent degradation of labeled PAF, first to 1-alkyl-2-acetyl-sn-glycerol (alkylacetylG) and then to alkylglycerol, which was reduced by the presence of CDP-choline (Francescangeli et al., 1997b, 2000).

5.2 PAF Biosynthesis by the Remodeling Pathway in Nervous Tissue LysoPAF AcT activity for PAF biosynthesis by the remodeling pathway was reported in chick retina (Bussolino et al., 1986), rat-brain microsomes (Goracci and Francescangeli, 1991), and immature rabbit brain (Baker, 2000). The specific activity of rat-brain microsomal lysoPAF-AcT was almost one-third of that of PAF-PCT when their activities were assayed under optimal conditions (Francescangeli et al., 2000). LysoPAF-AcT activity was also detected in neuronal nuclei, mainly located in the envelope, and it is more active in the presence of reducing agents similar to alkylGP-AcT (Baker and Chang, 1997). The presence of a PLA2 in rat brain and human cerebral cortex preparations, which was able to hydrolyze alkyl(long-chain)acylGPC producing 1-alkyl-2-lyso-GPC, a compound later called lysoPAF, was first reported by Woelk et al. (1974). On the basis of more recent knowledge, lysoPAF production could be due to more than a single enzyme belonging to different families because cPLA2, iPLA2, sPLA2, and plasmalogen-selective PLA2 (PlsEtn-PLA2) are present in brain tissue and neural cells (Bazan, 2003; Farooqui and Horrocks, 2006, 2007; Sun et al., 2007). However, it is still unclear what their relative contribution to the production of lysoPAF from alkylacylGPC is. Due to the relative high content of arachidonic acid at C-2 of ether lipids, group IV cPLA2 is often indicated as the isoform most likely involved in lysoPAF production, but enzymes belonging to other PLA2 families could also participate in this process. For instance, PlsEtn-PLA2 has been purified from bovine brain (Hirashima et al., 1992) and it might also contribute to the production of lysoPAF whenever it could hydrolyze alkylacyl-GPC. Different isoforms of secretory PLA2 are also expressed in neural cells and might contribute to the formation of lysoPAF as well. Particularly, group IIA sPLA2 is the main isoform present in particulate ratbrain fraction (Yang et al., 1999) and it has been detected in rat-brain mitochondria (Macchioni et al., 2004). Another isoform (group V sPLA2) is localized in the nuclear fraction of neuronal and glial cells from rat-brain cortex (Nardicchi et al., 2007). Moreover, upregulation of group IIA sPLA2 mRNA expression was observed in astrocytes in response to transient focal cerebral ischemia in the rat brain (Lin et al., 2004). Interestingly, a crosstalk between secretory sPLA2 and cPLA2 in neuronal signal transduction has been proposed (Kolko et al., 2003) because exogenous sPLA2 activates cPLA2 and leads to the formation and accumulation of bioactive lipids, such as PAF, free arachidonic acid, and eicosanoids. An alternative route for the production of lysoPAF is present in the brain (Blank et al., 1995), which also possesses CoA-independent transacylase activity localized in the microsomal fraction (Ojima et al., 1987; Masuzawa et al., 1989) and neuronal nuclei (Baker and Chang, 2000). In this case, lysoPAF is produced from alkylacylGPC by the transfer of a polyunsaturated fatty acid from its C-2 to 2-lyso-phosphoglycerides produced by a PLA2 action on other glycerophospholipids. 1-AlkenylGPE is a better acceptor of the acyl group than 1-acylGPC for the neuronal nuclear transacylase even in the presence of inhibitors of lysophospholipases (Baker and Chang, 2000).

5.3 Degradation of PAF in the Nervous Tissue Mammalian brain contains at least three intracellular isoforms, and PAF-AH(Ib) is the best characterized. This isoform contains a heterodimer of two homologous catalytic subunits a1 and a2, with a relative molecular mass of 26K each, and a non-catalytic 45K b-subunit, a homolog of the b-subunit of trimeric G proteins (Ho et al., 1997; Manya et al., 1999; McMullen et al., 2000; Sheffield et al., 2001). The amino acid sequences of the three subunits showed extremely high homologies among mammalian species (Hattori et al., 1994a, 1994b, 1995; Adachi et al., 1995; Albrecht et al., 1996; Watanabe et al., 1998). Only one amino acid substitution has been observed in the human a2 subunit in comparison with the other three species,

Metabolism and functions of platelet-activating factor (PAF) in the nervous tissue

13

and similarly the amino acid sequences of the b subunit from the mouse, rat, and cow are identical and only one amino acid substitution has been observed in the human b subunit. The a1 and a2 subunits, which show approximately 60% amino acid homology, have a catalytic center (Hattori et al., 1994b, 1995; Adachi et al., 1995). As will be discussed below with more details, the b subunit gene was found to be identical to the human LIS1 gene, which is the causative gene of type I lissencephaly (Hattori et al., 1995; Reiner et al., 1995; Peterfy et al., 1998). All three subunits are expressed in embryonic brain, whereas only the a2 and b subunits were detected in the adult brain. Molecular characterization of a translocation t(1;19)(q21.3; q13.2) has been reported in a woman with mental retardation, ataxia, and atrophy of the brain (Nothwang et al., 2001). Sequence analysis of the breakpoints revealed an Alu-repeat-mediated mechanism of recombination that led to the truncation of two genes: the kinase CLK2 and a1 subunit of PAF-AH (I). One expressed fusion gene encodes the first 136 amino acids of the a1 subunit followed by the complete CLK2 protein. The truncated a1 protein loses its hydrolytic activity and the possibility to interact with the b subunit suggesting the key role of the catalytic subunit PAF-AH (I) in brain development. Expression of all three PAF-AH (I) proteins (1, 2, and LIS1) as well as PAF-AH (II), but not plasma PAFAH or PAF receptor (PAFR), has been demonstrated in PC12 cells (Bonin et al., 2004). Induction of PAFAH (I)a2, but not PAF-AH (I) a1, regulates the duration of apoptotic signaling initiated by PAF challenge. By pharmacological inhibition of PAF-AH I and II activity and downregulation of PAF-AH (I) catalytic subunits by RNA interference, the PAFR-independent death pathway has been shown to be regulated by PAF-AH (I) and, to a lesser extent, by PAF-AH (II). Moreover, the anti-apoptotic actions of PAF-AH (I) are subunit-specific. PAF-AH (I)a1 regulates intracellular PAF concentrations under normal physiological conditions, but its expression is not sufficient to reduce an acute rise in intracellular PAF levels. PAF-AH (I) a2 expression is induced when cells are deprived of serum or exposed to apoptogenic PAF concentrations, limiting the duration of pathological cytosolic PAF accumulation (Bonin et al., 2004). Type II PAF-AH is a monomeric enzyme of 40 kDa with significant homology to plasma-type PAF-AH and not to any subunit of PAF-AH (I) (Arai, 2002; Karasawa et al., 2003). Both monomeric enzymes hydrolyze not only PAF but also the short-chain phospholipid and oxidized fragment of polyunsaturated fatty acid at the sn-2 position. Thus they terminate PAF-mediated signaling and attenuate the toxicity induced by oxidized PAF-like lipids (Matsuzawa et al., 1997; Arai, 2002; Karasawa et al., 2003; Kono et al., 2008). In contrast to PAF-AH (I), PAF-AH (II) is easily inactivated by sulfhydryl agents such as 5,5’-dithio-bis (2-nitrobenzoic acid), suggesting the presence of a free cysteine residue essential for catalysis. PAF-AH (II) has been shown to be an antioxidant phospholipase in a number of systems. This enzyme recognizes the redox state of the cell environment, changes location in response to cellular requirements, and prevents damages from lipid peroxidation. Indeed, PAF-AH (II) translocates to the membrane during oxidative stress and inhibits oxidative stress-induced apoptosis, presumably by scavenging oxidized phospholipids. Interestingly, as described by Bae et al. (2000), this enzyme possesses transacetylase as well as hydrolase activity under certain conditions, thus diversifying the biological function of PAF by producing different lipid mediators such as analogs of PAF and C2-ceramide. Transfection of the plasma-type PAF-AH gene attenuates glutamate-induced apoptosis in cultured rat cortical neurons (Prescott et al., 1990), and overexpression of PAF-AH (II) in Chinese hamster ovary K1 cells suppresses apoptotic death induced by tert-butylhydroperoxide, most likely by its antioxidant effects (Venable et al., 1993). More recently, it has been shown that PAF-AH (I)I overexpression exerts neuroprotective effects in mouse models of focal cerebral ischemia, suggesting that the delivery of recombinant PAF-AH (II) to post-ischemic neurons might have a therapeutic significance to prevent neuronal death after transient ischemia (Umemura et al., 2007).

5.4 Regulation of PAF Metabolism in Neural Cells The presence of two distinct pathways, both potentially able to contribute to the production of PAF in neural cells under physiological and pathological conditions, raises the question whether they correlate to specific cell functions and might be differently involved in the onset or aggravation of neurological diseases.

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PAF is directly produced by the reactions catalyzed by PAF-PCT and lysoPAF-AcT, and the rate of its production depends on the activation status of the enzymes and on the availability of sufficient amounts of their substrates: CDP-choline and alkylacetylG for the last reaction of the de novo pathway and lysoPAF and acetyl-CoA for the remodeling pathway. CDP-choline is produced in neural cells by choline cytidylyltransferase from phosphorylcholine and CTP, and this is considered the limiting step for the synthesis of cholinephosphoglyceride, including PAF (Blank et al., 1988). The availability of alkylacetylG depends on the activity of alkylGP-AcT, which represents the first committed step for the synthesis of PAF by the de novo route. In rabbit brain, the enzyme has a microsomal and nuclear localization. Evidence for the upregulation of the microsomal enzyme by phosphorylation has been reported (Baker and Chang, 1994), but the protein kinase responsible for the activation of the enzyme was not identified. Further studies (Baker and Chang, 1997) have shown that the nuclear fraction of neuronal cells from rabbit immature brain possesses both alkylGP and lysoPAF acetyltransferases. Both enzymes are more active when phosphorylated because protein phosphatase inhibitors preserve their activities. However, different protein phosphatases seem to be involved in the inactivation of the two nuclear enzymes. The presence of group V -sPLA2 in the nuclei of neural cells (Macchioni et al., 2004; Nardicchi et al., 2007) might provide the substrate for lysoPAF-AcT and consequently contribute to the synthesis of PAF in the nuclei. Evidence for the upregulation of lysoPAF-AcT by phosphorylation, apparently by PKC, has been also reported for the rat-brain microsomal enzyme as well as rat leukocytes (Hayashi et al., 1991). However, the involvement of other enzymes cannot be excluded such as Ca2+-calmodulin protein kinase (Domenech et al., 1987), PKA (Nieto et al., 1988b), and p38 MAPK (Nixon et al., 1999; Owen et al., 2005a; Shindou et al., 2007), which activate lyso-PAF AcT in other cell types. The concentration of intracellular Ca2+ is also an important factor for the regulation of the two PAFsynthesizing pathways in neural cells and in other cell types. Following an increase of intracellular Ca2+, the remodeling pathways should provide the most important contribution to PAF biosynthesis because cPLA2, sPLA2, and lysoPAF-AcT become more active (Francescangeli et al., 2000). This assumption is also supported by the observation that the treatment of chick retinas with PLA2 inhibitors blocked the production of PAF induced by A23187. In addition, this Ca2+ ionophore increases the activity of chick retina lysoPAF-AcT, but it has no effect on PAF-PCT (Bussolino et al., 1986). Another relevant difference between the two pathways is the energy requirement. The de novo pathway needs ATP and CTP for the production of CDP-choline, which is the substrate for PAF-PCT, whereas microsomal lysoPAF-AcT is inhibited by ATP (Baker and Chang, 1994; Francescangeli et al., 2000). Thus, it has been suggested that under conditions of low cellular energy, as during cerebral ischemia, the decrease of ATP and the increase of Ca2+ concentration should switch the biosynthesis of PAF from the de novo to the remodeling pathway (Francescangeli et al., 2000). Baker’s group has extensively studied the acetylation reaction of alkylGP and lysoPAF acetyltransferases in rabbit brain (Baker, 2000) and shown that microsomal and nuclear alkylGP-AcT of rabbit brain are inhibited by MgATP (Baker and Chang, 1994, 1996, 1998). These studies have also suggested an important contribution of the de novo pathway to PAF formation when ATP cellular levels decrease. However, in vivo, the removal of this inhibitory effect would lead to the accumulation of alkylacetylG, but the scarce availability of the other substrate for PAF-PCT, i.e., CDP-choline, would reduce its conversion to PAF. The cellular accumulation of alkylacetyG might activate PKC (Stoll et al., 1991) and lead to the activation of lysoPAF-AcT by phosphorylation. Other possible modulators for PAF biosynthetic pathways are free fatty acids, acyl-CoAs, and lysophosphoglycerides other than lysoPAF. A concentration-dependent inhibition of rat-brain microsomal lysoPAF-AcT was observed with arachidonate at micromolar concentrations. This effect was greater with its CoA-thioester, and PAF-PCT was almost unaffected (Francescangeli et al., 2000). Interestingly, oleic acid increases the production of lysoPAF from alkylacylGPC in the presence of 1-acyl-GPC by transacylation, and it has been hypothesized that this mechanism for the production of lysoPAF might be relevant during ischemia where both free fatty acids and lysophospholipids are produced by the activated PLA2 (Baker and Chang, 2000).

Metabolism and functions of platelet-activating factor (PAF) in the nervous tissue

13

The increased availability of lysoPAF, produced by direct hydrolysis of alkylacyl-GPC or by transacylase, favors the production of PAF by the remodeling pathway in ischemic brain because lysoPAF-AcT is also more active (Francescangeli et al., 1996). On the other hand, lysoPAF availability is reduced by its conversion to alkylacylGPC by the lysoPC acyltransferase activity. This enzyme activity coexists with that of lysoPAF-AcT in the same protein, which is expressed in brain and appears to predominate in the resting conditions of the cells (Shindou et al., 2007). Levels of PAF are also modulated by the activities of the three intracellular isoforms of PAF-AH present in nervous tissue. Particularly, as reported above in more detail, PAF-AH (I) appears to play a primary role in the control of PAF concentration in the brain counteracting the continuous PAF production (Chen et al., 2007). This allows the maintenance of physiological levels avoiding its accumulation, which is detrimental for normal brain functions.

6

PAF Receptors and Cell Signaling

High-affinity binding sites were identified by kinetic and pharmacologic studies in microsomal and synaptosomal fractions of rat-brain homogenate (Marcheselli et al., 1990). PAF binding to the intracellular receptor (microsomal) is specifically reduced by ginkogolide BN 50730, whereas BN 52021 is a specific antagonist of the extracellular receptor (synaptosomal). The latter corresponds to the plasma membrane receptor (PAFR), which has been well characterized (Domingo et al., 1988; Marcheselli et al., 1990; Bito et al., 1992, 1993; Diserbo et al., 1995; Lalouette et al., 1995; Shimizu et al., 1996; DeCoster et al., 1998; Aihara et al., 2000; Clark et al., 2000; Ishii et al., 2002). Structural features and properties of PAFR, as well as the cellular effects of its activation, have been reviewed (Ishii and Shimizu, 2000). Attempt for isolating PAFR from tissues was unsuccessful. However, molecular biology techniques have allowed cloning PAF receptor cDNAs from various cell types of mammalian tissues including human tissues (Nakamura et al., 1991; Kunz et al., 1992; Sugimoto et al., 1992). Human PAFR gene is localized in chromosome 1 and its expression generates two different mRNA species driven by distinct promoters. However, only one transcript is generated in human brain (Mutoh et al., 1993). Human and guinea pig PAFRs consist of a single polypeptide chain composed of 342 amino acids with seven transmembrane domains typical of the G-protein coupled receptor superfamily (Ishii et al., 2002). Three-dimensional (3D) models of PAFR were constructed on that of bacteriorhodopsin (Kajihara et al., 1994) and, more recently, another model has been developed by a hierarchial approach integrating homology modeling, molecular docking, and molecular dynamics (Gui et al., 2007). The mechanisms regulating PAFR by agonists and antagonists were then studied by using this 3D model. PAFR is expressed in numerous mammalian tissues, and its expression is induced during differentiation of HL-60 cells (Muller et al., 1991; Nakamura et al., 1991) and by cytokines in inflammatory cells (Weber et al., 1993). In the nervous tissue, the expression of PAFR has been demonstrated by radioligand binding assay (Bito et al., 1992) and by Northern analysis (Bito et al., 1994; Izumi et al., 1995). In rat brain, PAFR is expressed in almost all areas but its level is rather low in thalamus and cerebellum (Bito et al., 1992). In situ hybridization has shown that PAFR is highly expressed in rat microglial cells and, to a lesser extent, in the neuronal population (Mori et al., 1996). Cell responses to the activation of PAFR are mediated by PTX-sensitive (Gq/11) or PTX-insensitive G-proteins (Gi and/or G0) (Ali et al., 1994; Honda et al., 1994; Ishii and Shimizu, 2000). The interaction of activated PAFR with G-proteins triggers a complex intracellular signaling. Indeed, studies in many different cellular models have revealed the participation of Ca2+, cAMP, diacylglycerols, and IP3 as second messengers in the activation of several protein kinases (MAPK, PKC, PI3K, tyrosine kinases and G-protein receptor kinases) and phospholipases (PLCb, PLCg, PLA2, PLD) (reviewed by Ishii and Shimizu, 2000). The stimulation of rat-brain cortical slices with PAF at nanomolar concentration produced a significant increase of the incorporation of labeled phosphate into phosphoinositides and phosphatidic acid and induced a rapid and transient accumulation of inositol-1,4,5-trisphosphate, indicating the activation of the hydrolysis of PIP2 by phosholipase C (Catalan et al., 1992). Further studies have shown that the stimulation

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of an immortalized hippocampal cell line (HN33.11) with PAF also leads to the accumulation of inositol phosphates by a receptor-mediated mechanism and the involvement of GaI 1/2 and Gaq (Shi et al., 1996). This mechanism is responsible for the PAF-induced elevation of intracellular Ca2+ concentration observed in other neural cell lines NG108–13 as well as PC12 cells (Kornecki and Ehrlich, 1988, 1991), NCB-20 (Yue et al., 1992) and microglial cells (Mori et al., 1996). The activation of PAFR in neural cells induces also the expression of immediate-early genes (Dell’Albani et al., 1993; Bazan and Allan, 1996), but the mechanisms eliciting nuclear effects induced by PAF are still not completely elucidated. The possibility exists that the activated PAFR is internalized by endocytosisassociated b-arrestin, leading to the activation of Ras and MAPKs similar to other G protein-coupled receptors (Pierce et al., 2001). PAF might also induce its own production by one of the biosynthetic pathways (Bussolino and Camussi, 1995) and produce nuclear effects by its binding to intracellular receptors (Marcheselli et al., 1990; Bazan, 1998). The presence of PAF-synthesizing enzymes in cell nuclei (Baker and Chang, 1996) and the nuclear localization of a PAF receptor support this hypothesis (Marrache et al., 2002, 2005; Zhu et al., 2006). Interestingly, the stimulation of isolated nuclei of porcine cerebral microvascular endothelial cells with non-hydrolysable PAF analog induces the expression of COX-2 and iNOS, which is blocked by a selective PAFR antagonist (Marrache et al., 2002). The creation of PAFR overexpressing mice and PAFR knock-out mice might provide another approach for studying the role of PAF in brain functions and dysfunctions (Ishii and Shimizu, 2000). However, the utility of PAFR-KO mice for this purpose is questionable because they did not show relevant brain malformation and had normal excitatory synaptic transmission (Ishii et al., 1998).

7

PAF Receptors Agonists and Antagonists

Natural and synthetic agonists of PAFRs have been identified and their potency was compared with that of 16:0 PAF, which is the most effective in inducing platelet aggregation and release of labeled serotonin (Braquet and Godfroid, 1987; Shen et al., 1987). The structural requirements for inferring PAF-like activities have been extensively investigated by chemical modifications. The L (R) configuration of the chiral center at C-2 of the glycerol backbone is absolutely necessary for the activity of PAF (Wykle et al., 1982).The substitution of the alkyl chain at C-1 with alkenyl or acyl groups, or the elongation of the acyl chain at C-2, greatly reduces the potency of the agonist. The modification of the polar head group at C-3 position decreases the potency in the order choline > dimethyl ethanolamine > monomethylethanolamine > ethanolamine, the last one being completely inactive (Satouchi et al., 1981). A synthetic non-hydrolysable analog, 1-O-alkyl-2-N-methylcarbamyl-GPC (mc-PAF), is a potent PAFR agonist with a dissociation constant 5-times greater than that for PAF, which correlates with the potency of its biological effects (O’Flaherty et al., 1987). The resistance of this synthetic agonist to the metabolic degradation by cellular and plasma PAF-AH is particularly useful for its use in experimental studies. The involvement of PAF in a variety of pathophysiological events has led to intense research for the identification of natural and synthetic antagonists for possible clinical applications (Koltai et al., 1994; Maclennan et al., 1996b). Among natural antagonists, ginkgolides are by far the most known and have been used both in experimental studies and in clinical applications (Birkle et al., 1988; Guinot, 1994; Maclennan et al., 1996a; Debek et al., 1999; Wittwer et al., 2001; Hostettler et al., 2002). Ginkgolides are terpenoids isolated from the leaves for Gingko biloba, the ancient tree with known beneficial effects on human health, according to Chinese traditional medicine. They are cage molecules with 20 carbons and six rings and differ from each other in the number and position of hydroxyl groups. Ginkgolide B (BN 52021) is the most potent PAFR antagonist that locks PAFR in its inactive state as shown by molecular dynamics simulation studies (Gui et al., 2007). Ginkgolides are components of the standardized G. biloba leaves extract (EGb761) possessing also antioxidant properties and exerting neuroprotective effects in cellular models of Alzheimer’s disease (Bastianetto and Quirion, 2002; Eckert et al., 2005). Structurally related PAF antagonists have been also synthesized. This class of compounds includes CV-3988 and CV-6209 (Handley, 1990; Gati et al., 1991), TCV-309 (Yamaguchi et al., 1999), SDZ-64–412 (Handley et al., 1988), and others (Koltai and Braquet, 1994).

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Other synthetic compounds, unrelated to PAF structure, are potent PAFR antagonists. Among this class, WEB-2086 (Apafant) is more potent than BN- 52021 (Ikegami et al., 1992). WEB-2170 (bepafant) and WEB-2347 were then developed and these compounds have greater potency or a longer half-life than apafant (Heuer et al., 1990; Heuer, 1991). ABT-491 (Albert et al., 1997) is another synthetic selective PAFR antagonist that has been shown to inhibit neuronal apoptosis in neonatal rats following hypoxic ischemic insults (Bozlu et al., 2007). PMS777 is a tetrahydrofurane derivative, which, in addition to the blockade of PAFR, also inhibits acetylcholine esterase (Letexier et al., 1996). Thus, it has been proposed as a drug with beneficial effects on Alzheimer’s disease (Li et al., 2007). Recently, a piperazine derivative with dual PAF-antagonist and anti-HIV properties has been reported (Sallem et al., 2006).

8

Functions of PAF in the Nervous Tissue

Several lines of evidence indicate the participation of PAF in a number of brain functions. These include the synaptic transmission and, particularly, the mechanisms involved in long-term potentiation (LTP), a phenomenon that is associated with learning and memory formation. In addition, PAF participates in some mechanisms involved in the proliferation and differentiation of neural cells but its role in these processes is still largely unknown.

8.1 Synaptic Transmission The first evidence indicating a possible connection of PAF with neurotransmission was the observation that psychoactive drugs such as triazolobenzodiazapines inhibited PAF-induced platelet aggregation (Kornecki et al., 1984). Then, Bussolino and colleagues (1986, 1988) reported that PAF synthesis was increased by stimulation of mature or immature chicken retina with neurotransmitters, particularly acetylcholine and dopamine. Under their experimental conditions, PAF was not detectable in the medium but another study demonstrated its release from rat cerebellar granule cells (Yue et al., 1990). The release of dopamine from PC12 cells stimulated by PAF (Bussolino et al., 1988) provided further evidence for the involvement of this mediator in synaptic transmission. Electrophysiological studies demonstrated that the addition of 1-Oalkyl-2-N-methylcarbamyl-GPC (mc-PAF) to cultured primary hippocampal neurons specifically enhanced excitatory synaptic transmission but did not interfere with inhibitory responses mediated by GABA. Because PAF-induced increase of excitatory effect was blocked by the PAFR antagonist BN 52021 and not altered by exogenously added glutamate, a link between the presynaptic action of PAF and the release of glutamate was postulated (Clark et al., 1992).

8.2 LTP and Memory The inhibition of long-term potentiation (LTP) by extracellular PAFR antagonist in the CA1 region of hippocampus was first reported by Del Cerro et al. (1990). Further studies have shown that various PAFR antagonists, having different potency, blocked or reduced LTP induced by high-frequency stimulation (HFS) in slices of rat hippocampus. This effect was partially reversed by mc-PAF. Evidence was also provided for the involvement of NMDA receptors (NMDAR) (Arai and Lynch, 1992). In the same experimental model, Wieraszko et al. (1993) reported that PAF-induced LTP was also blocked by the PAFR antagonists BN52021 and WEB 2086. PAF (100 nM) caused a fourfold increase in the population spike, which was almost double that induced by HFS stimulation which was not affected by PAFR antagonists. The involvement of NMDAR was clearly demonstrated, because the increase of population spikes induced by PAF was almost completely blocked by APV, a competitive antagonist of glutamate NMDAR, and by MK 801, a blocker coupled to cation channel of NMDAR. The role of PAF as a retrograde messenger in the induction of LTP was then demonstrated in the CA1 region of hippocampus (Kato et al., 1994) and in dentate gyrus (Kato and Zorumski, 1996). Support to the hypothesis that post synaptically released PAF,

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upon NMDAR activation, could induce a pre-synaptic PAFR-mediated increase of Ca2+ concentration and a consequent further release of glutamate was brought by Kornecki et al. (1996). Indeed, HFS of hippocampal slices induced the release of PAF in the extracellular medium, and the stimulation with 50 nM PAF of hybrid pyramidal neurons (HN33 cells), which releases glutamate by a Ca2+-dependent mechanism, increased the Ca2+ concentration. The generation of PAFR-deficient mice (Ishii et al., 1998) provided a potential tool for investigating the role of PAF in brain function. However, LTP was only slightly attenuated in dentate gyrus neurons of PAFR (–/–) mice (Chen et al., 2001). Using the same model, it was concluded that PAF is not required for LTP in hippocampal CA1 region (Kobayashi et al., 1999). The involvement of PAF in LTP is not restricted to hippocampal neurons, but it regards other neuronal populations of different brain areas such as medial vestibular nuclei of brain stem (Grassi et al., 1998) and somatosensory cortex (Heusler and Boehmer, 2007). HFS stimulation of primary vestibular afferents induces LTP in the medial vestibular nuclei through the activation of post-synaptic glutamate NMDAR (Capocchi et al., 1992). In this experimental model, HFS and mc-PAF induced LTP. The exposure of rat brain stem slices to synaptosomal PAFR antagonist BN52021 did not alter basal vestibular potentials but reduced their amplitude after HFS or addition of mc-PAF (Grassi et al., 1998). A similar effect, but at higher concentrations, was obtained by the intracellular binding-site antagonist CV-6209 (Marcheselli et al., 1990). Once LTP was induced, both antagonists had no effect on LTP itself, indicating that PAF is necessary for LTP induction but not for its maintenance. The onset of LTP, induced by PAF analog, was prevented by the NMDAR antagonist AP-5, confirming the participation of glutamate in the mechanism. The exposure of rat brain stem slices to AIDA, an antagonist of group I metabotropic glutamate receptors (group I-mGluR), prevented mc-PAF-induced LTP in medial vestibular nuclei (Grassi et al., 1999). However, once LTP was established, the same compound reversibly reduced it. This observation indicates that vestibular LTP requires pre-synaptic events for its consolidation. The interaction of PAF, which is synthesized at post synapsis and released in the extracellular medium, with group I mGluRs is necessary for a long-lasting increase of synaptic effeciency. Then, the following mechanism for the participation of PAF in the induction of LTP as retrograde messenger can be hypothesized (> Figure 13‐4): (1) The activation of NMDAR stimulates the synthesis of PAF at the post-synaptic neuron; (2) PAF is then released, activates its presynaptic receptor, and induces a further release of glutamate. The observation that the specific activities of the enzymes catalyzing the last reaction of PAF biosynthesis (lysoPAF-AcT and PAF-PCT), assayed under optimal conditions in the homogenates of rat brain stem slices, are both elevated during HFS-induced LTP in medial vestibular nuclei supports this hypothesis (Francescangeli et al., 2002). These effects are specific because DTTinsensitive phosphocholinetransferase, the enzyme synthesizing phosphatidylcholine, was not affected by HFS stimulation. Both lysoPAF-AcT and PAF-PCT resulted already activated 5 min after application of HFS but the activation of the latter enzyme lasted longer. Superfusion of rat-rain slices with AP-5 before the induction of LTP by HFS in medial vestibular nuclei did not allow the activation of PAF-synthesizing enzymes were not activated when slices of medial vestibular nuclei from rat brain were superfused with AP-5 before the induction of LTP by HFS (Francescangeli et al., 2002). This indicates that the interaction of glutamate with NMDAR is required for increasing the activity of PAF-synthesizing enzymes, very likely, by phosphorylation (Francescangeli et al., 2000). On the basis of these results, it was proposed that the activation of NMDAR should increase the postsynaptic activity of Ca2+-dependent PLA2, which produces the substrate for lysoPAF-AcT, which is also more active due to the higher Ca2+ concentration and, very likely, is phosphorylated. According to this hypothesis, during the first minutes after stimulation of NMDAR, PAF biosynthesis should be essentially due to the remodeling pathway but the long-lasting activated status of PAF-PCT might be important for maintaining elevated post-synaptic levels of PAF. LTP and certain forms of learning and memory are believed to share some common cellular mechanisms (Collingridge and Bliss, 1995; Medina and Izquierdo, 1995; Bazan et al., 1997; Cain, 1997). Evidence for a link between PAF-induced LTP, learning, and memory formation was provided by experiments with animal models (Jerusalinsky et al., 1994; Izquierdo et al., 1995; Packard et al., 1996; Teather et al., 1998, 2001).

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. Figure 13‐4 Schematic representation of the involvement of PAF in the induction and maintenance of LTP. Depolarization of pre-synaptic terminals releases glutamate which induces the increase of post-synaptic Ca2+ concentration by the activation of NMDAR. The increase of the concentration of this cation activates post-synaptic sPLA2, very likely sPLA2-IIA, and cPLA2 producing lysoPAF from alkylacylGPC and polyunsaturated fatty acids including arachidonic acid (AA). LysoPAF is then converted to PAF by lysoPAF-AcT which is also activated by Ca2+. AA could be used for eicosanoid synthesis. Thus, post-synaptic PAF concentration increases and, subsequently activates intracellular PAF receptor (PAFIR) inducing the expression of genes involved in long-term effects of synaptic functions. In addition, PAF is released in the extracellular space and activates presynaptic PAF receptor (PAFR) with a further increase of Ca2+ concentration and a consequent release of glutamate in the synaptic cleft. Once established the synaptic potentiation is maintained by the activation of presinaptic metabotropic glutamate receptor (mGluR)

Particularly, the infusion of PAF antagonist (BN 52021) into hippocampus or amygdale before or immediately after training in step-down inhibitory avoidance resulted to be amnesic (Jerusalinsky et al., 1994), and the post-training intra-striatal infusion of mc-PAF enhanced memory in the cued water maze task (Packard et al., 1996).

8.3 Proliferation and Differentiation of Neural Cells The addition of 50 nM PAF to the culture medium of hybrid neural cells (NG108-15) arrested their growth and caused their morphological differentiation, inducing neurite extension (Kornecki and Ehrlich, 1988). At higher concentrations, PAF becomes neurotoxic and causes cell death. The involvement of PAF in neuronal differentiation was also reported in cell cultures from embryonic rat cerebra in which it stimulates the formation of axon-like extensions (Ved et al., 1991). Following these observations and considering that neural cells possess both the substrates and the enzymes for the endogenous production of PAF, it was investigated whether the metabolism of this lipid mediator undergoes changes during proliferation and differentiation (Francescangeli et al., 1993, 1996, 1997a). To this aim, PAF-synthesizing enzymes and PAF-AH were assayed in primary cultures of neuronal and glial cells obtained from chick embryo hemispheres. The specific activity of PAF-PCT was nearly

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3-times higher than that of lysoPAF-AcT in the homogenate of cell suspensions of 8-day chick embryo hemispheres, consisting essentially of undifferentiated neuronal cells, whereas they were similar in the homogenates of brains from 14-day embryos which consisted of glial and differentiated neuronal cells. A sharp decrease of PAF-PCT was observed when neurons ceased to proliferate. A decrease of the activity of PAF-PCT was also observed in LA-N-1 neuroblastoma cells when they were induced to differentiate by retinoic acid (Francescangeli et al., 1997a). In chick embryo neuronal primary cultures, the activities of both PAF-PCT and lysoPAF-AcT were similar and increased during the period of cell maturation and the formation of synaptic-like junctions, whereas the activity of PAF-AH remained constant. On the other hand, the specific activity of PAF-PCT did not change during glial cell proliferation, but that of lysoPAFAcT increased reaching a maximum that was five- to sixfold that of PAF-PCT at cell confluence. Considering that both enzymes were assayed at their optimal conditions and with saturating concentrations of substrates, it was concluded that both the de novo and the remodeling pathways might be participating in the production of PAF in neuronal and glial cells but their relative contribution depending on the concentration of the substrates (i.e., CDP-choline and alkylacylG for PAF-PCTand lysoPAF for lysoPAF-AcT) and on the activation status of PAF-synthesizing enzymes. On the basis of these observations, it may be hypothesized that PAF-PCT is in an activated status during the proliferation of undifferentiated neuronal cells and becomes downregulated when they start to differentiate. At a later developmental stage of neuronal cells, both the de novo and the remodeling pathways may equally contribute to PAF synthesis. In this case, the availability of their substrates may limit their relative contribution. The involvement of PAF in neuronal differentiation has been also demonstrated by the observation that the addition of PAF or mc-PAF to the complete medium of cultured NB-2a neuroblastoma cells induced neurite outgrowth, which was inhibited by PAFR antagonist when cells were maintained in a serum-free medium (Herrick-Davis et al., 1991). Under this condition, PAF levels and neurite outgrowth almost paralleled. In this experimental model, PAF was essentially synthesized by the de novo pathway due, perhaps, to the lack of lysoPAF because PLA2 activity was unchanged following serum deprivation. The increase of PAF levels, C16:0-PAF but not C18:0 PAF, takes place also in PC12 cells induced to differentiate by NGF (Whitehead et al., 2007). Interestingly, PAF induced a dose-dependent secretion of NGF form cultured cortical astrocytes and this effect was further increased by TNFa (Brodie, 1995). Thus, it has been suggested that PAF may mediate the neurothophic effects exerted by astrocytes through the induction of NGF expression. PAF might be produced and released by neuronal cells or by astrocytes themselves. The latter possibility is supported by the observation that developing astrocytes posses a relatively high activity of lysoPAF-AcT which increases during development of glial cells, suggesting that the remodeling pathway is mainly responsible for PAF synthesis in these cells (Francescangeli et al., 1997a).

8.4 Gene Expression Transcription of genes encoding transcription factors are initiated by various brain physiological and pathological conditions and PAF is one of the possible mediators of these processes. In fact, in human SH-SY5Y neuroblastoma cells, PAF induces a receptor-mediated rapid and transient activation of protooncogenes c-fos and c-jun (Squinto et al., 1989). A similar effect was also observed in rat astroglial cells where the addition of PAF in the culture medium induces the expression of mRNAs for c-fos and the zinc-finger trascription factor zif/268 (Dell’Albani et al., 1993). A rapid induction of the expression of prostaglandin synthase 2 (PGHS-2) by nanomolar concentration of PAF was also reported in neural and non-neural cells (Bazan et al., 1994). This effect was synergic with that of retinoic acid and, very likely, required the interaction of PAF with the intracellular receptor. These and other observations led to hypothesize a sequence of events following the stimulation of PAF biosynthesis in neuronal cells by the remodeling pathway (Bazan and Allan, 1996). According to this hypothesis, the interaction of PAF with the BN-50730-sensitive intracellular receptor would stimulate the expression of immediate-early gene expression (i.e., Zif-268, c-fos, c-jun, etc.), the synthesis of transcription factors, and then the expression of secondary genes coding for proteins involved in long-term neuronal changes. Furthermore, the induc-

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tion of PGHS-2 would lead to the increase of prostaglandin E2 (PGE2), which would then be involved in receptor-mediated mechanism particularly relevant in inflammatory responses (Bazan and Allan, 1996).

8.5 Chemotaxis Evidence has been reported that PAF mediates the interactions between neuronal and microglial cells (Aihara et al., 2000). Indeed, microglial cells migrate toward a medium containing 10 nM PAF or the lipid extract of NMDA-stimulated primary neuronal cells and their culture medium. The chemotactic response of microglia requires the PAFR-mediated activation of Gi-dependent and MAPK-dependent pathways. These observations suggest that PAF released by neuronal cells upon NMDA receptors activation recruits microglial cells, which might contribute to cell death or removal of damaged cells.

9

Role of PAF in Brain Dysfunctions

Following the early observation of the direct neurotoxic effects of relatively high PAF concentrations (Kornecki and Ehrlich, 1988), substantial evidence has been accumulated on the participation of this lipid mediator to neuronal damage and death in various pathological conditions (Birkle et al., 1988; Bazan et al., 1991; Nogami et al., 1997; Tong et al., 2000; Bate et al., 2004c; Xu and Tao, 2004). The detrimental effects of the excessive accumulation of PAF in the brain are also supported by the neuroprotection exerted by PAF receptor antagonists (Panetta et al., 1987; Ohmori et al., 1996; Mukherjee et al., 1999; Brewer et al., 2002). Here we provide a short survey of the observations demostrating the involvement of PAF on the mechanisms causing the onset and/or the progression of some neurological diseases. These effects are generally characterized by increased levels of PAF due to upregulation of PAF biosynthetic pathways or downregulation of PAF-AH. In some cases, an altered response to PAFR activation has been also observed.

9.1

Neuroinflammation

Neuroinflammation is a common feature of many neurological diseases involving also an increased production of lipid mediators such as eicosanoids and PAF (Farooqui et al., 2007). Microglial cells and also astrocytes are the mediators of neuroinflammatory responses to insults generated by acute trauma, neurodegenerative diseases, and infections. Thus, these cells participate in the protective mechanism to limit brain damage. Both cell types are able to produce and secrete a number of immune system modulators including inflammatory cytokines such as interleukins, TNF-a, and TGFs. These factors activate a variety of mechanisms propagating the effects of neuroinflammation. Among these effects, increased expressions of PLA2 (Oka and Arita, 1991; Tong et al., 1995) and COX-2 (Phillis et al., 2006) have been reported. The increased levels of these enzymes are responsable for the augmented production of eicosanoids and PAF. In an immortalized astrocyte cell line (DITNC), it has been demonstrated that LPS, TNF-a, IL-1b, and INFg induce the expression of group II sPLA2 (Tong et al., 1995). In the same experimental model, the intracellular PAF antagonist BN50730 reduced PLA2 activity in the extracellular medium, indicating a decreased release of sPLA2 (Wang and Sun, 2000). NF-kB-mediated induction of sPLA2 gene expression and the involvement of PAF in these phenomena were then hypothesized. An interesting connection between PAF and PGE2 production by COX-2 has been also demonstrated in rat primary astrocytes (Teather and Wurtman, 2003). The stimulation of these cells with mc-PAF increased the release of PGE2, which was reduced by COX-2 inhibitors but was unaffected by inhibitors of COX-1. Even though no information is so far available on the direct effects of inflammatory cytokines on PAF synthesizing pathways in neural cells, it is hypothesized that the increased expression of sPLA2 in glial cells might lead to increased production of PAF mainly by the remodeling pathway. This lipid mediator might be

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in part released and further contribute to the activation of glial cells intiating the cascade of events leading to an increased activity of other PLA2 isoforms such as cPLA2 and the expression of COX-2. These combined effects would then cause an increased production of PGE2. On the other hand, PAF might further potentiate the expression of sPLA2-IIA through its binding to the microsomal receptor. In conclusion, PAF participates to a self-reinforcing mechanism, which, if persisted long enough, would turn the original defence function of inflammation into an offensive action against neurons.

9.2

Glutamate Neurotoxicity

The excessive activation of post-synaptic glutamate receptors leads to necrotic and apoptotic neuronal cell death (Ankarcrona et al., 1995), which is a consequence common to many neurological disorders. Several studies have indicated the existence of a link between glutamate excitotoxicity and PAF. A direct evidence was provided by Nogami and colleagues who reported that the exposure of rat neuronal primary cells to 1 mM glutamate caused cell damage and increased PAF concentration in the culture medium (Nogami et al., 1997). Cell damage induced by glutamate was reduced by pretreatment with PAFR antagonists. The addition of 10 nM PAF to the medium caused a significant increase in the number of damaged cells. The cytotoxic effect of PAF is reduced not only by PAFR antagonist BN52021 but also by the NMDA glutamate receptor antagonist (MK-801) and by the inhibition of nitric oxide (NO) synthase (Xu and Tao, 2004). Thus, the excessive stimulation of NMDA receptor should activate the formation of PAF and NO by a Ca2+-mediated mechanism, and both should further contribute to the damage of neurons induced by glutamate. The increased levels of PAF, due to stimulation of biosynthetic pathways or to an inhibition of its degradation, would exert an excessive release of glutamate, activation of NMDA receptors (Nishizawa, 2001), overloading of cytosolic Ca2+ in the post-synaptic cell, and a consequent neuronal injury (Choi, 1994). It has been recognized that mitochondria dysfunctions play an important role in neuronal cell death (Ankarcrona et al., 1995; Anderson and Sims, 1999; Cassarino and Bennett, 1999; Nicholls et al., 1999; Beal, 2000; Duchen, 2000; Fiskum, 2000; Atlante et al., 2001). The release of cytochrome c from neuronal mitochondria is a pro-apoptotic signal (Cai et al., 1998) and it is induced by glutamate (Atlante et al., 1999), by the increase of cytosolic Ca2+ concentration (Andreyev and Fiskum, 1999; Brustovetsky et al., 2002), by the accumulation of ROS (Annunziato et al., 2003) and by the excessive production of NO (Bal-Price and Brown, 2000). A possible involvement of PAF in mitochondrial damage has been investigated in isolated rat-brain mitochondria (Parker et al., 2002), in which its addition causes a dose-dependent release of cytochrome c by a mechanism apparently dependent on the activation of transition pores (PT). Since the antagonist of intracellular PAF binding site (BN50730) inhibits both PAF-induced PT and cytochrome c release, the presence of another binding site in rat-brain mitocondria, tentatively identified as the peripheral benzodiazepine receptor (PBR), has been hypothesized (Parker et al., 2002). The glutamate-induced increase of PAF formation should be the consequence of the activation of the remodeling biosynthetic pathway because the activation of Ca2+-dependent PLA2s also takes place and contributes to neuronal damage (Sun et al., 2004; Farooqui and Horrocks, 2007). Indeed, a partial protection of neuronal cells from death, induced with 50 mM NMDA, is exerted by inhibitors of cPLA2 and sPLA2 (Kowara et al., 2008), which synergistically contribute to the damage of neurons induced by glutamate (Kolko et al., 1996, 1999, 2003). The exposure of primary cortical neurons to 80 mM glutamate induced cPLA2 gene and protein expression (Kolko et al., 2003). A similar effect was also observed by exposing the cells to neurotoxic sPLA2 from Taipan snake venom (sPLA2-OS2), which binds to N-type receptors with high affinity (Lambeau and Lazdunski, 1999). Both glutamate and sPLA2-OS2 induced the release of arachidonate (AA) from membrane phospholipids, which was reduced by the pretreatment of cells with cPLA2 inhibitor AACOCF3. The release of AA and cell damage induced by glutamate or sPLA2OS2 was also significantly reduced by the PAFR antagonist BN52021 or by recombinant PAF-AH, indicating the participation of PAF in the mechanisms causing cell death by the two distinct treatments (Kolko et al., 2003). The hypothetical mechanisms of PAF-mediated neurotoxicity are summarized in > Figure 13‐5.

. Figure 13‐5 Schematic and hypothetical representation of the neurotoxic effect of PAF in response to inflammatory factors and glutamate excitotoxicity.The expression of group IIA sPLA2 is increased in microglia and astrocytes activated by inflammatory factors. This enzyme, together with Ca2+-activated cPLA2 and lysoPAF-AcT contributes to the production of PAF by the remodeling pathway. The increased PAF concentration causes a further increase of the expression of group IIA sPLA2 and COX-2 by its binding to intracellular PAF receptor (PAFIR). Both group IIA sPLA2 and PAF are released in the extracellular space and activate their presynaptic receptors causing a massive release of glutamate. The activation of NMDAR leads to the activation of postsynaptic Ca2+- dependent enzymes including sPLA2, cPLA2 and lysoPAF-AcT leading to the increase of PAF production. PAF can be released causing a further increase of its concentration in the extracellular space. The activation of PAF presynaptic receptors creates a self-reinforcing excitotoxic mechanism which may lead to neuronal death by apoptosis. The release of group IIA sPLA2 from mitochondria induced by the increased Ca2+ concentration, by PAF or NO might also accelerate the apoptotic process. Dotted lines indicate hypothetical mechanisms Metabolism and functions of platelet-activating factor (PAF) in the nervous tissue

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The toxicity of PAF is not restricted to neurons because prolonged exposure of astrocytes to 20 nM PAF significantly reduced cell viability (Hostettler et al., 2002). A similar effect was also observed in mature oligodentrocytes but at higher PAF concentration (2 mM). Pretreatment of both cell types with PAFR antagonist reduced cell death of both cell types by apoptosis because the activation of caspase-3 was also observed following the treatment with PAF (Hostettler et al., 2002).

9.3

Seizures

Intraperitoneal injection of chemoconvulsant drugs or electroconvulsions greatly enhance the production of PAF in the rat brain (Kumar et al., 1988). Electroconvulsive shock (ECS) increases the production of free fatty acids, due to the activation of brain phospholipases, which is reduced by the administration of PAFR antagonist BN52021 30 min before the treatment (Birkle et al., 1988). ECS also induce the expression of c-fos and zif-268 in rat hippocampus (Marcheselli and Bazan, 1994) and this effect is reduced by the intracellular receptor PAF antagonist BN50730. The involvement of PAF as a lipid mediator in convulsive phenomena was also reported inducing seizures by injecting kainic acid in rats. This causes long-lasting effects leading to delayed neuronal cell death in various brain areas including hippocampus through an excessive stimulation of kainate/AMPA glutamate receptors (Bennett et al., 1996). The systemic injection of kainate causes changes in the expression of extracellular PAFR detected by in situ hybridization (Bennett et al., 1998). In control rats PAFR is essentially localized in healthy neurons but after 24 h after the injection of kainate is found only in apoptotic neurons and glial cells in the proximity of damaged neurons. One week later, neuronal expression of PAFR was replaced by a marked labeling of glial cells in CA1 and CA3 pyramidal field. The expression of PAFR was reduced in surviving neurons. These observations might indicate a downregulation of PAFR expression in neurons destined to survive kainate toxicity, whereas neurons expressing the receptor are more susceptible to apoptosis.

9.4

Brain Ischemia

Following the observation that PAFR antagonist BN52021 and other ginkgolides administered to Mongolian gerbils before the induction of brain ischemia by bilateral carotid artery ligation reduced the morbidity of the animals (Spinnewyn et al., 1987), extensive investigations were carried out on the role of PAF during brain ischemia and reperfusion (reviewed by Goracci, 1990; Lindsberg et al., 1991; Yue and Feuerstein, 1994; Bazan et al., 1995; Bazan, 1998, 2006; Farooqui and Horrocks, 2006). Direct evidence for a possible involvement of PAF in brain dysfunctions due to an ischemic insult was provided by the observation that the concentration of PAF increased during ischemia due to an increased endogenous synthesis by neural cells (Kumar et al., 1988). Synthesized PAF is then released because a delayed accumulation of PAF in the hippocampal extracellular space was observed by microdialysis (Pettigrew et al., 1995). A rapid increase of PAF levels was also observed during global brain ischemia (10 min) induced by bilateral ligation of common carotid arteries of the Mongolian gerbil (Gerbil model) (Domingo et al., 1994). The levels returned to basal levels after 30 min of reperfusion. As mentioned above, the formation of PAF in the ischemic brain should be mainly the consequence of the remodeling pathway due to the rapid drop of ATP and the activation of PLA2s (Farooqui and Horrocks, 2006) as demonstrated by an immediate and massive release of free fatty acids from brain phospholipids (Bazan, 1970). The extent of the contribution of the various PLA2 isoforms and the mechanisms involved in their activation during brain ischemia are still somewhat controversial. However, evidence for the involvement of cPLA2, sPLA2, and EtnPlsPLA2 has been reported (Edgar et al., 1982; Rordorf et al., 1991; Clemens et al., 1996; Bonventre, 1997). More recently, upregulation of sPLA2-IIA in reactive astrocytes in response to ischemia–reperfusion has been demonstrated (Lin et al., 2004). Even though it has not been definitively demonstrated, the activation of the various isoforms of PLA2 should lead to increased levels of lysophospholipids in parallel with the production of FFA and particularly with an increased availability of lysoPAF for the synthesis of PAF by the remodeling pathway. The possibility that the increased PAF levels observed

Metabolism and functions of platelet-activating factor (PAF) in the nervous tissue

13

during brain ischemia might be mainly due to the activation of this pathway is also supported by the activation of lysoPAF-AcT in ischemic gerbil brain. However, during reperfusion, the de novo pathway could also contribute to maintaining relatively high PAF concentration due to the elevated specific activity of PAF-PCT and the availability of substrates (Francescangeli et al., 1996). These effects are particularly evident in the hippocampus. This hypothesis is supported by a recent study with another rat model of brain ischemia (middle cerebral artery occlusion) (Zhang et al., 2007). In these experiments, PAF concentration increased almost fivefold in the region of ischemic infarction after 90 min with respect to sham-operated animals and remained above basal levels 2 days after reperfusion. Interestingly, PAFR mRNA was reduced significantly after 6 h of reperfusion and up to 5 days, reaching the lower level after 2 days. Due to the clear evidence of the involvement of PAF in ischemia-induced brain injury, a number of studies were carried out for evaluating whether PAFR antagonists might be beneficial for reducing brain damage (Braquet et al., 1989a; Oberpichler et al., 1990; Gilboe et al., 1991; Zablocka et al., 1995; Aspey et al., 1997; Kinter et al., 1997). Recently, another approach for reducing neuronal cell death in ischemic brain has been proposed on the basis of the results obtained with transgenic mice overexpressing PAF-AH II, where both neurological deficit scores and apoptotic neurons were significantly reduced (Umemura et al., 2007). This study suggests the possibility of using recombinant PAF-AH II for future therapy of stroke.

9.5

Oxidative Stress and Aging

Multiple lines of evidence support the concept that oxidative stress contributes to neuronal cell death following brain trauma, ischemic insult, neurodegenerative diseases, and age-related decline of cognitive functions (Reynolds et al., 2007). In fact, the involvement of free radical, peroxide, NO, superoxide anion, and peroxynitrite in neuronal injury has been widely recognized (Dawson and Dawson, 1996; Chan, 2005). Because PAF levels are also elevated and contribute to brain dysfunctions caused by the onset of pathological conditions, its effects should overlap with those of oxidative stress-induced mediators. Furthermore, it might be also possible that PAF metabolism is altered by oxidative radicals. The first possibility was investigated by adding non-neurotoxic concentration of PAF to primary cortical neurons in the presence of oxidative radical donors at concentrations causing modest cellular damage (Zhu et al., 2004). In these experiments PAF potentiates H2O2-induced neurotoxicity, but had no effect on that induced by NO or superoxide donors. PAF potentiation of H2O2-induced cell damage required its interaction with plasma membrane PAFR. The concentration of PAF in rat brain decreases with age (16 weeks old) by more than 50% compared to younger animals (4–8 weeks old) (Tokumura et al., 1992). A decrease of PAF levels, measured by bioassay, was also reported in the striatum of aged rats (19 months old; 20.8  7.2 pmol/g) in comparison to younger animals (2 months old; 62.8  9.6 pmol/g) (Gimenez and Aguilar, 1998). The supplementation of the diet of aged rats with of CDP-choline (0.5 g/kg per day) caused a significant further decrease of striatum PAF levels already after 10 days of treatment. With the same experimental model, it was found that the specific activity of microsomal PAF-PCT did not change with age, whereas that of PAF-AH was significantly increased (Gimenez and Aguilar, 2001). In addition, it was also reported that the specific activity of PAF-PCT in the striatum was reduced by 55% after only 8 days of treatment with CDP-choline supplemented diet, whereas the same treatment did not affect the specific activity of the phosphocholine transferase-synthesizing phosphatidylcholine. PAF-AH activities, assayed in the same samples, were not significantly reduced. It was concluded that the decrease of PAF-PCT activity was responsible for the reduced levels of PAF in the striatum of aged animals and that diet supplemented with CDP-choline might be beneficial for the reduction of PAF-mediated neuronal damage. The decreased levels of PAF observed in the healthy aged brain might correlate with a moderate loss of PAF-mediated fuctions, such as LTP and memory formation, which appears to be mainly due to an increased PAF-AH activity more than a reduced biosynthetic efficiency by the de novo pathway. On the other hand, the effects attributed to the administration of CDP-choline per se are questionable and difficult to explain. First of all, it is very unlikely that CDP-choline might reach unchanged brain regions and consequently modulate PAF metabolism. Furthermore, the increased PAF levels observed in brain injury and neurodegerative diseases are

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mainly due to the activation of the remodeling pathway where CDP-choline might be beneficial by modulating the activities of phospholipases A2 (Adibhatla et al., 2006).

9.6

Hypoxia

The role of lipid mediators in relation to hypoxia-induced brain damage has been extensively reviewed (Bazan et al., 2002b). Normoxia, following brief periods of hypoxia, generates reactive oxygen species (ROS) and causes the induction of inflammatory genes (Bazan and Lukiw, 2002). Evidence for the potential involvement of PAF in hypoxia-induced damage, and particularly in rat hippocampal cultured neurons, has been reported (Ohmori et al., 1996). The correlation of PAF with brain dysfunctions was also demonstrated in a rodent model of intermittent hypoxia, where an increase of oxidative stress, inflammation, and neurodegenerative changes could be observed (Row et al., 2002). In comparison with their wild-type littermates, PAFR –/– mice were protected from spatial learning deficits when subjected to intermittent hypoxia. In addition, this treatment did not increase the expression of inducible NO synthase and reduced the production of PGE2 (Row et al., 2004). It can be concluded that the intermittent hypoxia might lead to neurological dysfunctions through PAFRmediated mechanisms, which might contribute to memory deficits due to loss of regional gray matter observed in obstructive sleep apnea (Macey et al., 2002).

9.7

Alzheimer’s Disease

Pathophysiological events very likely involved in the onset and development of Alzheimer’s disease (AD) and alterations of brain lipid metabolism, observed in patients or in animal models, have been extensively described (Shimohama et al., 1993; Farooqui et al., 1995a; Lehtonen et al., 1996; Stephenson et al., 1996; Kanfer et al., 1998; Ross et al., 1998; Blass, 2000; Talbot et al., 2000; Han et al., 2001; Bazan et al., 2002a; Chen et al., 2002; Chalimoniuk et al., 2006; Moses et al., 2006; Butterfield and Sultana, 2007; Crouch et al., 2007; Forlenza et al., 2007; Mukherjee et al., 2007; Sun et al., 2007). Specific alterations of the metabolism of PAF in AD brains are not available so far. However, in addition to the evidence of its contribution to neuronal cell death common to neurodegenerative diseases, some observations lead to the hypothesis that PAF might participate in mechanisms characterizing AD. One of these is the activation of sPLA2 by Ab peptides in vitro (Lehtonen et al., 1996). The treatment of primary cortical neurons with Ab also increases the release of arachidonic acid due very likely to the activation of cPLA2 mediated by ERK 1/2 (Shelat et al., 2008). Thus, an increase of Ab-mediated PLA2 activity might produce lysoPAF and consequently stimulate PAF synthesis by the remodeling pathway. Furthermore, neurons pretreated with PLA2 inhibitors or PAFR antagonists were more resistant to the neurotoxic effect of Ab peptide (Bate et al., 2004b, c).

9.8

Parkinson’s Disease

Parkinson’s disease (PD) is characterized by a gradual loss of dopaminergic neurons in the substantia nigra where an intense lipid peroxidation and superoxide dismutase activity has been reported. At present, no specific information is available on the role of PAF in the pathogenesis of PD, which, however, might contribute to the progressive degeneration of dopaminergic neurons following oxidative damage (Thomas and Beal, 2007). Several recent findings indicate that inflammation may contribute to the pathogenesis of PD. A study on genetic variants of genes coding for components involved in immune reactions in the brain and potentially influencing the risk of developing PD was carried in patients affected by this disease (Hakansson et al., 2005). No correlation was found between the variant of platelet-activating factor acetylhydrolase (PAF-AH; Val379Ala) and PD.

Metabolism and functions of platelet-activating factor (PAF) in the nervous tissue

9.9

13

Miller–Dieker Lissencephaly

PAF has been implicated in the human neuronal migration disorder, the so called Miller–Dieker lissencephaly, which is due to the haploinsufficiency of the gene LIS-1 encoding a 45-kDa subunit of PAF acetyl hydrolase (PAF-AH 1b) (Hattori et al., 1994a) causing a defect of PAF catabolism. PAF affects neuronal migration by receptor-dependent and receptor-independent mechanisms (Tokuoka et al., 2003). This disease is responsible for severe defects of brain development in children due to disruption of the specific distribution of neurons within the cerebral cortex, which causes mental retardation, epilepsy, and usually early death. This condition arises from an inadequate migration of neuronal progenitors to their cortical destination, resulting in a low number of neurons in the cortex and in a smooth rather than folded surface of the brain. LIS1/b interacts with the catalytic subunit of PAF-AH (I) indicating that the b subunit functions as a subunit of intracellular PAF-AH and also exhibits important cellular functions such as induction of nuclear movement and control of microtubule organization. Moreover, overexpression of the PAF-AH (I) catalytic subunits has been shown to induce centrosomal amplification and microtubule disorganization by disturbing intracellular localization of LIS1 (Yamaguchi et al., 2007). The cellular function of the catalytic subunits is still under investigation. Several results support the hypothesis that the PAF-AH (I) catalytic subunits also play a role in brain development and diseases. The a1 expression was found to be restricted to actively migrating neurons and alteration of catalytic subunits from the a1/a2 heterodimer to the a2/a2 homodimer occurring in these cells during brain development, suggesting that these catalytic subunits play a role in neuronal migration (Manya et al., 1998).

9.10 Multiple Sclerosis PAF levels were assayed in the cerebral spinal fluid (CSF) and plasma from patients with relapsing/remitting or secondary progressive multiple sclerosis (MS), and they were significantly more elevated with respect to controls (Callea et al., 1999). In both plasma and CSF, PAF concentration was higher in relapsing/remitting than in secondary progressive MS and correlated with markers of blood–brain barrier (BBB) injury but not with the number of white matter lesions or with the expanded disability status score. It was suggested that PAF might be a marker of BBB injury in the early stages of MS, rather than a marker of its progression and severity. The role of PAF in MS has been investigated in experimental allergic encephalomyelitis (EAE), which is considered an animal model of MS, and a correlation was found between levels and expression of PAFR mRNA in spinal cord with EAE symptoms (Kihara et al., 2005). Furthermore, PAFR-KO mice were better protected against the onset or the severity of symptoms. Because cPLA2 has been detected in EAE lesions (Kalyvas and David, 2004) and cPLA2 ( / ) mice are more resistant to EAE (Marusic et al., 2005), the remodeling pathway for PAF production should be mainly responsible for it production.

9.11 HIV Infection PAF levels are elevated in CSF of patients affected by HIV-1-associated dementia (HIV-D) (Gelbard et al., 1994). Mechanisms involved in neurodegeneration induced by HIV infection and AIDS have been extensively reviewed, and a model for neuronal apoptosis has been proposed (Kaul et al., 2005). According to this model, a central role in initiating neuronal injury and apoptosis is played by the overstimulation of NMDAR, similar to other neurodegenerative diseases. In this case, HIV-infected or immune-stimulated macrophages/microglia release neurotoxins, causing an impaired clearance (or release) of glutamate, which, under normal conditions, would have been taken up by astrocytes. However, when activated by antigenic stimuli, HIV-1-infected monocytes release PAF in addition to TNF-a (Nottet et al., 1995), and a link between PAFR activation and glutamate neurotoxicity has been proposed as discussed above. An interesting hypothesis for neuronal apoptosis induced by HIV infection has been proposed by Perry et al. (1998), who suggested that TNF-a neurotoxicity was mediated by ceramide and required the activation of PAFR. Indeed, ceramide levels were elevated in a neuronal cell line exposed to TNF-a, and

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the PAFR antagonist WEB 2086 abrogated both TNF-a and ceramide-induced neurotoxicity. Furthermore, co-incubation of human neuronal cells with PAF-AH abolished the increase of cell death induced by the medium from activated HIV-1-infected macrophages. A randomized, double-blind, placebo-controlled clinical trial was conducted to investigate whether treatment of HIV-infected patients with PAFR antagonist (Lexipafant) might be beneficial for improving their cognitive dysfunctions. The outcome from this investigation indicated a trend toward improvement in neurophycological performance in patients treated with the drug (Schifitto et al., 1999).

9.12 Prion Disease A study for determining the metabolic pathways responsible the formation of b-sheet-rich disease-related isoform (PrPSc) from host-encoded cellular prion protein (PrPC) was carried out using scrapie-infected neuroblastoma cell lines as a cellular model of the disease (Bate et al., 2004a). The treatment of infected cells with inhibitors of various PLA2 isoforms or PAF antagonists caused a marked reduction of the formation of the protease-resistant core (PrPres) of PrPSc. Furthermore, the exposure to PAF or mc-PAF increased the formation of PrPres. The effects of various PAF antagonists were also tested on the cellular levels of PrPC necessary for the formation of PrPres. When N2a cells were grown in the presence of PAF antagonists, a dose-dependent reduction of PrPC was observed. Conversely, 2 mM PAF or mc-PAF increased PrPC content in the same cell type. These observations indicate that drugs counteracting PAF production by the remodeling pathway or PAF antagonist, together with drugs targeted at the structure of PrPSc itself, might have beneficial effects in preventing or limiting neurodegeneration in prion disease.

10

Conclusions

The presence of PAF in mammalian brain and the demonstration that this lipid mediator participates in the mechanisms involved in important physiological functions have led to the investigation of the metabolic pathways for its synthesis and degradation. The overall reactions of these pathways are similar to those reported for other organs and cell types. However, it has been also well documented that PAF levels increase both in acute and chronic pathological conditions and that increased PAF concentrations cause neuronal cell death, mainly by apoptosis. Thus, it appears to be of primary importance to unravel the mechanisms involved in the regulation of the levels of this lipid mediator in the brain tissue. This task is far from complete because of the complexity of the organ, which is greatly heterogeneous for anatomy, morphology, and functions. Neural cells posses both the substrates and the enzymes of the pathways for PAF biosynthesis and it has been proposed that the de novo pathway might be mainly responsible for providing the levels of PAF necessary for physiological mechanisms. On the basis of this dichotomic view of the biosynthetic pathways, the remodeling pathway had to be mainly responsible for the increased levels of PAF under pathological conditions. This concept is also supported by the numerous observations that indicate a correlation between increased PLA2 and the onset of neurological dysfunctions. However, there is also substantial evidence for the production of PAF by the remodeling pathway in physiological events such as glutamatergic neurotransmission during the formation of LTP and its maintenance. Levels of PAF depends also on the activity of PAF-Ahs, and, consequently, the complete panel of information necessary for understanding the mechanisms regulating PAF levels in different situations should include (1) how the expressions of key enzymes of PAF metabolism are regulated and (2) how their activities change in response to specific stimuli. Both aspects are still to be defined. Cellular responses to PAF are mediated by plasma membrane (PAFR) and intracellular receptors. While PAFR has been extensively studied and characterized, little information is available on the molecular properties of intracellular PAF binding sites in neural cells even though they appear to mediate PAF-induced nuclear responses. Thus, the identification at molecular levels of intracellular PAF binding sites and the elucidation of the following steps leading to the expression of genes appear to be of primary importance for further understanding how PAF specifically regulates or, depending on its levels, interferes with normal cellular functions.

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The increased information on PAF metabolism and on the mechanisms involved in cellular responses will allow developing new molecules that will be able to modulate its levels or antagonize its effects. These molecules will then provide new tools for maintaining or improving PAF-mediated physiological functions as learning and memory formation or, in pathological conditions, for reducing its neurotoxic effects.

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Lipid Anchors to Proteins

N. N. Nalivaeva . A. J. Turner

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 354

2 2.1 2.1.1 2.1.2 2.2 2.2.1 2.2.2 2.3 2.3.1 2.3.2 2.3.3 2.3.4 2.3.5

Types of Lipid Modifications of Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 354 Protein Acylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 354 Protein Myristoylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 355 Protein Palmitoylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 357 Protein Isoprenylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 358 Farnesyltransferase and Geranylgeranyltransferase I . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359 Rab Geranylgeranyl Transferase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359 Glycosyl-Phosphatidylinositol (GPI) Anchors of Membrane Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . 361 Shedding and Roles of GPI-Anchored Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 362 Prion Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 Neural GPI-Anchored Cell Adhesion Molecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364 GPI-Anchoring of AChE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364 Other GPI-Anchored Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365

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Functions of Lipid Modifications of Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 366

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_14, # Springer ScienceþBusiness Media, LLC 2009

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Lipid anchors to proteins

Abstract: A variety of lipid anchor mechanisms are used to target and stabilise the membrane association of proteins, many of which play important roles in synaptic development, function and plasticity. These lipid anchors include long-chain acyl or prenyl groups, glycosyl-phosphatidylinositol (GPI) and cholesterol. The lipid association with proteins can be either reversible or irreversible and an individual protein may have more than one lipid anchor attached. This chapter summarises some of the main mechanisms of lipid protein anchoring, including myristoylation, palmitoylation, isoprenylation and GPI addition and the enzymes involved in the covalent addition of lipid moieties to proteins. Lipid anchoring can influence protein targeting, especially to membrane lipid rafts, which can have pathological consequences, e.g. in Alzheimer’s disease. Specific examples of lipid-anchored proteins of neurochemical significance will be described in detail, including the prion protein, neural cell adhesion molecules and acetylcholinesterase. List of Abbreviations: AChE, acetylcholinesterase; ADAM, A disintegrin and metalloproteinase; AMPA, α-amino-3-hydroxyl-5-methyl-4-isoxazole-propionate; APP, amyloid precursor protein; BACE, β-site APP cleaving enzyme (β-secretase); EGFR, epidermal growth factor receptor; GAP-43, Growth Associated Protein 43 (neuromodulin); GPI, glycosyl-phosphatidylinositol; HMG, hydroxymethylglutaryl; MARCKS, myristoylated alanine-rich C-kinase substrate; MBOAT, membrane bound O-acyltransferase; N-CAM, neural cell adhesion molecule; PI-PLC, phosphatidylinositol specific phospholipase C; PLA2, phospholipase A2; PPT, palmitoyl-protein thioesterase; PRiMA, proline-rich membrane anchor; PrP, prion protein; PSD, post-synaptic density; REP, rab escort protein; RGGT, rab geranylgeranyl transferase; Shh, sonic hedgehog; SNAP, synaptosome-associated protein; SNARE, soluble NSF Attachment protein receptors

1

Introduction

Post-translational modification is a critical process that regulates the structure, properties, cellular location, and functions of numerous proteins. It can occur at various stages of protein metabolism and can be transient or permanent. The modifications range from amino acid changes through to the addition of macromolecules: carbohydrate, protein or lipid. Lipid modifications of proteins include covalent attachment of long-chain acyl groups, particularly myristate or palmitate, modification by long-chain prenyl groups, addition of cholesterol and C-terminal anchorage of proteins via glycosyl-phosphatidylinositol (GPI) moieties. Such modifications alter dramatically the hydrophobic character of proteins, affecting their structure and folding characteristics, facilitate their interactions with cell membranes and hence can play a role in cellular signaling events, reversible cell targeting of proteins and thus cell function, growth and differentiation. For example, key growth factor proteins, G proteins, cell adhesion molecules such as neural cell adhesion molecule (N-CAM), cell surface receptors, and cytoskeletal proteins can be so modified. > Table 14‐1 summarizes the major lipid modifications and provides a number of specific examples particularly of neurochemical significance, and > Figure 14‐1 illustrates their chemical diversity. More detailed discussion of relevant processes and proteins so modified are provided in the succeeding sections. Over the last 10 years many of the specific signals that direct lipid modification have been revealed and the enzymology of these processes has been unraveled. Such enzymes provide potential targets for novel therapeutic agents in a variety of different conditions.

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Types of Lipid Modifications of Proteins

2.1 Protein Acylation The process of protein acylation has been known for some three decades and was originally described for the bacterial proteolipids but is now known to be widespread in both prokaryotic and eukaryotic organisms. The most common fatty acids found attached covalently to eukaryotic proteins are myristate (C14:0) and palmitate (C16:0), although their underlying biochemistry are quite distinct. Other fatty acids are occasionally found, for example, both palmitate and stearate (C18:0) are attached in S-acyl linkage to the protein kinase C substrate GAP-43 (neuromodulin) (Liang et al., 2002). The attachment of such lipids often appears to be required for optimal activity of proteins and can facilitate membrane interaction (e.g., for

Lipid anchors to proteins

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. Table 14‐1 Some examples of lipid modifications of proteins Palmitoylation Structural proteins: Ankyrin

Fibronectin GAP-43

Postsynaptic density-95 protein (PSD-95) SNAP and SNARE proteins

Synaptotagmin Vinculin Signal transduction proteins: G-protein-coupled receptors (some) GABAA receptors Na+ channel, a-subunit NCAM140 Nicotinic receptors: Potassium channel Kv1.1

Myristoylation Signal transduction proteins: Cyclic AMP-dependent protein kinase (catalytic subunit) Calcineurin G proteins (a subunits of G0, Gi)

Guanylate cyclase-activating proteins (GCAPs)

Isoprenylation G protein g subunit Lamin B and prelamin A Ras and some ras-related proteins Rab1 and Rab2

GPI-anchored proteins Enzymes: Alkaline phosphatase

Acetylcholinesterase (erythrocytes) 50 -nucleotidase

Aminopeptidase P Carboxypeptidase M

Membrane dipeptidase Gravin (A-kinase anchoring protein) MARCKS (myristoyl alaninerich C-kinase substrate) Src family kinases TRAM (TRIF-related adaptor molecule) Other enzymes: Endothelial nitric oxide synthase

ras proteins Rhodopsin Src family protein tyrosine kinases Enzymes: Active g-secretase complex BACE-1 Ca2+-ATP-ase (sarcoplasmic reticulum) GAD65

Mammalian antigens: Thy-1 Qa (mouse lymphocytes) Ly-6 (mouse lymphocytes) Carcinoembryonic antigen Cell adhesion molecules: Fasciclin I (Drosophila) Fibronectin receptors (subset) LFA-3 (lymphocytes) Neural cell adhesion molecule Other: Prion protein

GAP-43, see Denny, 2006). Myristic and palmitic acids appear to label different pools of cellular proteins and occur by very different pathways: myristoylation occurs as an irreversible co-translational event whereas palmitoylation is a post-translational, reversible process.

2.1.1 Protein Myristoylation The addition of myristic acid to proteins requires the N-terminal sequence Met1-Gly2- and the myristate is attached to the glycine residue via an amide bond during translation, after removal of the initiating Met

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Lipid anchors to proteins

. Figure 14‐1 Schematic representation of lipid-modified proteins. (a), S-palmitoylated (e.g., nicastrin and Aph-1 subunits of g-secretase complex; Cheng et al., 2009); (b) – N-palmitoylated (e.g., human growth hormone; Sachon et al., 2007); (c), farnesylated (e.g., cPLA2g; Hirabayashi et al., 2004); (d), O-acylated (e.g., Wnt-3a; Takada et al., 2006); (e), N-myristoylated (e.g., calcineurin, Selvakumar et al., 2007); (f), geranylgeranylated (e.g., ras proteins; Zhang and Casey, 1996); (g), GPI-anchored (e.g., prion protein; Baldwin, 2005; AChE in erythrocytes or in Drosophila nervous system, Incardona and Rosenberry, 1996); (h), cholesterol-modified (e.g., sonic hedgehog protein, Huang et al., 2007)

residue by methionine aminopeptidase. This modification was first detected on the catalytic subunit of cyclic AMP-dependent protein kinase (Carr et al., 1982) and, shortly afterwards, on the brain calcium-dependent phosphoprotein phosphatase, calcineurin (protein phosphatase 2B) (Aitken et al., 1982). Since myristate is a relatively minor eukaryotic fatty acid, its structure must presumably be critical to the functioning of the myristoylated protein. Not all proteins with an N-terminal glycine become myristoylated since there is a consensus sequence at the N-terminus for myristoylation consisting of (M1)G2XXXS6/T6. Structural studies of N-myristoyl transferase, which is a therapeutic target for a number of disorders, revealed a novel catalytic mechanism and the dimensions of the myristoyl-CoA binding site limit the size of the fatty acid which can be bound, hence favouring the relatively short chain and less abundant, myristate (Bhatnagar et al., 1998, 1999). It is common for signaling proteins to be N-myristoylated, for example, many protein kinases and phosphatases, various G proteins and Ca2+-binding proteins. Myristoylation is often required for membrane binding of a protein, although myristoylation alone does not secure membrane association and a second modification is normally required to stabilize membrane association. This can either be the addition of palmitate, protein-protein interactions, or the presence of a cluster of positively charged amino acids that interact with acidic phospholipid headgroups in the membrane. Although the covalent attachment of myristate to a protein is irreversible, its membrane binding can be reversed. Phosphorylation of the myristoylated protein can destabilize the membrane interaction releasing the protein as a soluble form: the so-called ‘‘myristoyl switch’’ for reversible membrane association of a protein. A well-characterized example is the myristoylated MARCKS (myristoyl alanine-rich C-kinase substrate) protein (Aderem, 1992), which, when phosphorylated by protein kinase C in its basic region, causes its release into the cytosol. Many neuronal calcium sensor proteins are membrane targeted via myristoylation (Burgoyne, 2004). Other

Lipid anchors to proteins

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neural protein substrates for myristoylation include src family kinases, the catalytic subunit of cAMPdependent protein kinase, the A-kinase anchoring protein gravin (Wang et al., 2006), and the phosphoprotein phosphatase, calcineurin. Myristoyl-transferases appear to be up-regulated in epilepsy and have been suggested as potential therapeutic targets in the disorder (Lakshmikuttyamma et al., 2008).

2.1.2 Protein Palmitoylation The reversible S-acylation of proteins by palmitate occurs on cysteine residues and is an important component of signal transduction pathways. Other functions of palmitoylation include protein trafficking and vesicle fusion as well as organelle inheritance (Smotrys and Linder, 2004). S-palmitoylation can occur at a variety of sites in a protein, near the N-or C-terminus, or close to transmembrane domains. The incorporation of [3H]-palmitic acid into proteins was first shown in tissue culture thirty years ago by Schmidt and Schlesinger (1979) using viral membrane glycoproteins although subsequently normal cellular proteins were also found to be modified, for example, the transferrin receptor. Metabolic labeling studies have revealed that many of the palmitoylated proteins are associated with the inner face of the plasma membrane from which they are not readily released, indicating the integral nature of their hydrophobic association with the membrane. Some of the palmitoylated proteins exist as transmembrane glycoproteins, for example, rhodopsin and some other G-protein linked receptors. > Table 14‐1 indicates the different classes of proteins that can be palmitoylated. In addition to transmembrane proteins, many cytosolic proteins can be palmitoylated. Often these may contain an additional lipid modification, e.g., an N-terminal myristate or a C-terminal prenyl group. The difficulties in purifying and assaying palmitoyltransferases meant that they were first identified only in 2002 and this required the application of yeast genetics (Lobo et al., 2002; Roth et al., 2002). Although the two palmitoyltransferases cloned differed in substrate specificity, they shared features common to all this family of enzymes, multiple transmembrane regions and a consensus motif Asp-His-His-Cys in a cysteine-rich domain (generally now referred to as a DHHC-CRD motif). During the course of reaction the cysteine residue in the DHHC motif is autopalmitoylated which probably serves as an acyl-enzyme intermediate in the reaction mechanism (Lobo et al., 2002; Roth et al., 2002). Palmitoylation serves key roles in the brain and substrates include receptors and ion channels, SNARE and SNAP proteins, the postsynaptic density protein PSD-95, GAP-43 and glutamic acid decarboxylase (GAD65). The clustering of receptors at synapses involves palmitoylation and treatment of cells with the palmitoylation inhibitor 2-bromopalmitate causes diffusion of AMPA receptors and PSD-95 from the clusters (El-Husseini et al., 2002). Glutamate and GABA-A receptor subunits are also subject to palmitoylation and the enzyme responsible for palmitoylation of the GABA-A g2 subunit has been identified as a Golgi-specific DHHC zinc finger protein (GODZ) (Keller et al., 2004). The roles of lipid and other modifications of GABA-A receptors in their stabilization, trafficking, and internalization are reviewed in (Chen and Olsen, 2007). Another neuronal signaling protein interacting with, and targeted by, a palmitoyl-transferase is nitric oxide synthase (Saitoh et al., 2004). However, unlike endothelial NOS (Robinson and Michel, 1995), direct palmitoylation of n-NOS has not been demonstrated. Quite separate from the process of S-palmitoylation is that of N-palmitoylation in which palmitate is covalently and irreversibly linked to the amide group of an N-terminal cysteine residue, a modification that is found in secreted signaling molecules such as ‘‘sonic hedgehog’’ (Shh) and Wnt proteins as well as the EGFR ligand Spitz (Miura and Treisman, 2006). Lipid modification is one of the mechanisms that have evolved to restrict the range and increase the activity of diffusible, secreted morphogen ligands. Shh, which also carries a C-terminal cholesterol modification, plays a critical role in developmental patterning of the human brain. The Shh palmitoylating enzyme (Hhat) has been isolated and characterized as a transmembrane protein O-acyltransferase (MBOAT) related to the enzymes that transfer fatty acids and other lipids onto hydroxyl groups of membraneembedded lipids (Buglino and Resh, 2008). Only two other MBOAT proteins, Porcupine (Porc) and GOAT, have been implicated in the transfer of fatty acids to proteins. Porc is probably the Wnt O-acyltransferase and GOAT catalyses the attachment of octanoate to proghrelin, the appetite-stimulating hormone precursor. Palmitoylation may provide a mechanism, like GPI-anchoring, for targeting proteins to lipid microdomains (‘‘lipid rafts’’) in cell membranes, where they can regulate signaling events (Simons and Toomre,

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2000; Huang and El-Husseini, 2005). However, palmitoylation does not necessarily target proteins to such domains. All isoforms of the membrane metalloproteinase and amyloid-degrading enzyme, endothelin converting enzyme, contain a conserved cysteine residue close to the membrane spanning region which is palmitoylated, yet the enzyme is excluded from such microdomains (Schweizer et al., 1999). In this case, palmitoylation may stabilize against protein turnover and/or delivery to lysosomes. Raft localization probably requires a minimum of dual acylation (e.g., palmitate plus myristate). A similar situation arises with the phagocyte protein tyrosine kinase, p59Hck, where lack of palmitoylation redirects the protein from the plasma membrane to lysosomes (Carreno et al., 2000). The biochemistry and functions of protein acylation have recently been reviewed by Planey and Zacharias (2009). Studies are currently underway to define the full ‘‘palmitoyl proteome’’ of cells. This was first studied in yeast (Roth et al., 2006) but recently the rat neural palmitoyl proteome has been characterized and, in addition to the 68 known such palmitoylproteins, has identified over 200 other palmitoyl-protein candidates (Kang et al., 2008). The dynamics of palmitoylation, and the substrates for this process, strongly suggest that palmitoylation participates in synaptic activity, morphology and function. More is known in structural and functional terms of the deacylating enzymes, the palmitoyl-protein thioesterases (PPT). The structure of PPT-1, a major lysosomal enzyme, has recently been reported (Bellizzi et al., 2000) and deficiency of this enzyme results in a severe neurodegenerative condition, infantile neuronal ceroid lipofuscinosis, emphasizing the critical importance of reversible protein acylation. PPT-1 contains a serine lipase consensus sequence at its active site, although it is insensitive to the serine reagent, phenylmethylsulphonyl fluoride (Das et al., 2000).

2.2 Protein Isoprenylation Isoprenoids are ubiquitous in nature and have very diverse roles. The basic building block is the 5-carbon compound, isopentenyl pyrophosphate formed from mevalonate, the product of HMG CoA reductase. By combining precursors with 5 carbons, the pathway subsequently produces geranyl pyrophosphate (10 carbons), farnesyl pyrophosphate (15 carbons) and geranylgeranyl pyrophosphate (20 carbons). Isopentenyl pyrophosphate is the precursor to squalene in cholesterol biosynthesis. This means that statins, which inhibit HMG CoA reductase, inhibit the production of both cholesterol and isoprenoids and the physiological effects of statins may not necessarily relate to reduction in cholesterol levels. For example, the ability of statins to reduce the production of the amyloid b-peptide is due, at least in part, to inhibition of protein isoprenylation (Ostrowski et al., 2007). The condensation of two isopentenyl groups produces the ten-carbon geranyl-pyrophosphate and successive addition of isopentenyl groups then generates the intermediates farnesyl pyrophosphate and geranylgeranyl pyrophosphate. It is these latter isoprene units that act as protein modifying groups. The first prenyl-containing peptide identified was that of a yeast-mating factor, an 11-amino acid peptide which was shown to contain a farnesyl group attached to a C-terminal cysteine residue (Kamiya et al., 1978). Other fungal mating factors were also shown to be modified on cysteine residues with farnesyl groups and the cysteine itself was further carboxymethylated. Both these modifications were essential for mating-factor activity. Sequencing of the gene encoding Saccharomyces cerevisiae mating factors revealed the C-terminal sequence Cys-Val-Ile-Ala. This sequence, or rather the more general ‘‘CAAX box’’ (where C = Cysteine, A = aliphatic residue and X = any other amino acid), serves as the signal for isoprenylation which, in turn, signals the sequential carboxypeptidase removal of the X,A,A residues followed by carboxymethylation of the Cys. The structure of a farnesylated cysteine residue is shown in > Figure 14‐1. Studies of post-translational processing of ras-related proteins established unequivocally the role of the CAAX box in directing isoprenylation (Hancock et al., 1989) and which confirmed that ras proteins and fungal mating factors were processed by a common pathway. Inhibitors of HMG-CoA reductase, and hence of mevalonate synthesis, block the farnesylation of ras proteins and subsequent processing steps. More than 100 cellular proteins have been observed as prenylated. Of significance are the proteins of the nuclear lamina: lamin B, has a C-terminal CAAX box and is farnesylated which may play a role in association of the protein with the nuclear envelope (Farnsworth et al., 1989). The lamin A precursor (prelamin A) also contains a CAAX box and is transiently farnesylated but uniquely the mature protein is truncated at the C-terminus in a

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farnesylation-dependent proteolytic process which removes the 15 C-terminal amino acids, including the farnesyl group. In some rare, inherited forms of premature ageing, progeria, a mutation in the prelamin A gene produces an alternatively spliced form of prelamin A (progerin) which lacks 50 amino acids near the Cterminus including the proteolytic site and hence farnesylated progerin accumulates (De Sandre-Giovannoli et al., 2003). Another isoprenyl group, the 20-carbon isoprenoid, geranylgeranyl, can also modify proteins and is, in fact, more common than farnesylation. Examples of geranylgeranylated proteins include the g subunits of G proteins, some of which contain the C-terminal sequence, Cys-Ala-Ile-Leu (CAIL) (Yamane et al., 1990) and the ras related rab subfamily of G proteins which are involved in the mammalian secretory pathways (Khosravi-Far et al., 1991). The rab proteins do not have a CAAX box but have a pair of cysteines at or near the C-terminus (XXCC, XCXC, or CCXX). The observation that tetrapeptides corresponding to the C-termini of farnesylated proteins inhibit protein farnesyltransferase in vitro, whereas CAIL does not, first suggested that the two isoprenes (farnesyl and geranylgeranyl) may be transferred by different enzymes (Reiss et al., 1991). There are now known to be three mammalian protein prenyltransferases: farnesyltransferase and geranylgeranyltransferase type I which transfer their respective isoprenoids to the cysteine of a C-terminal CAAX sequence (Lane and Beese, 2006), and geranylgeranyltransferase type II that exclusively acts on rab family proteins (Leung et al., 2006).

2.2.1 Farnesyltransferase and Geranylgeranyltransferase I Protein farnesyltransferase can be assayed by its ability to transfer [3H]-farnesyl pyrophosphate to the protein substrate p21Ha-ras and was originally isolated from rat brain (Reiss et al., 1990). The purified protein exists as a heterodimer. The a subunit appears to be common to both farnesyltransferase and geranylgeranyltransferase since antibodies directed against the a subunit of farnesyltransferase are able to immunoprecipitate both activities (Seabra et al., 1991). The b subunit confers specificity for the protein acceptor and is different for the two enzymes, although homologous (25% sequence identity). These enzymes recognise the CAAX box at the C-terminus of the target protein. The two transferases exhibit different specificities, farnesyltransferase preferring Met, Ser, Gln, or Ala at the C-terminus and geranylgeranyltransferase type I preferring Leu or Phe. Additional features that may contribute to substrate specificity have been identified in a refinement of protein prenylation motifs which are available as the prenylation prediction suite, PrePS (http://mendel.imp.univie.ac.at/sat/PrePS) (Maurer-Stroh and Eisenhaber, 2005). Both proteins are essential and their deletion is usually lethal. The structures and mechanisms of these enzymes, which are both zinc metalloenzymes, have been elucidated (Long et al., 2002; Taylor et al., 2003). In general, prenylated proteins play pivotal roles in signal transduction and intracellular trafficking pathways and hence inhibitors of prenylation could have applications in a number of diseases. Many prenyltransferase inhibitors have been discovered or developed either as anticancer drugs or in the treatment of parasitic, fungal, or viral infections. Farnesyl transferase inhibitors have shown some promise in treating a mouse model of progeria.

2.2.2 Rab Geranylgeranyl Transferase Rab geranylgeranyltransferase, or geranylgeranyl transferase type II (RGGT), transfers (usually) two geranylgeranyl groups to the cysteine(s) at the C-terminus of Rab proteins (ras genes from rat brain). RGGT was first purified from rat brain cytosol as a protein complex comprising the catalytic component, which consists of two subunits like farnesyltransferase, and a Rab binding protein, the ‘‘Rab escort protein’’, REP. The Rab proteins, of which there are more than 60, represent the largest group of the ras superfamily of small G proteins and regulate organelle biogenesis and vesicle transport. The C-terminus of Rab proteins varies in length and sequence and is referred to as hypervariable. Thus rab proteins do not have a consensus sequence, such as the CAAX box, which the Rab geranylgeranyl transferase can recognise. Instead rab proteins are bound by the escort protein REP over a more conserved region of the Rab protein and then presented to RGGT (Leung et al., 2006). Once Rab proteins are prenylated, the lipid anchor(s) ensures that Rabs are no longer soluble. REP therefore plays an important role in binding and solubilizing the geranylgeranyl

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groups and delivers the Rab protein to the relevant cell membrane. The crystal structure of RGGT has been solved to 2.0 A˚ resolution and has helped to elucidate the reaction mechanism (Zhang et al., 2000). Prenyl proteins have a relatively short half-life (approx. 20 h) and their turnover requires removal of the prenylcysteine moiety. This is effected by a prenylcysteine lyase, a flavin adenine dinucleotide-dependent thioether oxidase with a completely novel mechanism (Tschantz et al., 2001). It is located in lysosomes like the palmitoylprotein thioesterase that turns over palmitoylated proteins (Tschantz et al., 1999). The ras small G proteins, H-, N-, and K-ras 4A/B, are lipid-anchored, peripheral membrane guanine nucleotide-binding proteins. The dual modification of ras small GTPase oncogene proteins provides a mechanism for their membrane association since their amino acid sequences do not indicate any hydrophobic membrane-spanning domains. Palmitoylation of these proteins appears to be dependent on prior isoprenylation and requires the presence of a second cysteine residue close to the CAAX box. All ras proteins are polyisoprenylated but only some are palmitoylated. The dynamic trafficking of ras proteins, and other neural proteins such as GAD65, to and from the plasma membrane (ras proteins) or synaptic vesicles (GAD65) is intimately associated with their lipidation status (> Figure 14‐2). The ras proteins appear to be laterally segregated into nonoverlapping, dynamic domains of the plasma membrane referred to as nanoclusters. This lateral segregation is important to specify ras interactions with membrane-associated proteins, effectors and scaffolding proteins and is critical for ras signal transduction (Abankwa et al., 2007). The role of lipid modifications in the membrane targeting and trafficking of ras proteins has been reviewed in Wright and Philips (2006). . Figure 14‐2 Schematic representation of the anterograde and retrograde intracellular transport of dual lipidated proteins such as ras proteins between cytosol, ER/Golgi and plasma membrane. The irreversible isoprenylation is coupled with reversible cycles of S-palmitoylation and depalmitoylation to regulate trafficking

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2.3 Glycosyl-Phosphatidylinositol (GPI) Anchors of Membrane Proteins Another highly specialized form of covalent lipid modification is the attachment of a complex glycophospholipid (GPI) tail to the C-terminus of certain plasma membrane proteins. This modification affects at least 10% of proteins in the secretory pathway and provides a mechanism to anchor proteins to the cell-surface and facilitate regulated release of their ectodomains. It provides an alternative to the regulated shedding of cell-surface proteins by proteolytic sheddases such as ADAMs (a disintegrin and metalloproteinase) protein family. Attempts at defining the full human GPI-proteome have had limited success to date (see e.g., Elortza et al., 2006). A large scale predictor of GPI precursor sequences in eukaryotic proteins (FragAnchor) is, however, publicly available (Poisson et al., 2007) and predicts the percentage of highly probable GPI-anchored proteins in the eukaryotic proteome as between 0.2 and 2%. This predictor, however, generally fails to identify the o attachment site at which the anchor is attached and a newer algorithm (PredGPI) is superior in this regard (Pierleoni et al., 2008). The ability of phosphatidylinositol-specific phospholipases C (PI-PLC) from various bacterial species to release specific membrane proteins selectively from the cell surface was first recognised for alkaline phosphatase (Ikezawa et al., 1976). The PI-PLC from B. cereus was shown to release alkaline phosphatase in a hydrophilic form which led to intensive studies to isolate and characterize the hydrophobic membrane anchor itself (Low and Zilversmit, 1980). The release process can also be tissue-specific and affected by differential mRNA splicing leading to alternative anchored forms of a protein: transmembrane polypeptideanchored or a GPI-anchored form of the same protein, relevant examples being acetylcholinesterase (AChE) and NCAMs. Although GPI-anchored proteins were first recognized over 20 years ago (in the early 1980s), relatively few GPI anchors have been analyzed in detail. Molecular characterization of various PI-PLC-released proteins indicated that there existed a covalent link between the protein and a phosphatidylinositol (PI) moiety in the membrane bilayer. Subsequent studies established that the anchor contained a core phospholipid-glycan chain consisting of PI-glucosamine-mannose3-phosphoethanolamine attached to the C-terminal amino acid of the mature protein. This core structure appears to be highly conserved among species. The fatty acid associated with the phospholipid can also be species-specific. For example, the trypanosome variant surface glycoprotein differs from mammalian GPI-anchored proteins in containing exclusive myristate. Structural details of GPI-anchored proteins are extensively reviewed in (Paulick and Bertozzi, 2008) and their phospholipase cleavage in Sharom and Lehto (2002). Although release of a membrane protein by PI-PLC is often taken as sufficient evidence for a glycolipid anchor, more rigorous criteria should ideally be used, including immunological analysis. Some GPI anchors are even resistant to hydrolysis by PI-PLC due to additional acylation of the inositol ring (for example erythrocyte AChE) or exist in a double anchored form (GPI and transmembrane anchors) such as the protein Bst-2 (Kupzig et al., 2003) and a mutant, pathogenic form of the prion protein (Stewart and Harris, 2001). Considerable interest has focused on the mode of biosynthesis and attachment of the glycolipid anchor. The deduced C-terminal amino acid sequence of the precursor forms of glycolipid-anchored proteins reveal a predominantly hydrophobic sequence of 20–30 amino acids that is not present at the C-terminus of mature proteins. This C-terminal hydrophobic domain appears to act as a temporary anchor which is exchanged rapidly for a pre-formed precursor glycolipid in a reaction catalyzed by a multisubunit membrane bound transamidase complex whose activity resides on the lumenal face of the endoplasmic reticulum. The transamidase is composed of five essential subunits of which PIG-K (GP18) is the catalytic core being related to a family of cysteine proteases (Orlean and Menon, 2007). This transfer process occurs within a minute or so of translation. Only Asp, Gly, Ala, Cys, Ser and Asn have been identified as the anchor attachment sites (the o site) in naturally occurring proteins suggesting that the residue to which the glycolipid attachment occurs must be small and aliphatic. GPI membrane anchors are present in organisms at most stages of eukaryotic evolution and are found on a functionally and evolutionarily diverse range of proteins (see > Table 14‐1). In addition to NCAMs, these include lymphoid antigens (e.g., Thy-1), cell-surface receptors (e.g., folate receptor, ciliary neurotrophic factor receptor), the prion protein, tumour markers and ectoenzymes. Among the enzymes anchored in

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this way are some forms of AChE (see below) and several peptidases discovered in our laboratory (e.g., membrane dipeptidase and X-Pro aminopeptidase) (Hooper and Turner, 1988; Turner, 1994). GPI-anchored proteins are functionally diverse, but one of their most striking features is their association with lipid microdomains, which consist mainly of sphingolipids and sterols, so-called lipid rafts (for review see Hoetzl et al., 2007). The GPI anchor is a key determinant of raft localization. Much attention has focused on the role of lipid rafts in the amyloidogenic processing of the Alzheimer’s amyloid precursor protein (APP). The attachment of a GPI anchor to the APP b-secretase dramatically enhances amyloid production (Cordy et al., 2003) and the disruption of rafts by lowering cholesterol concentrations decreases amyloid production in vitro (Fassbender et al., 2001; Cordy et al., 2003) and in vivo (Fassbender et al., 2001; Refolo et al., 2001). The b-secretase (BACE-1) is itself highly sensitive to its lipid environment being modulated by glycosphingolipids, glycerophospholipids and sterols (Kalvodova et al., 2005). BACE-1 localization is also regulated by covalent lipid modifications. The cytoplasmic domain of BACE-1 is subject to multiple palmitoylation which aids its localization to rafts (Benjannet et al., 2001). The g-secretase complex components nicastrin and Aph-1 are also S-palmitoylated facilitating the consecutive amyloidogenic processing of APP in rafts (> Figure 14‐3) while the non-amyloidogenic (a-secretase) processing mediated by ADAMs proteinases appears to occur outside of rafts (Reid et al., 2007; Cheng et al., 2009). However, the picture may be more complex since amyloid production is still detectable in cells expressing non-palmitoylated BACE-1 suggesting that this process is not exclusive to rafts (Vetrivel et al., 2009) and, in development, notch processing by g-secretase is mediated in the non-raft phospholipid milieu (Reid et al., 2007). The stability and turnover of BACE-1 is additionally regulated by reversible lysine acetylation in its N-terminal (lumenal) domain (Costantini et al., 2007; Jonas et al., 2008). A more detailed review of the cell biology of BACE-1 can be found in Hunt and Turner (2009).

2.3.1 Shedding and Roles of GPI-Anchored Proteins GPI-anchored proteins are composed of a relatively large hydrophilic moiety which is held relatively loosely in the membrane by a small lipid tail which is located only in the outer leaflet of the bilayer. Hence, . Figure 14‐3 The partitioning of amyloid precursor protein (APP) processing between lipid raft and non-raft domains. Non-amyloidogenic (a-secretase) processing of APP takes place in the bulk phospholipid (non-raft) region of the bilayer whereas amyloidogenic processing is favoured in, but not exclusive to, lipid raft domains where both BACE-1 and components of the g-secretase complex (nicastrin and Aph-1) are S-palmitoylated (Cheng et al., 2009)

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GPI-anchored proteins can transfer spontaneously from the membrane of one cell and be associated with other cells by insertion of their lipid anchors into the cell membrane. This can occur both in vivo and in vitro and has allowed the engineering of specific cell surfaces with GPI-anchored proteins (Medof et al., 1996). The normal function of this ability of GPI-anchored proteins to transfer between membranes is unknown although tumour cells and some pathogens can exploit this process to their own advantage (for review see Lauc and Heffer-Lauc, 2006). Some GPI-anchored proteins can be shed by proteolytic cleavage, usually mediated by an ADAMs type protein, rather than by hydrolysis of the lipid, an example being the prion protein itself (Taylor et al., 2009). The study of GPI-anchored protein sorting has led to some surprising new findings and concepts. Evidence is accumulating that, during their delivery to the surface, different types of plasma membrane protein might be sorted from each other early in this pathway, in the endoplasmic reticulum. Furthermore, membrane-lipid composition and microdomains might have a role in the process of protein sorting in both the secretory and endocytic pathways (Mayor and Riezman, 2004). The exact function of the GPI lipid anchor is still a matter of much speculation and little resolution. It is, however, more than just a simple membrane anchor. It appears to act as a targeting signal directing proteins to the apical surface of polarized cells (Lisanti and Rodriguez-Boulan, 1990), including neurons as first shown for the polarised sorting of Thy-1 in hippocampal neurons (Dotti et al., 1991), although it is not an exclusive signal for this purpose since N-glycans may also play a role (Benting et al., 1999; Pang et al., 2004). GPI-anchored proteins are sorted into lipid rafts before transport to the membrane surface and their localization in such lipid microdomains may explain their involvement in signal transduction processes (Mayor and Riezman, 2004). Since GPI-anchored proteins lack any cytoplasmic domain, however, this signalling is probably mediated through interaction with specific transmembrane proteins. Substantial evidence suggests that GPI-anchored proteins may interact closely with the bilayer surface, so that their functions may be modulated by the biophysical properties of the membrane and the anchor may affect the conformation and activity of its cognate protein. The release of GPI-anchored proteins from the cell surface by specific phospholipases (GPI-specific phospholipases C and D) may play a key role in regulation of their surface expression and functional properties. GPI-anchored proteins are particularly abundant in protozoa and represent the major carbohydrate modification of many cell-surface parasite proteins. Although the core glycan is highly conserved, subtle differences both to GPI structures and biosynthetic pathways occur allowing opportunities for therapeutic intervention (for review see de Macedo et al., 2003). Examples of some specific neural GPI-anchored proteins are given briefly below.

2.3.2 Prion Proteins Prions are responsible for degenerative encephalopathies through inducing conformational changes from the normal cellular form of the prion protein (PrPc) to the scrapie isoform (PrPsc) PrPc itself is a cell surface GPI-anchored glycoprotein. Although highly conserved among species and with expression in most tissues but, in particular, with high levels in the nervous system, a role for PrPc has remained obscure over the years. Initial scepticism about such a role was mainly due to the absence of a gross phenotype alteration in PrPc null mice. More recently, some feasible biological functions for PrPc have emerged. The protein binds copper and the resulting complex may be responsible for cell protection against oxidative stress (Watt et al., 2005). PrPc is also a high-affinity ligand for laminin, and induces neuronal cell adhesion, neurite extension and maintenance (Lima et al., 2007). Since PrPc recycles from the plasma membrane to an intracellular compartment, which is induced by copper binding, it is also possible that the internalization mechanism serves to terminate signaling events induced by PrPc ligand binding (for reviews see Martins and Brentani, 2002; Baldwin, 2005; Taylor and Hooper, 2006). PrPc acts to inhibit the activity of the APP b-secretase thereby inhibiting the formation of Ab peptide and hence providing a protective role against amyloidogenesis in the brain (Parkin et al., 2007; Hooper and Turner, 2008). PrPc has recently been shown to act as a receptor for Aβ oligomers leading to impairment of synaptic plasticity (Laure´n et al., 2009).

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2.3.3 Neural GPI-Anchored Cell Adhesion Molecules An important class of GPI-anchored proteins in the nervous system is represented by the NCAMs, which mediate cell–cell and cell–matrix interactions and play a vital role in embryonic development of the nervous system as well as in the maintenance and regeneration of nerves in adults. The NCAMs exist in alternatively spliced forms that can possess either a transmembrane or GPI anchor, or be secreted as a hydrophilic species. In addition to NCAM itself in the CNS, other adhesion molecules include L1, N-cadherin, Drosophila fasciclin as well as the mammalian glycoproteins Thy-1, TAG1, N-CAM, F11 and F3/contactin, OBCAM and neurotrimin (Walsh and Doherty, 1991; Struyk et al., 1995; Karagogeos, 2003). Such adhesion molecules are also implicated in cell migration, axonal growth and synaptogenesis through initiating transmembrane-signaling cascades and thereby modulating synaptic plasticity, including learning and memory. Neuronal cell adhesion molecules commonly contain one or more immunoglobulin-like domains and are hence generically referred to as IgCAMs (Walsh and Doherty, 1997) and play key roles in neuronal circuitry and its development. Certain human inherited neuronal disorders correspond to mutations in IgCAM genes emphasizing their fundamental developmental roles (Katidou et al., 2008). The modular protein structure of the IgCAMs provides multiple binding sites for interaction with a variety of ligands. Thus, the GPI-anchored neuronal protein F3/contactin mediates neuronal-glial contacts through its association with extracellular matrix components (e.g., the tenascins) and hence influencing axonal growth and fasciculation (Falk et al., 2002). Another major role of this GPI-anchored protein, ranging from humans to Drosophila, is in organising axonal insulation of myelinated fibers at specific subdomains of the nodes of Ranvier. A distinctive group of the Ig superfamily of GPI-anchored NCAMs is represented by the IgLON family proteins, which include the opiate binding-cell adhesion molecule (OBCAM or OPCML), neurotrimin (HNT), Kilon and limbic-associated membrane protein (LSAMP) (Hachisuka et al., 2000; McNamee et al., 2002; Hashimoto et al., 2008). The IgLON family proteins consist of three Ig domains, the third of which is attached to the GPI anchor. These proteins are distributed differently in the CNS during neuronal development and are important in the specification of neuronal connectivity. OBCAM expression is down-regulated in human gliomas and other brain tumours (Reed et al., 2007) suggesting its possible role as a brain tumour suppressor.

2.3.4 GPI-Anchoring of AChE The key synaptic membrane-associated enzyme regulating acetylcholine levels, acetylcholinesterase (AChE), is a globular, glycosylated protein that lacks a transmembrane peptide anchor region. Its attachment to the membrane occurs through several different mechanisms, including lipid modification, and varies between cell types and organisms (see e.g., Nalivaeva and Turner, 2001). Alternative splicing of the AChE gene regulates the occurrence of several different molecular forms of the enzyme, which all contain the same catalytic domain but with varied C-termini (Massoulie´ et al., 2005). In mammals, a read-through product of the AChE gene (AChER) generates a soluble monomer which was shown to be up-regulated in the brain under stress conditions and is characteristic for some neurological disorders (Zimmerman and Soreq, 2006). In the brain the main form of AChE is a membrane-bound variant, AChET, which is also present in muscle. This CNS common transcript encodes an isoform characterized by the presence in the C-terminal peptide of 40 amino acid residues, called the T peptide. Disulphide bonding between AChET units via the T peptide gives rise to amphipathic homodimers and homotetramers of AChE. The T peptide, which is organised as an alpha-helix, also allows the AChE complexes to bind to hydrophobic, proline-rich domains of specific membrane-anchoring proteins such as ‘‘collagen-like Q subunit’’ (ColQ) at the neuromuscular junction and the ‘‘proline-rich membrane anchor’’, PRiMA, in the brain (Massoulie, 2002). Most of the AChE in the CNS is thus in the form of tetrameric AChET (designated G4) bound to PRiMA (Navaratnam et al., 2000; Perrier et al., 2002; Massoulie´ et al., 2005). It has been proposed that the interaction of AChE with PRiMA forms the basis of a larger multiprotein complex (Navaratnam et al., 2000). In the muscles, AChE-T subunits are linked to the cell membrane via a collagen tail (ColQ)

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represented mostly by asymmetric species A4, A8, and A12. The A12 symmetric form of AChET is dominant at neuromuscular junctions (Tsim et al., 2008). Another form of AChE, possessing the H domain (AChEH), contains a signal for GPI-anchoring and is characteristic of blood cells. The GPI-anchored form of AChE is found predominantly in erythrocytes in humans and is represented by G2 dimers. The structures of some GPI anchors of AChEH have been solved but their functions, apart from membrane attachment and dynamics, still remain to be determined. The GPI anchor consists of a core glycan identical to those observed in GPI anchors of other proteins and has the following structure: ethanolamine-P-6Mana1-2Mana1-6Mana1-4GlcN attached to an inositol phospholipid (Deeg et al., 1992). Attached to the conserved core are variable side chains, which may be proteinor tissue-specific and can vary between species (Brewis et al., 1995). The phospholipid component of GPI anchors may differ in different species. Thus, AChE from bovine erythrocytes contains mainly stearic acid (C18:0), whereas in human erythrocytes it possesses C22:4, 22:5 or 22:6 acyl groups at the C-2 position of glycerol (Roberts et al., 1987). The human GPI anchor also contains a plasmanylinositol with a palmitoyl group on an inositol hydroxyl which makes it resistant to phospholipase C cleavage (Roberts et al., 1988). The physiological significance of the GPI-anchored form of AChE on the surface of erythrocytes relates not only to the hydrolytic activity of the enzyme but also provides the cell surface with specific antigenic determinants. AChE is one of a number of GPI-anchored molecules of the human erythrocyte cell surface that contribute to the blood group antigen properties. Thus, the Cartwright (Yt) antigens reside on AChE. In erythroid cells, a small exon that encodes the signal for attachment of the GPI anchor is retained in a tissue-specific process. Identification of the AChE gene has also allowed assignment of the Cartwright Yt locus to chromosome 7 and identification of the molecular basis of the Yt(a/b) polymorphism which occurs in exon 2 of the AChE gene (Telen, 1995). By contrast, in Drosophila, a GPI-anchored form of AChE encoded by the Ace locus is the only form of this enzyme. To assess what role the GPI anchor plays in the physiology of AChE, Incardona and Rosenberry (1996) replaced the wild-type GPI-AChE with a chimaeric, transmembrane form AChE (TM-AChE) in the fly nervous system. The individuals expressing TM-AChE at about 30% of normal levels showed no gross behavioural differences from wild-type flies but had a reduced lifespan and diminished locomotor activity. However, at the morphological level, there were no significant differences in localization of GPI-AChE compared with TM-AChE in the flies, suggesting that cellular targeting of AChE is not a function of the GPI anchor in this case. It was therefore suggested that the Drosophila GPI anchor of AChE plays some subtle cellular role in neuronal physiology (Incardona and Rosenberry, 1996).

2.3.5 Other GPI-Anchored Proteins There are a number of other families of GPI-anchored proteins of relevance to the nervous system. For example, one group is represented by ligands for the Eph-related tyrosine kinase receptors. Tyrosine kinase receptors are abundant in the CNS playing a critical role in development and the Eph family represents the largest cohort of these being expressed exclusively or predominantly in the nervous system (Snider, 1994; Tuzi and Gullick, 1994). The Eph family consists of two subclasses distinguished by the nature of their ligands: the EphA receptors bind the GPI-anchored ephrin-As, and the EphB receptors bind the transmembrane-tethered ephrin-Bs (Klein, 2001; Kno¨ll and Drescher, 2002). Ephrin/Eph signaling appears to be involved in axon bundle formation, repulsive axon guidance, cell migration, topographic mapping and angiogenesis (Winslow et al., 1995). They also play an important role in neural stem-cell proliferation and migration in adult mouse brain. The GPI-anchored forms appear to act via lipid raft domains, for example, ephrin-A5 is involved in compartmentalized cell signallng within a caveolae-like membrane microdomain and ephrin A2 interacts with caveolin-1 (Davy et al., 1999; Vihanto et al., 2006). Recently, it was shown that ephrin-A2, -A3 and -A5 are involved in the correct organization of projections from retina to superior colliculus (Cang et al., 2008). During cell–cell interactions, GPI-anchored ephrin-As form strong complexes with Eph receptors inducing various cell signaling events. This interaction and, consequently, the effects of ephrins (e.g., ephA2), is terminated by a proteolytic event which involves an ADAMs metalloproteinase, most likely ADAM10 (Hattori et al., 2000).

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The Thy-1 antigen of rodent thymocytes and brain was the first eukaryotic membrane molecule for which biochemical data clearly suggested membrane integration via a non-protein tail, which was further characterized as a glycophospholipid structure attached to the C-terminal cysteine residue (Tse et al., 1985). The detailed structure and comparative analysis of the GPI anchor for Thy-1 from analysis of Trypanosoma brucei and rat brain was solved by Homans and colleagues (1988). Thy-1, a member of the Ig superfamily, is one of the most abundant glycoproteins on mammalian neurons. Thy-1 appears to play a role in synapse formation and cell adhesion, as well as in axonal or dendritic development and regeneration (Hill et al., 1988; Dreyer et al., 1995). Nevertheless, Thy-1 appears not to be essential for the normal development and maintenance of functional synaptic connections, as shown from studies of Thy-1 null mice (Barlow et al., 2002). Presumably there is a level of redundancy and other guidance and signaling molecules can compensate for the loss of Thy-1. A fuller discussion of the regulatory roles of Thy-1 in the CNS is to be found in Rege and Hagood, (2006).

3

Functions of Lipid Modifications of Proteins

As shown by this survey of the properties of lipid-anchored proteins of the CNS, the biological functions of lipid modifications of proteins are likely to be many and varied, dependent on the combination of lipid modifications, but with a primary role in permitting the stable membrane association of proteins. This clearly is not an exclusive role since some myristoylated proteins are cytosolic and some palmitoylated proteins are transmembrane polypeptides (for example, the adrenergic receptor). In theses cases the acyl group(s) may stabilize protein-protein interactions, allowing regulation of protein functions. For example, palmitoylation of the adrenergic and related receptors may modulate their interaction with G proteins. Another function for palmitate may be in directing intracellular transport of proteins since palmitoylation of the neuronal protein GAP-43 targets it to growth cones. That numerous proteins involved in cellular signaling exist in lipid-modified forms strongly suggests a role for these modification processes in cell regulation and development (see James and Olson, 1990; Resh, 2006). Inhibitors of acylation and isoprenylation could therefore have considerable therapeutic potential as yet not effectively exploited. The reversibility of some lipid modifications provides a dynamic nature to their role permitting cycling between membrane and cytosol, or the extracellular environment as with the phospholipase release of GPI-anchored proteins. The range of lipid modifications and their widespread occurrence attests to their biological importance and new modifications, as well as new combinations and permutations of lipid modifications, probably remain to be discovered. How these may influence neuronal function in health and disease, and how they may be exploited therapeutically, remains to be determined

Acknowledgments We thank the UK Medical Research Council, Yorkshire Cancer Research, the Wellcome Trust, the Royal Society and the E.U. INTAS scheme for support of our research.

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Lipid anchors to proteins Tuzi NL, Gullick WJ. 1994. eph, the largest known family of putative growth factor receptors. Br J Cancer 69: 417-421. Vetrivel KS, Meckler X, Chen Y, Nguyen PD, Seidah NG, et al. 2009. Alzheimer Disease Ab Production in the Absence of S-Palmitoylation-dependent Targeting of BACE1 to Lipid Rafts. J Biol Chem 284: 3793-3803. Vihanto MM, Vindis C, Djonov V, Cerretti DP, Huynh-Do U. 2006. Caveolin-1 is required for signaling and membrane targeting of EphB1 receptor tyrosine kinase. J Cell Sci 119: 2299-2309. Walsh FS, Doherty P. 1991. Glycosylphosphatidylinositol anchored recognition molecules that function in axonal fasciculation, growth and guidance in the nervous system. Cell Biol Int Rep 15: 1151-1156. Walsh FS, Doherty P. 1997. Neural cell adhesion molecules of the immunoglobulin superfamily: Role in axon growth and guidance. Annu Rev Cell Dev Biol 13: 425-456. Wang HY, Tao J, Shumay E, Malbon CC. 2006. G-Proteincoupled receptor-associated A-kinase anchoring proteins: AKAP79 and AKAP250 (gravin). Eur J Cell Biol 85: 643-650. Watt NT, Taylor DR, Gillott A, Thomas DA, Perera WS, et al. 2005. Reactive oxygen species-mediated b-cleavage of the

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Bioactive Sphingolipids: An Overview on Ceramide, Ceramide 1-Phosphate Dihydroceramide, Sphingosine, Sphingosine 1-Phosphate

J. M. Kraveka . Y. A. Hannun

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 374

2 2.1 2.2 2.3 2.4

Overview of Sphingolipid Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 374 The De Novo Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Formation of Complex Sphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Sphingomyelinase Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Salvage/Recycling Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378

3 3.1 3.2 3.3 3.4 3.5

Bioactive Sphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378 Dihydroceramide (dhCer) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378 Ceramide (Cer) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378 Sphingosine (Sph) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 379 Ceramide 1-Phosphate (C1P) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 379 Sphingosine 1-Phosphate (S1P) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 379

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Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 379

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_15, # Springer ScienceþBusiness Media, LLC 2009

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Abstract: Bioactive sphingolipids such as ceramide, ceramide 1-phosphate, dihydroceramide, sphingosine, and sphingosine 1-phosphate have key roles in cell proliferation, differentiation, senescence, apoptosis, migration, carcinogenesis, inflammation, and angiogenesis. There has been much progress made in understanding the complex pathways of sphingolipid metabolism and identifying the enzymes involved in sphingolipid production. This chapter reviews the structure, metabolism and functions of these bioactive sphingolipids. List of Abbreviations: alk-SMase, alkaline sphingomyelinase; aSMase, acid sphingomyelinase; C1P, ceramide 1-phoshpate; CDase, ceramidase; Cer, ceramide; CERK, ceramide kinase; CerS, (dihydro)ceramide synthases; DAG, diacylglycerol; DEGS-1, dihydroceramide desaturase; dhCer, dihydroceramide; ER, endoplasmic reticulum; GalCer, galactosylceramide; GCS, glucosylceramide synthase; GluCer glucosylceramide; GSLs; glycosphingolipids; haCER, human alkaline ceramidase; haPHC, human alkaline phytoceramidase; MAMs, mitochondria-associated membranes; nSMase, neutral sphingomyelinase; PKC, protein kinase C; Rb, retinoblastoma protein; S1P, sphingosine 1-phosphate; SK, sphingosine kinases; SM, sphingomyelin; SMases, sphingomyelinases; Sph, sphingosine

1

Introduction

Sphingolipids comprise a class of lipids that share the presence of a sphingosine (or related sphingoid) base in the backbone of their structures. Research in the past two decades has shown that sphingolipids, in addition to their roles as structural components of cell membranes, play important roles as regulators of signal transduction in cell differentiation, cell proliferation, inflammation and apoptosis (Spiegel and Milstien, 2003; Futerman and Hannun, 2004; Ogretmen and Hannun, 2004; Taha et al., 2006; Zeidan and Hannun, 2007; Hannun and Obeid, 2008; Wymann and Schneiter, 2008). The most studied bioactive sphingolipids have been ceramide (Cer), sphingosine 1-phosphate (S1P), and sphingosine (Sph). Recent studies have also focused on dihydroceramide (dhCer), ceramide 1-phoshpate (C1P) and glucosylceramide (GluCer). The metabolic pathways of sphingolipids are very complex with several potential points of regulation and modulation. Most enzymes of sphingolipid metabolism have specific subcellular localization(s), thereby exerting profound effects on the signaling and regulatory functions of the generated sphingolipid in a specific compartment. Cer, one of the most studied bioactive sphingolipids, functions as the central building block for sphingolipids; it is generated in cells by multiple regulated metabolic pathways. Functionally, Cer participates in regulation of cell growth, apoptosis, stress responses, and other signaling pathways (Hannun and Obeid, 2008). DhCer was previously thought to be an inactive precursor of Cer; however, recent studies have demonstrated it to be involved in growth arrest and autophagy (Zheng et al., 2006; Kraveka et al., 2007). Sph inhibits protein kinase C (PKC) and induces cell cycle arrest and apoptosis (Taha et al., 2006). Both S1P and C1P are involved in cell proliferation and can counteract the effects of Cer and Sph; S1P is one of the best studied intra and intercellular messengers (Chalfant and Spiegel, 2005).

2

Overview of Sphingolipid Metabolism

Structurally, sphingolipids are based on Cer which is composed of a sphingosine base and an amide-linked fatty acyl chain varying in length from 14 to 26 carbons. (> Figure 15-1) The various complex sphingolipids contain various substituents at the 1-OH position of Cer (e.g., phosphocholine in sphingomyelin and glucose in GluCer). There are multiple pathways for the generation and/or regulation of the levels of ceramide, S1P, and the other bioactive sphingolipids (> Figure 15-2). Many of these pathways reside in specific sub-cellular compartments and respond to various extra- and intracellular stimuli. Of these pathways, the best studied are the de novo pathway, the sphingosine kinase (SK) pathway, and the breakdown of sphingomyelin (SM) through the action of sphingomyelinases (SMases). Other emerging pathways include the Cer kinase pathway and the salvage pathway which recycles complex sphingolipids.

. Figure 15-1 Bioactive sphingolipids. Ceramide is composed of a sphingosine base (18 carbons) and an amide-linked fatty acid (14–26 carbons). Complex sphingolipids are composed of a hydrophilic headgroup (R) attached to the lipophilic ceramide backbone. These headgroups may be phosphocholine in sphingomyelin or sugars in glycosphingolipids

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. Figure 15-2 Sphingolipid metabolism. Ceramide is the central building block for complex sphingolipids. Enzymes are shown in italics

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2.1 The De Novo Pathway This pathway has been well studied (Merrill, 2002; Futerman and Hannun, 2004; Futerman and Riezman, 2005). The condensation of serine and palmitoyl CoA by serine palmitoyltransferase in the endoplasmic reticulum (ER) initiates the pathway and generates ketosphinganine, which is then reduced to dihydroshingosine (sphinganine), followed by the acylation to dhCer by (dihydro)ceramide synthases (CerS). CerS are also referred to as Lass (longevity-assurance homolog) family members. Six mammalian CerS have been identified molecularly, and each CerS protein exerts specificity for the generation of distinct endogenous Cers with fatty acid chain length specificity (Ogretmen, 2006; Pewzner-Jung et al., 2006). In mammals, dhCers with distinct fatty acid chain lengths are then desaturated by dihydroceramide desaturase (DEGS-1) (Michel et al., 1997) to form the corresponding Cers. The de novo synthesis of Cer occurs in the ER and possibly in ER-associated membranes, such as the perinuclear membrane and mitochondria-associated membranes (MAMs). Cer formed in this compartment is transported to the Golgi, which is the site of synthesis of SM and GluCer, with the latter also serving as the precursor for complex glycosphingolipids (GSLs). The transport of Cer to the Golgi occurs either through the action of the transfer protein CERT (Hanada et al., 2003), which specifically delivers Cer for SM synthesis, or through vesicular transport, which delivers Cer for the synthesis of GluCer.

2.2 Formation of Complex Sphingolipids Once Cer is generated, it can be glycosylated by GluCer synthase (GCS) to form GluCer on the cytoplasmic surface of Golgi. GluCer then serves as the precursor for GSLs. Cer may also be galactosylated to galactosylceramide (GalCer) by GalCer synthase in the ER. Sulfatides and Gala-series GSLs are formed from GalCer. In turn, GSLs are hydrolyzed by b-glucosidases and galactosidases to regenerate Cer (Tettamanti, 2004). In addition, Cer can be converted into a number of other bioactive sphingolipids, including C1P, Sph, and S1P.

2.3 Sphingomyelinase Pathways Cer can be also generated from the hydrolysis of SM through the action of either acid or neutral SMases (Marchesini and Hannun, 2004; Clarke and Hannun, 2006). These enzymes break down SM to produce Cer and phosphocholine, and are stimulated in response to TNF-a (Schwandner et al., 1998; Luberto et al., 2002), Fas ligand (Lin et al., 2000), or oxidative stress (Goldkorn et al., 1998). The SMase-mediated hydrolysis of SM has emerged as a major pathway of stress-induced Cer generation. Conversely, sphingomyelin synthase (SMS) transfers the headgroup of phosphatidylcholine to Cer, and generates SM and diacylglycerol (DAG) in the process. This pathway has been suggested to regulate the levels of not only SM and Cer, but also DAG (Villani et al., 2008) as well as the activation of NFkB (Luberto et al., 2000; Hailemariam et al., 2008). SMases are characterized according to their pH optima, metal ion dependence, and subcellular localization. Several have been characterized: zinc ion-dependent acid Smases (lysosomal aSMase and secretory acid Smase (aSMase or SMPD1), neutral magnesium ion-dependent SMases (nSMase1, nSMase2, and nSMase3), and alkaline SMase (alk-SMase). aSMase is deficient in Niemann-Pick disease (Schneider and Kennedy, 1967; Yamaguchi and Suzuki, 1977). nSMase1 (SMPD2) is localized to the ER and is expressed in most tissues (Tomiuk et al., 2000); however, its cellular functions appears to be more related to its activity as a lysoPAF phospholipase rather than as an SMase (Sawai et al., 1999; Tomiuk et al., 2000). nSMase2 (SMPD3) has been localized to the Golgi and plasma membrane and is highly expressed in neuronal tissues (Hofmann et al., 2000). nSMase3 (SMPD4) is localized in the ER and Golgi; it is expressed in most tissues, with the greatest abundance in muscle (Krut et al., 2006). Alk-SMase is localized in the Golgi and expressed in the gastro-intestinal tract (Duan et al., 2003).

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2.4 Salvage/Recycling Pathway The sphingolipid recycling or salvage pathway refers to the various mechanisms of Cer generation from the catabolism of complex sphingolipids which are broken down into Sph, which is then reused through reacylation to produce Cer (Kitatani et al., 2008). This pathway involves a number of key enzymes that include SMases, cerebrosidases, ceramidases (CDases), and CerS. It has been estimated to contribute 50– 90% of sphingolipid biosynthesis. Degradation of sphingolipids and GSLs takes place mostly in the acidic subcellular compartments, the late endosomes and the lysosomes. SM is converted to Cer by acid SMase. Cer can be deacylated with loss of the fatty acid from the amide bond through the action of acid CDases to yield Sph. Sph can then translocate across the lysosome where it can be either re-acylated to Cer or phosphorylated by SK 1 or 2 to generate S1P. S1P can be cleared by the action of specific phosphatases that regenerate Sph or by the action of a lyase that cleaves S1P into ethanolamine-1-phosphate and a C16fatty-aldehyde. Like SMases, CDases are characterized according to their pH optima and subcellular localization. Three types of CDases have been described: acid, neutral and alkaline. Acid CDase, a lysosomal enzyme, is deficient in Farber’s disease (Gatt, 1963; Bernardo et al., 1995). Neutral CDase has been localized to the plasma membrane (Hwang et al., 2005), mitochondria (El Bawab et al., 2000) and is extracellularly secreted from endothelial cells. Three alkaline CDases have been identified in humans. The haCER1 is localized to the ER and preferentially hydrolyzes Cers with a very long-chain unsaturated fatty acid (C24:1) (Sun et al., 2008). haCER2 is localized to the Golgi complex (Xu et al., 2006). The human alkaline phytoceramidase (haPHC) utilizes phytoceramide as substrate and has not been demonstrated to play a role in the regulation of the generation of Sph or S1P (Mao et al., 2001). Interestingly, some of these enzymes demonstrate reverse activity, i.e., CoA-independent ceramide synthase function (El Bawab et al., 2001; Okino et al., 2003). In mammals, two isoforms of SK (1 and 2) have been cloned and characterized (Kohama et al., 1998; Liu et al., 2000; Pitson et al., 2000). SK1 is located in the cytoplasm, but stimuli such as PDGF and PKC can change its localization to intracellular membranes or the plasma membrane (Rosenfeldt et al., 2001; Johnson et al., 2002). SK2 is a also localized to both cytosol and nucleus (Igarashi et al., 2003). SK1 is most highly expressed in the brain, heart, thymus, spleen, kidney, and lung (Melendez et al., 2000) whereas SK2 is highest in the kidney and the liver (Liu et al., 2000). S1P lyase has been localized to the ER and is mist highly expressed in small intestine, colon, thymus, and liver (Zhou and Saba, 1998; Ikeda et al., 2004).

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Bioactive Sphingolipids

3.1 Dihydroceramide (dhCer) Previous studies on the biological activity of dhCers using their short-chain analogs concluded that dhCers were inactive in inducing cell death and apoptosis (Bielawska et al., 1993; Sugiki et al., 2000; Ahn and Schroeder, 2002). However, recent publications are beginning to show biologic functions of dhCers in cell cycle arrest, hypophosphorylation of the retinoblastoma protein (Rb), and autophagy (Jiang et al., 2004; Stiban et al., 2006; Zheng et al., 2006; Kraveka et al., 2007). Inhibitors of the DEGS-1 enzyme lead to the accumulation of endogenous dhCers. These inhibitors include the cyclopropene ceramide, GT-11 (also known as C8-Cyclopropenylceramide or C8-CPPC) (Triola et al., 2004). The synthetic retinoid, fenretinide (Schulz et al., 2006; Zheng et al., 2006; Kraveka et al., 2007), and g-tocopherol (Jiang et al., 2004) have also been shown to inhibit the desaturase although it is not known if these are direct effects or not.

3.2 Ceramide (Cer) Cer has been shown to be involved in the cellular stress responses, differentiation, cell cycle arrest, senescence, and apoptosis, but also in neurodegeneration, inflammation, cancer, and infection.

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Many stimuli and especially chemotherapeutic agents activate the de novo (cannabinoids, daunorubicin, etoposide, camptothecin, fludarabine, gemcitabine) or SM (cytarabine, actinomycin D, etoposide, cisplatin) (Strum et al., 1994; Bose et al., 1995; Suzuki et al., 1997; Biswal et al., 2000; Perry et al., 2000; Chalfant et al., 2002; Gomez Del Pulgar et al., 2002; Lacour et al., 2004) pathways. TNF, UV and g radiation have been shown to activate SMases (Kim et al., 1991; Haimovitz-Friedman et al., 1994; Liu et al., 1998; Zhang et al., 2001). Cer exerts its downstream effects via activation of ceramide-activated serine–threonine phosphatases (CAPPs), such as protein phosphates 1 and 2 (PP1 and PP2A) (Dobrowsky et al., 1993; Wolff et al., 1994) In addition, Cer has been shown to activate PKCz, the kinase KSR, and cathepsin D (Lozano et al., 1994; Heinrich et al., 1999; Conway et al., 2000).

3.3 Sphingosine (Sph) Sph was the first bioactive sphingolipid to be identified, and it was reported to inhibit PKC (Hannun et al., 1986). Sph, like Cer, is a pro-apoptotic molecule (Taha et al., 2006). Sph mediates cell death through the inhibition of the prosurvival factors PKC, ERK (Jarvis et al., 1997) and Akt/Protein kinase B (Akt/PKB) (Chang et al., 2001) and induces death via SDK1, which leads to Bax activation through inhibition of 14-3-3 proteins (Hamaguchi et al., 2003). Sph also induces Bid cleavage and mitochondrial cytochrome C release, as well as the activation of caspases 3 and 7 and the cleavage of PARP (Hung et al., 1999; Cuvillier et al., 2000; Cuvillier et al., 2001).

3.4 Ceramide 1-Phosphate (C1P) C1P is produced in mammalian cells by the phosphorylation of Cer by ceramide kinase (CERK) (Sugiura et al., 2002). Since then, it has been reported to have several pro-proliferative functions (Gomez-Munoz, 2004; Chalfant and Spiegel, 2005). It has been reported to induce DNA synthesis (Gomez-Munoz et al., 1995), block caspases, inhibit ASMase (Gomez-Munoz et al., 2004) and promote phagosome formation (Hinkovska-Galcheva et al., 2005). It is involved in mast cell degranulation (Mitsutake et al., 2004). It has been shown to also serve as an activator of cytosolic phospholipase A2 (cPLA2a) (Pettus et al., 2004), and thus may function as a key regulator of eicosanoid synthesis.

3.5 Sphingosine 1-Phosphate (S1P) S1P is an anti-apoptotic molecule that blocks the effects of Cer. It has been demonstrated to play a role in cell proliferation and survival, adhesion, angiogenesis, cell migration, and inflammation (Spiegel and Milstien, 2003; Chalfant and Spiegel, 2005). S1P functions mainly via binding to the G-protein-coupled receptors (S1P1–5Rs). Many growth factors and cytokines (e.g., VEGF, PDGF, TNF-a) activate SKs, thereby leading to increases in S1P levels. S1P is also a key mediator in inflammation by activating cyclooxygenase2 (COX-2) (Pettus et al., 2003).

4

Conclusions

The role of sphingolipids as bioactive molecules in cell regulation and signal transduction has been established. However, more research is needed to fully understand the complexities of sphingolipid metabolism, its regulation, its subcellular compartmentalization, and signaling pathways. Understanding these mechanisms will help reveal the pathogenesis of many diseases and ultimately serve as the foundation for developing novel therapies.

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The Endocannabinoid System and its Manifold Central Actions

M. Maccarrone

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 386

2 2.1 2.1.1 2.2 2.2.1 2.2.2 2.2.3 2.3 2.3.1 2.4 2.4.1 2.4.2 2.5

Metabolism of AEA and 2-AG . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 388 Synthesis of AEA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 388 NAPE-PLD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 388 Degradation of AEA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 388 AMT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 390 FAAH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 390 NAAA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 391 Synthesis of 2-AG . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 391 DAGL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 391 Degradation of 2-AG . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 391 2-AG Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 392 MAGL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393 Transport Across the Blood–brain Barrier . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393

3 3.1 3.2

Molecular Targets and Signaling Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 394 Cannabinoid Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 394 Vanilloid Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 395

4

Modulation by Lipid Rafts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 395

5 5.1 5.2

Biological Activities within the CNS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 397 Interaction with Steroids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 398 Control of Cell Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399

6

Conclusions and Future Avenues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_16, # Springer ScienceþBusiness Media, LLC 2009

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The endocannabinoid system and its manifold central actions

Abstract: Endocannabinoids are amides, esters, and ethers of long-chain polyunsaturated fatty acids, which act as endogenous agonists of cannabinoid receptors. Thus, they are able to mimic several pharmacological effects of D9-tetrahydrocannabinol, the psychoactive principle of Cannabis sativa preparations like hashish and marijuana. Anandamide (N-arachidonoylethanolamine) and 2-arachidonoylglycerol are the best-studied members of this new class of lipid mediators. It is now widely accepted that the biological activity of endocannabinoids is largely dependent on a ‘‘metabolic control,’’ which modulates the effects of these substances by modulating their in vivo concentration. Therefore, the metabolic routes that allow synthesis, transport, and degradation of endocannabinoids, and that altogether form the so-called ‘‘endocannabinoid system,’’ are the focus of intense research. This new system will be reviewed in the chapter, along with the molecular targets of endocannabinoids, and the signal transduction pathways triggered thereof. The aim of this update is also to put in a better perspective the cross-talks of endocannabinoids with other signaling molecules, and their implications for the manifold biological activities of these lipids within the central nervous system. List of Abbreviations: AC, adenylyl cyclase; AEA, anandamide (N‐arachidonoylethanolamine); 2‐AG, 2‐arachidonoylglycerol; AMT, AEA membrane transporter; ASK1, apoptosis signal‐regulating kinase 1; BBM, bovine brain microvessels; CB1/2R, type 1/2 cannabinoid receptor; CNS, central nervous system; CPAE, calf pulmonary artery endothelium; CRE, cAMP response element; DAG, diacylglycerol; DAGL, diacylglycerol lipase; DSE, depolarization‐induced suppression of excitation; DSI, depolarization‐induced suppression of inhibition; E, estrogen (17b‐estradiol); ER, estrogen receptor; ERK, extracellular signal‐ regulated kinase; FAAH, fatty acid amide hydrolase; FAK, focal adhesion kinase; GABA, g‐aminobutyric acid; GPCR, G protein‐coupled receptor; HPA, hypothalamic‐pituitary‐adrenal; HUVEC, human umbilical vein endothelial cells; JNK, c‐Jun N‐terminal kinase; LTD, long term depression; LTP, long term potentiation; MAFP, methyl‐arachidonoyl fluorophosphonate; MAGL, monoacylglycerol lipase; MAPK, mitogen‐ activated protein kinase; MCD, methyl‐b‐cyclodextrin; mGluR, group I metabotropic glutamate receptor; NAAA, N‐acylethanolamine‐hydrolyzing acid amidase; NADA, N‐arachidonoyldopamine; NAE, N‐acylethanolamine; NAPE, N‐acyl‐phosphatidylethanolamine; NArPE, N‐arachidonoylphosphatidylethanolamine; NAT, N‐acyltransferase; NGF, nerve growth factor; NMDA, N‐methyl‐D‐aspartate; NO, nitric oxide; NOS, nitric oxide synthase; OEA, N‐oleoylethanolamine; P, progesterone; PEA, N‐palmitoylethanolamine; PI3K, phosphatidylinositol 3‐kinase; PKA/B, protein kinase A/B; PL A1/C/D, phospholipase A1C, D; PRL, prolactin; PVN, paraventricular nucleus; QSAR, quantitative structure‐activity‐relationship; RBE, rat brain endothelium; SEA, N‐stearoylethanolamine; STAT, signal transducer and activator of transcription; TRPV1, transient receptor potential channel vanilloid receptor subunit 1

1

Introduction

The psychoactive principle of C. sativa, D9-tetrahydrocannabinol (THC; > Figure 16-1), has two main molecular targets: type-1 (CB1R) and type-2 (CB2R) cannabinoid receptors (Howlett et al., 2002, 2004; Pertwee and Ross, 2002), which are both localized centrally and peripherally. In a few years a number of endogenous agonists of CB receptors were characterized, i.e., amides, esters, and ethers of long-chain polyunsaturated fatty acids collectively termed ‘‘endocannabinoids’’ (Mechoulam et al., 2002; Piomelli, 2003; De Petrocellis et al., 2004). The distribution of these bioactive lipids in mammalian cells reflects the membrane content of N-acylethanolamines (NAEs), which exist in almost all animal tissues and are ubiquitously present in the animal kingdom (Hansen et al., 2000). Two arachidonate derivatives, the amide N-arachidonoyl-ethanolamine (anandamide, AEA) and the ester 2-arachidonoyl-glycerol (2-AG), both shown in > Figure 16-1, are the most biologically active endocannabinoids described to date (Piomelli, 2003; De Petrocellis et al., 2004). In addition, the ether 2-arachidonoyl-glyceryl-ether (noladin ether; > Figure 16-1) has been shown to act as an endocannabinoid (Hanus et al., 2001), but its actual physiological relevance remains a matter of debate (Oka et al., 2003). Furthermore an ‘‘inverted anandamide,’’ O-arachidonoyl-ethanolamine (virodhamine; > Figure 16-1), has been shown to behave as a partial agonist or as a full agonist at CB1 or CB2 receptors, respectively (Porter et al., 2002). Instead, the amides N-oleoyl-ethanolamine (OEA), N-palmitoyl-ethanolamine (PEA), and N-stearoylethanolamine (SEA) are better considered ‘‘endocannabinoid-like’’ compounds, because they do not

. Figure 16-1 Chemical structures of plant-derived (THC) and endogenous (AEA, 2-AG, noladin ether, virodhamine, and NADA) cannabinoids

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activate CB receptors. However, these compounds seem to have an ‘‘entourage effect,’’ i.e., they might potentiate the activity of AEA or 2-AG by inhibiting their degradation (Ben-Shabat et al., 1998; Lambert and Di Marzo, 1999; De Petrocellis et al., 2004). In just one decade, endocannabinoids have been shown to play manifold roles, both in the central nervous system (CNS) and in the periphery, which recently have been the subject of comprehensive reviews (Di Carlo and Izzo, 2003; Iversen, 2003; Piomelli, 2003; Di Marzo et al., 2004; Fowler, 2005). In particular, it is now widely accepted that the biological activity of AEA and 2-AG is largely dependent on a ‘‘metabolic control,’’ which modulates the effects of these substances by modulating their in vivo concentration (or endogenous tone). Therefore, the main metabolic routes that allow synthesis, transport, and degradation of AEA and 2-AG will be reviewed in the next section, in order to put in a better perspective their implications for the biological activity of endocannabinoids at their receptors within the CNS.

2

Metabolism of AEA and 2-AG

2.1 Synthesis of AEA Unlike classical neurotransmitters and neuropeptides, AEA is not stored in intracellular compartments, but is produced ‘‘on demand’’ by receptor-stimulated cleavage of lipid precursors. There is now a general consensus that AEA is generated mainly by a transacylase-phosphodiesterase-mediated synthesis, starting from membrane phosphoglycerides and phosphatidylethanolamine. The main features of this biosynthetic pathway, schematically depicted in > Figure 16-2, have been recently reviewed (Ligresti et al., 2005). The AEA precursor N-arachidonoylphosphatidylethanolamine (NArPE) is believed to originate from the transfer of arachidonic acid from the sn-1 position of 1,2-sn-di-arachidonoyl-phosphatidylcholine to phosphatidylethanolamine, catalyzed by a calcium-dependent N-acyltransferase (trans-acylase, NAT). NArPE is then cleaved by a recently characterized N-acylphosphatidylethanolamine (NAPE)-specific phospholipase D (NAPE-PLD), which releases AEA and phosphatidic acid (> Figure 16-2). Alternative Biosynthetic pathways for AEA have recently energed, and are the subject of intense investigation (Lic et al., 2006; Simon and Cravatt, 2006).

2.1.1 NAPE-PLD The complementary DNAs of this enzyme have been cloned from mouse, rat, and human, and the deduced amino acid sequences were found to be composed of 393 (human) or 396 (mouse and rat) residues (Okamoto et al., 2004). The calculated molecular masses were 45.8 (mouse), 45.7 (rat), and 45.6kDa (human), with an amino acid identity of 95.5% (between mouse and rat), 90.4% (between rat and human), and 89.1% (between mouse and human). Interestingly, the sequences showed no homology with those of the reported phospholipases, and suggested that NAPE-PLD belongs to the zinc metallo-hydrolase family of the b-lactamase fold (Okamoto et al., 2004). This is a large superfamily of zinc metallohydrolases, which show a highly conserved motif presumably needed for zinc coordination (Okamoto et al., 2004). Assays of enzyme activity, messenger RNA and protein levels have demonstrated that NAPE-PLD is widely distributed in murine organs, with higher contents in the brain, kidney, and testis. A remarkable age-dependent increase of NAPEPLD expression has been recently shown in rat CNS, suggesting a central role in brain development (Morishita et al., 2005). Also of interest seems the observation that decreased NAPE-PLD in the uterus is instrumental to keep low the endogenous AEA levels, thus allowing blastocyst implantation and survival (Guo et al., 2005).

2.2 Degradation of AEA The biological activity of AEA at CB receptors is terminated by its removal from the extracellular space, which occurs through cellular uptake by a high-affinity transporter (AEA membrane transporter, AMT), followed by intracellular degradation by fatty acid amide hydrolase (FAAH).

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. Figure 16-2 Metabolism of AEA. Anandamide (AEA) is synthesized from phosphoglycerides and phosphatidylethanolamine via a calcium-dependent trans-acylase (NAT) activity. The resulting N-arachidonoyl-phosphatidyl ethanolamine (NArPE) is then cleaved by a calcium-dependent NAPE (N-acyl-phosphatidylethanolamine)-specific phospholipase D (NAPE-PLD), which releases AEA from membrane lipids. Finally, this endocannabinoid is hydrolyzed by fatty acid amide hydrolase (FAAH) into arachidonic acid and ethanolamine

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2.2.1 AMT A hot spot in endocannabinoid research is the mechanism by which AEA is taken up by cells. In fact, it is clear that AEA uptake has the features of a facilitated transport (i.e., it is dependent on concentration, time and temperature, and independent of external Na+ ions or ATP hydrolysis), yet the molecular identity of, and the gene encoding for, an AEA membrane transporter still remain elusive (Glaser et al., 2003, 2005; Hillard and Jarrhaian, 2003; Battista et al., 2005). In particular, the relationship between AMT and FAAH is still under debate, because FAAH might not quite need a transporter to get in contact with AEA (Bracey et al., 2002), and AMT might export (rather than import) AEA across the plasma membrane (Maccarrone et al., 2002a). However, recent evidence based on pharmacological, biochemical, and morphological data seem to favor the existence of an authentic AEA transporter distinct from the AEA hydrolase (Ligresti et al., 2004; Oddi et al., 2005). Moreover, the development of new drugs able to inhibit selectively AMT (reviewed by Ortega-Gutie´rrez, 2005) corroborates this concept.

2.2.2 FAAH Once taken up by cells, AEA is substrate for the enzyme fatty acid amide hydrolase (N-arachidonoylethanolamine amidohydrolase, EC 3.5.1.4; FAAH), which breaks the amide bond and releases arachidonic acid and ethanolamine (reviewed by McKinney and Cravatt, 2005). Studies of FAAH knockout mice have confirmed that this enzyme is a key regulator of in vivo signaling of AEA and related fatty acid amides. In fact, FAAH (/) animals have 15-fold higher levels of AEA than wild-type littermates (Cravatt et al., 2001). These observations have boosted research aimed at finding inhibitors of FAAH activity, as new ‘‘magic bullets’’ for a number of human diseases (reviewed by Maccarrone, 2006). Mammalian FAAH is a membrane-bound enzyme with a globular shape, and in all species the monomer has a molecular mass of 60–65 kDa and 579 aminoacids, of which 73% are identical. The biological unit of FAAH appears as a homodimer bound to membrane lipids via a-helices 18 and 19 (McKinney and Cravatt, 2005). In total, FAAH has 28 a-helices and 11 b-sheets, which account for 53 and 13% of the whole protein structure (Bracey et al., 2002), and contains an ‘‘amidase signature’’ sequence. In addition, the recently resolved x-ray crystal structures of FAAH have revealed an unusual serine–serine–lysine (S241-S217-K142) catalytic triad, along with a remarkable collection of channels that form a ‘‘cytosolic port’’ and a ‘‘membrane port’’: these ports might grant the simultaneous access to both the membrane and cytosolic compartments of the cell, useful for substrate entry and/or product exit during the catalytic reaction (Bracey et al., 2002). In this context, it seems noteworthy that a recent quantitative structure–activity relationship (QSAR) analysis of some natural hydroxy-derivatives of AEA or 2AG has shown that molecules able to interact with the cytosolic port of FAAH might be more suitable for the development of reversible inhibitors of enzyme activity, compared with hydrophobic compounds that interact with the membrane port (Dainese et al., 2005). On the other hand, the analysis of the FAAH promoter has revealed interesting features (reviewed by Puffenbarger, 2005), which might lead to the development of enzyme activators, rather than inhibitors. In fact, the promoter of human FAAH gene has a binding site for the transcription factor Ikaros, which in lymphocytes enhances FAAH gene expression in response to progesterone (Maccarrone et al., 2003a). Also leptin, a cytokine that controls immunity and reproduction in humans, is able to enhance FAAH gene transcription in lymphocytes, through a STAT3 (signal transducer and activator of transcription 3)-mediated up-regulation of the promoter (Maccarrone et al., 2003a). Therefore, drugs able to mimic the actions of Ikaros and STAT3 might open the avenue to the development of FAAH activators of potential therapeutic value, for instance in the treatment of female (Maccarrone and Finazzi-Agro`, 2004) and male (Schuel and Burkman, 2005) infertility. Yet, neuronal FAAH promoter, that does contain Ikaros and STAT3 binding sites, seems insensitive to progesterone and leptin, suggesting that FAAH expression is under cell-specific control (Gasperi et al., 2005). Additionally, inhibitors or activators may become useful tools to modulate the ability of FAAH to convert in vivo common drugs into bioactive compounds, as recently demonstrated for the conversion of the analgesic and antipyretic agent paracetamol into a potent vanilloid receptor agonist (Hogestatt et al., 2005). Recently a new FAAH-2

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has been characterized, that shows a lower selectivity than FAAH towards AEA and night contribute to endocannabinoid catabolism in vivo (Wei et al.)

2.2.3 NAAA Recently, a novel N-acylethanolamine-hydrolyzing acid amidase (NAAA), active only at acidic pH, has been cloned from human, rat, and mouse (Tsuboi et al., 2005). The primary structures deduced from complementary DNAs revealed that NAAA has no homology to FAAH, but rather belongs to the choloylglycine hydrolase family. In addition, NAAA was distinguishable from FAAH not only for the low pH optimum (5 vs. 8–9), but also for the preference of PEA compared with other NAEs, the activation by nonionic detergents, the lower sensitivity toward powerful inhibitors of FAAH and the lysosome localization within the cell (Tsuboi et al., 2005). Furthermore, the deduced amino acid sequence unveiled similarity of NAAA to acid ceramidase, an amidohydrolase that hydrolyzes ceramide to sphingosine and fatty acid. NAAA is composed of 359 (human) or 362 (rat and mouse) residues, with a molecular mass of 40 kDa, and shows 90.1% (between rat and mouse), 76.7% (between human and mouse), or 76.5% (between human and rat) amino acid identity (Tsuboi et al., 2005). Interestingly, there are six potential N-glycosylation sites in human, rat, and mouse sequences, and in fact NAAA is a glycoprotein, and glycosylation is essential for the full enzyme activity (Tsuboi et al., 2005). The organ distribution of NAAA messenger RNA revealed its wide distribution with the highest expression in lung. Yet, the physiological meaning of this acidic NAEhydrolase remains to be clarified, and further analysis is deemed necessary before NAAA can find its correct place among the proteins that metabolize endocannabinoids.

2.3 Synthesis of 2-AG Like anandamide, 2-AG is not stored in intracellular compartments, but it is produced on demand by receptor-stimulated cleavage of lipid precursors (reviewed by Sugiura and Waku, 2000; Ligresti et al., 2005). A biosynthetic pathway provides for quick hydrolysis of inositol phospholipids by a specific phospholipase C (PLC) for the generation of diacylglycerol (> Figure 16-3). In turn, this is converted to 2-AG by a sn-1-diacylglycerol lipase (DAGL). Another pathway for 2-AG formation involves the hydrolysis of phospholipids by phospholipase A1 (PLA1) into lysophospholipids, which are hydrolyzed by a lyso-PLC to produce 2-AG (> Figure 16-3).

2.3.1 DAGL Two sn-1-specific DAG lipases (DAGL) responsible for the synthesis of 2-AG have been recently cloned, by comparing human genome with Penicillium DAGL sequence (Bisogno et al., 2003). These two isoforms (a and b) have molecular masses of 120 and 70 kDa, respectively, and four transmembrane domains. They are members of the serine-lipase family, with serine and aspartatic acid (S443 and D495) participating in the catalytic triad. In addition, the a isoform is predominant in the adult brain, whereas the b isoform is expressed in the developing brain (Bisogno et al., 2003). Most recently, activation of group I metabotropic glutamate receptors (mGluR) has been shown to trigger the biosynthesis of 2-AG, but not AEA, in corticostriatal primary cultures and hippocampal slices, by enhancing DAGL (Jung et al., 2005). This interesting observation, points to a role for 2-AG as a primary endocannabinoid mediator of mGluRdependent neuronal plasticity, which is critical for synaptic transmission (Freund et al., 2003).

2.4 Degradation of 2-AG Like for AEA, the biological activity of 2-AG depends on its life span in the extracellular space, which in turn is limited by a rapid transport through the membrane, and subsequent intracellular hydrolysis.

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. Figure 16-3 Metabolism of 2-AG. Phospholipase C (PLC) releases diacylglycerol from membrane phospholipids. Diacylglycerol is then cleaved by diacylglycerol lipase (DAGL), which produces 2-arachidonoyl-glycerol (2-AG). Alternatively, phospholipase A1 (PLA1) releases from membrane phospholipids an sn-1 lysophospholipid, which is cleaved by lyso-PLC in order to generate 2-AG. Finally, this endocannabinoid is hydrolyzed by monoacylglycerol lipase (MAGL) into arachidonic acid and glycerol

2.4.1 2-AG Transporter It has been proposed that the 2-AG membrane transporter is the same as AMT (Beltramo and Piomelli, 2000; Bisogno et al., 2001). In fact, 2-AG accumulation is reduced by AM404 (N-(4-hydroxyphenyl)arachidonoylamide), an AMT inhibitor, and indirectly by high concentrations of arachidonic acid

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(Beltramo and Piomelli, 2000). The effect of AM404 is due to the inhibition of AMT and not to that of FAAH activity, because using two strong FAAH inhibitors, like URB597 and AM374, the concentration of 2-AG remained unaltered (Hajos et al., 2004).

2.4.2 MAGL Once accumulated in the cell, 2-AG can be degradated by FAAH (Goparaju et al., 1999). However, FAAH is not the main enzyme responsible for the metabolism of 2-AG, because FAAH (/) mice, which are unable to metabolize AEA, still have the ability to hydrolyze 2-AG (Lichman et al., 2002). The enzyme responsible for 2-AG degradation, monoacylglycerol lipase (MAGL), has been isolated from porcine brain (Goparaju et al., 1999), cloned and characterized in rat (Dinh et al., 2002) and human brain (Ho et al., 2002). Rat brain MAGL is a 33-kDa protein, showing 92% homology with mouse adipocyte MAGL (Karlsson et al., 1997). It has the catalytic triad serine-aspartic acid-histidine (S122-D239-H269) commonly found in lipases (Karlsson et al., 1997). Unlike FAAH, MAGL is localized in the cytosol and is discretely distributed throughout the CNS (Dinh et al., 2002). In general, it should be kept in mind that critical activities of 2-AG independent of those of AEA are emerging both in the CNS (Melis et al., 2004), and in the periphery (Oka et al., 2005), therefore the understanding of MAGL and DAGL regulation and of their role in maintaining the endocannabinoid tone in vivo can be of utmost importance, as it has been the case for NAPE-PLD (Okamoto et al., 2004) and FAAH (McKinney and Cravatt, 2005) with respect to AEA. In this line, recently a selective inhibitor of MAGL has been synthesized, which has identified this enzyme as a key modulator of endocannabinoid signaling within the hippocampus (Makara et al., 2005), and thus as an important target for drug development.

2.5 Transport Across the Blood–brain Barrier AEA has many neurovascular activities (for a comprehensive review see Battista et al., 2004); however it is not yet clear how AEA can be metabolized at the neurovascular interface, and how it can move through the vascular and the cerebral compartments. In order to shed some light on this important issue, isolated bovine brain microvessels (BBM) have been recently used as an ex vivo model of the blood–brain barrier (Maccarrone et al., 2006). BBM have been found to contain detectable levels of AEA and to possess the biochemical machinery to bind and metabolize it, namely CB1 and CB2 receptors, AMT, FAAH, and NAPEPLD. In addition, activation of CB1R was found to enhance AMT activity of BBM, through increased nitric oxide synthase (NOS) activity and subsequent release of NO, thus extending to an ex vivo model previous observations in human umbilical vein endothelial cells (HUVEC) in vitro (Maccarrone et al., 2000b). Instead, AMT activity was reduced by activation of CB2R, which inhibits NOS and NO release. Of major interest was the observation, based on functional assays and electronmicroscopic analysis, that different endothelial cells express differently CB1R and CB2R on their luminal and/or abluminal sides, and that such a different localization can lead to a different regulation of AMT activity on the luminal and abluminal membranes (Maccarrone et al., 2006). Taken together, these data suggest that CBR distribution may drive the directional transport of AEA through the blood–brain barrier and other endothelial cells. In fact, the CB1R-AMT coupling might represent a ‘‘trigger’’ for a fast AEA uptake through the transporter, whereas the CB2R-AMT coupling would represent a ‘‘brake’’ of the same process. Therefore, the presence of CB1R on one side (e.g., luminal) and that of CB2R on the other side (e.g., abluminal) of the barrier could drive AEA transport in the luminal ! abluminal direction, while preventing endocannabinoid movement in the opposite direction. If AMT is also a 2-AG transporter (see previous section), the same mechanism might direct the flow of 2-AG across the blood–brain barrier. Our experiments failed to substantiate this rather appealing working hypothesis for BBM, which did not show an asymmetric distribution of CB1R and CB2R. However, such an asymmetry did occur in calf pulmonary artery endothelial (CPAE) cells, where CB1R was found to be predominantly expressed on the luminal side membranes, whereas CB2R was almost exclusively expressed on the abluminal side (Maccarrone et al., 2006). In addition, our electronmicroscopic analysis

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showed that rat brain endothelial (RBE) cells failed to show any asymmetry in the distribution of their CBRs, as they did with AMT activity, and that HUVEC had only CB1 receptors symmetrically distributed on both sides of the cell (Maccarrone et al., 2006). Overall, these data support the hypothesis that the different functional link between AMT and CB1R or CB2R, and a different distribution of these receptors on the luminal and abluminal membranes, could generate a driving force for the directional transport of AEA through the endothelial cell layers of different districts.

3

Molecular Targets and Signaling Pathways

3.1 Cannabinoid Receptors As mentioned before, AEA and 2-AG bind to and activate type-1 (CB1R; 2.1:CBD:1:CB1:) and type-2 (CB2R; 2.1:CBD:2:CB2:) cannabinoid receptors (Howlett et al., 2002, 2004; Pertwee and Ross, 2002). CB1R is localized mainly in the CNS (Egertova et al., 2003), but is also expressed in peripheral tissues like heart, uterus, testis, liver and small intestine, and in immune cells (Maccarrone et al., 2001; Nong et al., 2001; Klein et al., 2003). Conversely CB2R is predominantly expressed peripherically, but is also present in the brain (Nunez et al., 2004; Van Sickle et al., 2005). In addition to CB1R and CB2R, AEA and 2-AG activate non-CB1/non-CB2 cannabinoid receptors also, which are still under characterization (Drmota et al., 2004; Howlett et al. 2004). Both CB1R and CB2R belong to the class A rhodopsin family of G protein-coupled seven trans-membrane spanning receptors (GPCR). They show 44% overall identity with 68% identity within the transmembrane regions, and are coupled mainly to the Gi/o family of G proteins (Howlett et al., 2004). The main signal transduction pathways triggered by CBRs are summarized in > Table 16-1, and

. Table 16-1 Main signaling pathways triggered by type-1 or type-2 cannabinoid (CB1/2) receptors, and by vanilloid receptors (TRPV1) CB1/2 receptors Inhibition of adenylyl cyclase Activation of mitogen-activated protein kinase Inhibition of voltage-gated L, N, and P/Q-type Ca2+ channels Activation of K+ channels Activation of focal adhesion kinase Activation of cytosolic phospholipase A2 Activation (CB1R) or inhibition (CB2R) of nitric oxide synthase

TRPV1 receptors Activation of nonselective ion channels Activation of protein kinases Increase in intracellular Ca2+ concentration Opening of intracellular Ca2+ stores Dissipation of mitochondrial membrane potential (mitochondrial uncoupling) Cytochrome c release from mitochondria Activation of caspases

include inhibition of adenylyl cyclase (AC), activation of mitogen-activated protein kinase (MAPK), regulation of ionic currents (inhibition of voltage-gated L, N and P/Q-type Ca2+ channels, and activation of K+ channels), activation of focal adhesion kinase (FAK) and of cytosolic phospholipase A2, and activation (by CB1R) or inhibition (by CB2R) of nitric oxide synthase (Howlett et al., 2002, 2004). Evidence has emerged that in addition to CB1R and CB2R there are other CB receptors through which the endocannabinoids might induce a biological activity, like a purported CB3 (or GPR55) receptor (Breivogel et al., 2001; Drmota et al., 2004). Another molecular target of AEA, but not 2-AG, which has attracted great interest, is the type-1 vanilloid receptor (now called transient receptor potential vanilloid 1, TRPV1).

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3.2 Vanilloid Receptors TRPV1 is a six trans-membrane spanning protein with intracellular N- and C-terminals and a pore-loop between the fifth and sixth transmembrane helices (Jung et al., 1999). TRPV1 is a ligand-gated and nonselective cationic channel, activated by molecules derived from plants, such as the pungent component of ‘‘hot’’ red peppers capsaicin, and also by noxious stimuli like heat and protons (Jung et al., 1999). In the last few years, a number of studies have suggested a physiological role for AEA as TRPV1 agonist, leading to the concept that AEA, besides being an endocannabinoid, is also a true ‘‘endovanilloid’’ (De Petrocellis et al., 2001; Van der Stelt and Di Marzo, 2004). Instead, 2-AG is unable to bind to and activate TRPV1 (Van der Stelt and Di Marzo, 2004). The interaction of AEA with TRPV1 occurs at a cytosolic binding side (De Petrocellis et al., 2001; Jordt and Julius, 2002), and triggers the intracellular responses listed in > Table 16-1. TRPV1 is expressed in peripheral sensory fibers and also in several nuclei of the CNS (Marinelli et al., 2003), thus suggesting the existence of brain endogenous agonists for its activation. In this context, recently an endogenous capsaicin-like substance has been identified, N-arachidonoyl-dopamine (NADA; > Figure 16-1), which activates TRPV1 with high potency and is also a potent cannabimimetic compound (Huang et al., 2002). Therefore, there seems to be quite an expanded overlap between the endogenous cannabinoid system and the vanilloid system. Overall, it can be stated that virtually all central and peripheral systems of mammals are affected by endocannabinoids, from CNS (Iversen, 2003; Freund et al., 2003; Piomelli, 2003; Di Marzo et al., 2004; Fowler, 2005), to immune system (Parolaro et al., 2002; Klein et al., 2003), digestive tract (Di Carlo and Izzo, 2003), and reproductive organs (Schuel and Burkman, 2005). Taken together AEA, 2-AG, and congeners, along with CB and non-CB receptors, synthetic and hydrolytic enzymes, and membrane transporters form the ‘‘endocannabinoid system,’’ which is schematically depicted in > Figure 16-4. This system is subject of intense investigation, also in the light of its possible therapeutic exploitation, for the treatment of neurodegenerative diseases (Piomelli, 2003), cancer (Bifulco and Di Marzo, 2002; Guzman, 2003), anxiety (Kathuria et al., 2003), eating disorders (Di Marzo et al., 2004), human infertility (Maccarrone and Finazzi-Agro`, 2004), and pain (Hohmann et al., 2005). Very recently, evidence has been accumulated that suggests a role for lipid rafts in modulating the endocannabinoid system.

4

Modulation by Lipid Rafts

Lipid rafts are subdomains of the plasma membrane that contain high concentrations of cholesterol and glycosphingolipids, and are well-known modulators of the activity of a number of GPCR (KunzelmanMarche et al., 2002; Simons and Ehehalt, 2002; Pike, 2003). As a consequence, they modulate signaling and membrane trafficking in many cell types (Gajate and Mollinedo, 2005; Le Roy and Wrana, 2005; Rodgers et al., 2005). Not surprisingly lipid rafts have been proposed as a potential regulator of CBR activity (Hinz et al., 2004; Howlett et al., 2004; Barnett-Norris et al., 2005). In this line, it has been shown that in rat C6 glioma cells raft perturbation by cholesterol depletion enhanced CB1R binding and signaling (Bari et al., 2005a), whereas raft perturbation by membrane cholesterol enrichment had the opposite effect (Bari et al., 2005b). In these studies methyl-b-cyclodextrin (MCD), a membrane cholesterol depletor that is widely used to disrupt the integrity of lipid rafts (Kunzelman-Marche et al., 2002; Pike, 2003), was found to double CB1R binding and subsequent signaling via AC and MAPK activity (Bari et al., 2005a), which instead were halved by membrane cholesterol enrichment (Bari et al., 2005b). In addition, two parallel studies have shown that cholesterol depletion by MCD reduces also the activity of AMT, possibly by promoting a faster endocytosis of the transporter molecules (McFarland et al., 2004; Bari et al., 2005a), overall suggesting that lipid rafts might modulate the endocannabinoid signaling (reviewed by McFarland and Barker, 2005). Of major interest was also the observation that CB1R activation after MCD treatment could account for the ability of this raft disruptor to block apoptosis induced in vitro by AEA in the same C6 cells (Sarker and Maruyama, 2003; Bari et al., 2005a), a biological activity of AEA that will be discussed in further detail later in this chapter. As mentioned above, the cannabinoid receptor subtypes, CB1 and CB2, have been classified into the class A rhodopsin-like family of GPCR (Howlett et al., 2002, 2004; Pertwee and Ross, 2002). In the absence

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. Figure 16-4 The endocannabinoid system. The intracellular localization of the proteins that bind, transport, synthesize, and degrade AEA or 2-AG is shown. Note that AMT works bidirectionally, moving AEA in and out across the plasma membrane, and that it might transport 2-AG also. In addition, note that AEA but not 2-AG binds TRPV1 receptors, which show an intracellular binding site. Instead CBRs, which are activated by both AEA and 2-AG, have an extracellular binding site. AEA, anandamide; 2-AG, 2-arachidonoylglycerol; AMT, AEA membrane transporter; CB1/2R, type 1/2 cannabinoid receptor; DAG, diacylglycerol; DAGL, diacylglycerol lipase; FAAH, fatty acid amide hydrolase; MAGL, monoacylglycerol lipase; NArPE, N-arachidonoyl-phosphatidylethanolamine; NAT, N-acyltransferase; TRPV1, transient receptor potential channel vanilloid receptor subunit 1

of crystals, studies have been carried out to understand the three-dimensional structure of CB receptors and their mechanisms of action by using computer molecular modeling and NMR approaches. Recently, a solid-state NMR study has shown that AEA undergoes a fast lateral diffusion within the bilayer outer leaflet before making a productive interaction with CB1R (Tian et al., 2005), giving ground to the concept that the membrane environment is critical for CB1R binding and signaling. The molecular basis of the sensitivity of CB1R to raft integrity might be complex, and need a thorough analysis of the lipid environment of the receptor, along with the characterization of its three-dimensional structure in the context of membrane bilayers. In this line, it seems noteworthy that it has been supposed that in the membrane outer leaflet AEA takes an extended conformation that enables it to interact with a hydrophobic groove formed by helices 3 and 6 of CB1R, where its terminal carbon is positioned close to a key cysteine residue (C47) in helix 6 (Tian et al., 2005). The C47-helix 6 interaction has been shown to be essential for receptor activation (Tian et al., 2005). In addition, a recent study using combined high-resolution NMR and computer modeling has shown that CB1 receptors have peculiar conformational properties and salt bridges in the so-called juxtamembrane segment (or helix 8), which is critical for their activity and regulation (Xie and Chen, 2005). Notably, helix 8 is under the influence of the surrounding chemical environment (Xie and

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Chen, 2005). Therefore, it is tempting to suggest that lipid rafts might regulate CB1R, by interacting with specific regions of its three-dimensional structure, like helices 3 and 6, or helix 8, thus modulating receptor folding within the bilayer. These studies on the role of lipid rafts on CB receptors have been extended to the CB2 subtype, which turned out to be insensitive to raft integrity (Bari et al., 2006). Since CB1 and CB2 receptors are both central and peripheral, are both activated by AEA and 2-AG, and both trigger common signaling pathways (> Table 16-1), exists it seens of utmost importance to that a possible differential regulation of CB1R and CB2R, also in view of the fact that these two subtypes have been recognized as distinct drug discovery targets for numerous potential therapeutic applications.

5

Biological Activities within the CNS

The endocannabinoid system serves important functions in the regulation of brain networks and synaptic transmission, which has been the subject of in-depth reviews (Freund et al., 2003; Iversen, 2003; Piomelli, 2003; De Petrocellis et al., 2004; Carrier et al., 2005; Paradisi et al., 2006). > Table 16-2 summarizes some

. Table 16-2 Biological actions of AEA and congeners in different brain areas Thalamus, hypothalamus, hippocampus Control of pain initiation Control of wake/sleep cycles Control of thermogenesis Control of appetite Impairment of working memory Impairment of memory consolidation Inhibition of long term potentiation Inhibition of glutamatergic transmission Basal ganglia, striatum, globus pallidus Control of psychomotor disorders Interference with dopaminergic transmission Inhibition of GABA (g-aminobutyric acid)-ergic transmission Induction of long term depression Potentiation of GABA (g-aminobutyric acid)-mediated catalepsy Cortex, cerebellum, spinal cord Blockade of NMDA (N-methyl-D-aspartate) receptors Control of tremor and spasticity Retina Control of scotopic vision

activities of AEA and 2-AG in different brain areas, an issue that is further expanded in two independent chapters of this book. Here, it seems important to recall a general activity of endocannabinoids in controlling brain synaptic transmission, i.e., their ability to inhibit for several seconds neurotransmitter release from presynaptic terminals, thus providing a physiological feedback mechanism for neurons to selfregulate the strength of their synaptic inputs (for a review, see Alger, 2002). This ‘‘retrograde signaling’’ of endocannabinoids (Wilson and Nicoll, 2002) results in depolarization-induced suppression of inhibition (DSI) at GABA (g-aminobutyric acid)-ergic synapses, and in depolarization-induced suppression of excitation (DSE) at glutamatergic synapses. In both cases, depolarization of a postsynaptic neuron induces the transient suppression of neurotransmitter release from presynaptic nerve terminals impinging

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on that neuron (Alger, 2002; Wilson and Nicoll, 2002). In addition to the regional actions shown in > Table 16-2, and to the general retrograde activity leading to DSI or DSE, endocannabinoid signaling shows two features broadly relevant for the CNS: (i) interaction with steroids, and (ii) regulation of cell survival or death.

5.1 Interaction with Steroids The endocannabinoid system has been shown to interact with a number of neurotransmitters within the CNS (Fride, 2002; Piomelli, 2003), and with various signaling molecules in the periphery (Parolaro et al., 2002). Growing evidence suggests that these cross-talks are based also on the interaction of endocannabinoids with steroids, another group of well-known biologically active lipids (Sheppard, 2003; Casserly and Topol, 2004). As a matter of fact, such an interaction has been proposed soon after the discovery of AEA, in a report describing the effect of this endocannabinoid on the hypothalamic-pituitary-adrenal (HPA) axis (Weidenfeld et al., 1994), and recently has found a strong biochemical background (Di et al., 2003; Wenger et al., 2003). Since then, cross-talks between endocannabinoids and steroids have been shown to regulate several central and peripheral functions, due to a direct effect of steroids on the proteins of the endocannabinoid system. > Table 16-3 shows that estrogen (17b-estradiol, E) can down-regulate CB1 expression at

. Table 16-3 Modulation of the proteins of the endocannabinoid system (ES) by steroids Cell type Rat pituitary gland Human endothelial cells Human endothelial cells Human endothelial cells Mouse uterus Mouse uterus Human T cells

ES protein CB1 ↓ AMT ↑ NAPE-PLD ↑ FAAH ↓a FAAH ↓ FAAH ↓ FAAH ↑b

Steroid Estrogen Estrogen Estrogen Estrogen Estrogen Progesterone Progesterone

CB1, type-1 cannabinoid receptor; AMT, anandamide membrane transporter; NAPE-PLD, N-acyl-phosphatidylethanolaminespecific phospholipase D; FAAH, fatty acid amide hydrolase a Nongenomic regulation b Genomic regulation

the mRNA level in the anterior pituitary gland of rats (Gonzalez et al., 2000), whereas it up-regulates AMT in human umbilical vein endothelial cells (HUVEC) in vitro (Maccarrone et al., 2002a). Additionally, in HUVECs estrogen also up-regulates NAPE-PLD and down-regulates FAAH through a rapid, nongenomic mechanism mediated by a surface estrogen receptor (ER) (Maccarrone et al., 2002a). Furthermore, E and progesterone (P) down-regulate FAAH activity in mouse uterus, while P genomically activates the same enzyme in human T lymphocytes, by enhancing FAAH promoter activity and subsequent gene expression (Maccarrone et al., 2003a). Within the CNS, glucocorticoids secreted by the adrenal cortex can exert inhibitory effects on several hypothalamic neuroendocrine systems, including negative feedback regulation of the HPA axis. Classical glucocorticoid actions, like those of other steroid hormones, are mediated by binding to intracellular receptors and regulation of gene transcription (Sheppard, 2003; Casserly and Topol, 2004). However, recent evidence suggests that in different species rapid glucocorticoid actions are mediated by membrane receptors and activation of nongenomic signaling mechanisms (Falkenstein et al., 2000). The molecular identity of glucocorticoid membrane receptors is still elusive, and it remains controversial how glucocorticoids mediate their rapid feedback regulation within the HPA axis. Yet, a recent report has given interesting

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clues on this issue, revealing an intriguing interplay between endocannabinoids and glucocorticoids (Di et al., 2003). In fact, it has been demonstrated that glucocorticoids exert a rapid inhibitory effect on glutamate release onto identified parvocellular neurons of the paraventricular nucleus (PVN), by activating a G-protein-dependent signaling mechanism that leads to the synthesis and retrograde release of an endocannabinoid messenger. This endocannabinoid binds to and activates G-protein-coupled CB1 receptors on presynaptic glutamate terminals, leading to the suppression of glutamate release onto the PVN neurons, and hence causing decreased neuronal activity within the PVN (Di et al., 2003). Taken together, these data demonstrate a mechanism of rapid glucocorticoid feedback inhibition of hypothalamic function via endocannabinoid release in the PVN, providing a molecular link between the actions of glucocorticoids and endocannabinoids that regulate stress, energy homeostasis, and feeding disorders. Such a glucocorticoid–endocannabinoid interface via activation of G-protein-dependent signaling pathways represents a novel mechanism of rapid steroid action in the brain. In addition, the endocannabinoid link between steroid hormonal and neuronal signaling expands the recently discovered role of endocannabinoids as retrograde modulators of synaptic function (Alger, 2002; Wilson and Nicoll, 2002), discussed above.

5.2 Control of Cell Death The anti-proliferative properties of THC have been known for decades (Parolaro et al., 2002), and also the ability of AEA to control cell proliferation and death has been the focus of intense research in the last few years (reviewed by Guzman, 2003; Maccarrone and Finazzi-Agro`, 2003b). In several models, AEA but not 2-AG has shown an anti-proliferative action due to the induction of apoptosis, which occurs through the activation of different receptors (> Table 16-4).

. Table 16-4 Nerve cells forced to apoptosis by (endo)cannabinoids, and receptors involved Cellular model Rat hippocampal neurons Rat glioma C6 cells in vitro

Rat glioma C6 cells in vivo Human astrocytoma U373MG cells

Rat pheochromocytoma PC12 cells Human neuroblastoma CHP100 cells Rat astrocytes

Receptor involved (role of activation) CB1 (pro-apoptotic) CB1 (pro-apoptotic) CB1 (anti-apoptotic), TRPV1 (pro-apoptotic) CB1 (weak pro-apoptotic), TRPV1 (proapoptotic) CB1 (pro-apoptotic), SBS (pro-apoptotic) CB2 (pro-apoptotic) CB1 (pro-apoptotic)

Reference (first report) Chan et al. 1998 Galve-Roperh et al., 2000 Maccarrone et al., 2000a Jacobsson et al., 2001

CB1 (anti-apoptotic) CB1 (pro-apoptotic)

Go´mez del Pulgar et al., 2000 Sarker et al., 2000

TRPV1 (pro-apoptotic)

Maccarrone et al., 2000a

CB1 (anti-apoptotic)

Sa`nchez et al., 2001b

Maccarrone et al., 2002b Sa`nchez et al., 2001a Rueda et al., 2000

CB1/2, type 1/2 cannabinoid receptor; TRPV1, transient receptor potential channel vanilloid receptor subunit 1; SBS, SEA (N-stearoylethanolamine)-binding site

On the one hand, programmed death of glioma cells in vitro has been shown to involve activation of CB1R followed by ceramide accumulation and Raf1/ERK (extracellular signal-regulated kinase) activation (Galve-Roperh et al., 2000); on the other hand, the activation of CB2R seems the critical event leading to

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inhibition of glioma growth in vivo (Sanchez et al., 2001a). In rat cortical astrocytes and human astrocytoma cells, AEA activates CB1 receptors leading to sphingomyelin breakdown through the adaptor protein FAN, suggesting a CB1 receptor-mediated pro-apoptotic signaling independent of Gi/o proteins (Sa`nchez et al., 2001b). In the same cells, CB1 receptor activation also leads to long-term activation of c-Jun N-terminal kinase (JNK) and p38 mitogen-activated protein kinase (MAPK), suggesting that a threshold might exist above which endocannabinoid-induced JNK and p38 MAPK activation would lead to apoptosis (Rueda et al., 2000). More recently, the JNK/p38 MAPK signaling cascade triggered by AEA has been shown to depend on the apoptosis signal-regulating kinase 1 (ASK1), a key player in the control of cell death (Sarker et al., 2003). In general it may be speculated that AEA binding to CB1 receptors modulates through ASK1 the balance among ERK, JNK, and p38 MAPK, thus regulating the cell choice between proliferation and death. In addition to the modulation of ceramide degradation through neutral sphingomyelinase (Galve-Roperh et al., 2000) and of ceramide synthesis through serine palmitoyltransferase (Go´mez del Pulgar et al., 2002a), it has been shown that cannabinoids are able to modulate, again through CB1 receptors, the phosphatidylinositol 3-kinase/protein kinase B (PI3K/PKB) pathway, which serves as a pivotal anti-apoptotic signal (Go´mez del Pulgar et al., 2000, 2002b). This finding is of particular interest, because it points toward a protective role of cannabinoid receptors against programmed cell death. This concept was originally proposed in neuroblastoma cells, where instead TRPV1 activation was shown to trigger apoptosis (Maccarrone et al., 2000a). Then, the anti-apoptotic role of CBR has found further grounds in astrocytoma cells (Galve-Roperh et al., 2002) and HUVECs, and in the latter cells TRPV1 was shown again to trigger apoptosis (Yamaji et al., 2003). In general, it seems noteworthy that AEA can exert opposite effects, i.e., a pro-apoptotic activity at vanilloid receptors and an anti-apoptotic action at cannabinoid receptors, but it remains to be clarified if the different localization of the binding sites of these receptors (intracellular for TRPV1 and extracellular for CBRs; see > Figure 16-4) may play a role in discriminating this dual action. It is tempting to speculate that modulation of intracellular and extracellular levels of AEA through fine tuning of the activity of FAAH (and possibly of AMT) is the ‘‘checkpoint’’ of this regulation, in line with the general consensus that FAAH controls the endogenous tone of AEA in vivo (Cravatt et al., 2001; McKinney and Cravatt, 2005). It seems also of interest the finding that activation of CB1R prevents apoptosis, and that lipid rafts, by controlling binding and signaling of this receptor, can modulate survival and death of neuronal cells (Sarker and Maruyama, 2003; Bari et al., 2005a, 2005b).

6

Conclusions and Future Avenues

A role for the endogenous cannabinoid system is emerging in regulating several aspects of human (patho) physiology, due to the activation of cannabinoid and vanilloid receptors, and/or to non-receptor mediated actions. In the case of AEA, the control of its cellular activity seems to be largely dependent on FAAH, rather than on NAT, NAPE-PLD, or AMT. Here, I have described the endocannabinoid system, in order to put in a better perspective its biological activity within the CNS, and the potential therapeutic applications of endocannabinoid-oriented drugs. In particular, I have presented and discussed new advances in this field, like transport across the blood–brain barrier, regulation of endocannabinoid signaling by lipid rafts, interaction with steroids, and regulation of cell survival or death. Overall, it seems that modulating endocannabinoid metabolism, rather than agonizing or antagonizing cannabinoid and noncannabinoid receptors, might be the way to better understand the pathophysiological implications of these bioactive lipids, and to exploit them for therapeutic purposes. In this context, I suggest that not only inhibitors of FAAH, but also drugs able to enhance its activity might become useful therapeutic tools for the treatment of human diseases. If not as therapeutic agents per se, FAAH inhibitors or activators could be used together with AEA analogues to lower the doses or to shorten the treatment necessary in vivo to observe an effect, and hence to minimize the possible psychotropic side effects of these substances when they are used as pharmaceuticals. On a final note, it seems necessary to remind the following: (i) several endogenous endocannabinoid(-like) compounds, whose functions are not yet understood, are present in our body, and their biological activity might be affected in unexpected ways by drugs that modulate FAAH or

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other known proteins of the endocannabinoid system; (ii) new metabolic enzymes have been recently identified, that catalyze the hydrolysis and synthesis of AEA and 2-AG, or the hydrolysis of PEA, and it remains to be elucidated how these pathways may contribute to the overall tone and biological activity of endocannabinoids.

Acknowledgments I wish to thank Prof. A. Finazzi-Agro` (Department of Experimental Medicine and Biochemical Sciences, University of Rome ‘‘Tor Vergata’’) for continuing interest and support, and all colleagues who gave their valuable contribution over the years to the studies on the endocannabinoid system in the CNS. I also thank Dr. A. Paradisi for the excellent production of the artwork. This investigation was supported by Ministero dell’Istruzione, dell’Universita` e della Ricerca (COFIN 2002 and 2003), and by Fondazione TERCAS (Research Programs 2004 and 2005).

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Simons K, Ehehalt R. 2002. Cholesterol, lipid rafts, and disease. J Clin Invest 110: 597-603. Simon GM, Cravatt BF. 2006. Endocannabinoid biosynthesis proceeding through glycerophospho-N-acyl ethanolamine and a role for alpha/beta-hydrolase 4 in this pathway. J Biol Chem 281: 26465-26472. Sugiura T, Waku K. 2000. 2‐Arachidonoylglycerol and the cannabinoid receptors. Chem Phys Lipids 108: 89-95. Tian X, Guo J, Yao F, Yang DP, Makriyannis A. 2005. The conformation, location, and dynamic properties of the endocannabinoid ligand anandamide in a membrane bilayer. J Biol Chem 280: 29788-29795. Tsuboi K, Sun YX, Okamoto Y, Araki N, Tonai T, et al. 2005. Molecular characterization of N‐acylethanolamine‐ hydrolyzing acid amidase, a novel member of the choloylglycine hydrolase family with structural and functional similarity to acid ceramidase. J Biol Chem 280: 11082-11092. Stelt Van der M, Di Marzo V. 2004. Endovanilloids. Putative endogenous ligands of transient receptor potential vanilloid 1 channels. Eur J Biochem 271: 1827-1834. Sickle Van MD, Duncan M, Kingsley PJ, Mouihate A, Urbani P, et al. 2005. Identification and functional characterization of brainstem cannabinoid CB2 receptors. Science 310: 329332. Weidenfeld J, Feldman S, Mechoulam R. 1994. Effect of the brain constituent anandamide, a cannabinoid receptor agonist, on the hypothalamo‐pituitary‐adrenal axis in the rat. Neuroendocrinol 59: 110-112. Wenger T, Ledent C, Tramu G. 2003. The endogenous cannabinoid, anandamide, activates the hypothalamo‐pituitary‐ adrenal axis in CB1 cannabinoid receptor knockout mice. Neuroendocrinol 78: 294-300. Wilson RI, Nicoll RA. 2002. Endocannabinoid signaling in the brain. Science 296: 678-682.

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Section 4

Diet, Brain Lipids and Brain Functions

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Diet, Brain Lipids, and Brain Functions: Polyunsaturated Fatty Acids, Mainly Omega‐3 Fatty Acids

J. M. Bourre

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 411

2 2.1 2.2 2.3 2.4 2.5 2.6 2.7 2.8

Brain Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 413 ALA: Animal Experiments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 413 ALA: Trials on Newborn Infants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 415 DHA and EPA: Animal Experiments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 416 EPA and DHA Alone in Baby Formulas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 417 DHA and EPA: Trials on Newborn Infants, the Importance of ARA and Breast‐Feeding . . . . . . . 417 Specific Actions on Sensory Organs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 419 Secondary Benefits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 420 The Special Case of Trans Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 420

3 3.1 3.2 3.3 3.4 3.5 3.6 3.7 3.8

Psychiatric Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421 Mood and Pain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421 Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421 Dyslexia in Young and Adult Subjects and Hyperactive Children . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 422 Drug Addiction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 422 Depression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 422 Dementias and Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 424 Schizophrenia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 425 Autism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 425

4 Aging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 426 4.1 Biochemical Changes and Diet . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 426 4.2 Cognitive Alterations During Aging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 426 5

Miscellaneous: Neurological Disorders and Handicaps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427

6

Desaturases and the Relationship between Omega‐3 Fatty Acids and Antioxidants . . . . . . . . . . . 428

7

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 428

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_17, # Springer ScienceþBusiness Media, LLC 2009

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Diet, brain lipids, and brain functions: polyunsaturated fatty acids, mainly omega‐3 fatty acids

Abstract: Fatty acids are directly involved in the structure of most lipids, including those in the nervous system, providing their chemical and biological characteristics. Besides saturated and monounsaturated fatty acids, two families of polyunsaturated fatty acids, the omega‐6 fatty acids, such as linoleic acid (LA), and the omega‐3 fatty acids, such as alpha linolenic acid (ALA), are most important. LA and ALA are essential nutrients, as the human body cannot synthesize them or convert one to the other. They were formerly called vitamin F, before their chemical structures were determined. The term ‘‘omega‐3 fatty acids’’ is plural as there are four main ones that have increasing numbers of double bonds and carbon atoms. ALA (18:3 o3) is the precursor of stearidonic acid (18:4 o3, SA), which gives rise to eicosapentaenoic acid (EPA, 20:5 o3) and docosapentaenoic acid (DHA, 22:6 o3). The omega‐3 polyunsaturated fatty acid content of the brain is extremely high, indicating that these fats are involved in brain physic chemistry, biochemistry, physiology and function; and, consequently, in brain development, in some neuropsychiatric diseases and in the cognitive decline of aging. This review examines all three aspects Some studies on perinatal cerebral development have focused on ALA, whereas others have examined long‐chain derivatives, DHA and, to a lesser extent EPA. A third group of studies has examined the influence of ALA and DHA, sometimes with the omega‐6 fatty acid, arachidonic acid (ARA). The studies on ALA provided the first demonstration of the effect of a dietary component on the structure and function of the brain that involved several scientific disciplines. These included cultures of dissociated brain cells; analyses of the fatty acids and lipids and cell types in the brain; regions and classes of phospholipids; physicochemical studies on brain membrane fluidity; biochemical and enzymological studies on enzymes such as ATPase; physiological studies on dopaminergic, serotoninergic, and cholinergic neurotransmission; toxicology of heavy metals and trans fatty acids; studies on vision, hearing, and taste; electrophysiological studies (ERG and EEG); and cognitive and behavioral studies, memory and habituation being specifically affected. The accumulation of considerable experimental evidence led to the inclusion of ALA in baby formulas. This decision has been confirmed by many studies on newborns. The nature of the polyunsaturated fatty acids (particularly the omega‐3 fatty acids) in baby formulas for both full term and premature infants influences the infant’s visual, neurological, cerebral, and intellectual capacities. Enrichment with long‐chain fatty acids such as DHA (and EPA) is based on the fact that human milk contains them, unlike animal milks, that the brain is extremely rich in these fatty acids, and there is little desaturase activity (which, together with elongases, transforms ALA to DHA). Clinical studies on EPA and DHA, both found in fish oils, preceded or paralleled those on animals, in contrast to the work on ALA. Despite a lack of exhaustive experiments, DHA and EPA were added to baby formulas, which may well have a limited or even negative effect because of competition with omega‐6 fatty acids. Formula supplemented with DHA and ARA has a positive effect on membranes, including nerve membranes, on physiological, electrophysiological, and sensory parameters, particularly vision and, most recently, on hearing. However, there is still debate about their influence on motor function, neurological capacity, behavior, and cognition. In fact, both DHA and ALA are essential fatty acids because of the low desaturase activities in the brain and liver. Dietary omega‐3 fatty acids from oily fish in the human diet (the efficiency of fish oil in capsule has not yet been clearly demonstrated) may help prevent some psychiatric disorders, mainly depression and dementia, particularly Alzheimer’s disease. The direct role of omega‐3 fatty acids in bipolar disorder (manic‐depressive disease) and schizophrenia has not yet been clearly established, although suspected in some aspects of these diseases. Omega‐3 fatty acids may also be involved in dyslexia and autism, but there is evidence that they appear to help prevent stress. Finally, their influence on mood and libido is a matter for discussion pending experimental proof in animals and humans. A lack of dietary omega‐3 fatty acid can prevent the renewal of membranes, and thus accelerate cerebral aging, but the roles of the vascular system (where the omega‐3 fatty acids are active) and the cerebral parenchyma itself have not yet been clearly resolved. The insufficient dietary supply of omega‐3 fatty acids (mainly ALA) in today’s occidental diet (and in a number of diets throughout the world) raises the problem of how to correct dietary habits so that the consumer selects foods that are rich in omega‐3 fatty acids; mainly rapeseed (canola) and walnut oils, oily fish, and certain eggs. Alpha tocopherol protects the omega‐3 fatty acids in the nervous system membranes and not other components of vitamin E.

Diet, brain lipids, and brain functions: polyunsaturated fatty acids, mainly omega‐3 fatty acids

17

List of Abbreviations: ALA, alpha‐linolenic acid; SA, stearidonic acid; EPA, eicosapentaenoic acid, also named timnodonic acid; DHA, docosahexaenoic acid also named cervonic acid; LA, linoleic acid; ARA, arachidonic acid

1

Introduction

The brain has a higher lipid content than any other body organ, except for adipose tissue. All its lipids, which are mostly phospholipids, are found in cell membranes, and are almost never sources of energy. Position 2 of the glycerol molecules in phospholipids generally bears a polyunsaturated fatty acid such as docosahexaenoic acid (DHA; 22:6 (n 3), 22:6o3), or arachidonic acid (ARA; 20:4(n 6), 20:4o6). There may well be smaller amounts of adrenic acid (22:4o6) and eicosapentaenoic acid (EPA; 20:5(n 3), 20:5o3). The brain contains very little alpha linolenic acid (ALA; 18:3(n 3), 18:3o3), although this is the precursor of all the other omega‐3 fatty acids, or linoleic acid (LA; 18:2(n 6), 18:2o6), the precursor of all the other omega‐6 fatty acids, mainly ARA. The families of fatty acids are shown in > Figure 17‐1. Brain structures also contain omega‐6 fatty acids, but the main feature of the omega‐3 (o3), (n 3)) and omega‐6 (o6, (n 6)) fatty acids is that they are essential and strictly complementary, while competing for the desaturases. The human diet usually contains enough omega‐6 fatty acid, but insufficient amounts of ALA and DHA; this is why dietary lipids and the nervous system focus on omega‐3 fatty acids. High concentrations of saturated and monounsaturated fatty acids are also present in the nervous system, but they appear to be endogenous rather than from the diet, with certain possible exceptions. Consequently, their origin and roles are rapidly examined in this review. The importance of the omega‐3 fatty acids for brain development is based on three findings. First, the brain is exceptionally rich in polyunsaturated fatty acids, including omega‐3 fatty acids (Svennerholm and Vanier, 1973). They account for 15–20% of cerebral fatty acids, and for as much as 40% in the neurons and nerve terminals (Bourre et al., 1984). Second, a lack of dietary ALA results in a reduced cerebral DHA content, which is offset by an increase in 22:6 omega‐6 (Galli et al., 1972; Bourre et al., 1984). Finally, human mother’s milk is particularly rich in ALA, but the cow’s milk used to prepare baby formula in the early eighties is not. This was because producers in many countries began to use sunflower oil (devoid of omega‐3 fatty acids) rather than rapeseed oil (canola, containing 9% ALA) in their formulations. Human fetuses and newborns accumulate considerable quantities of omega‐3 fatty acids, mainly DHA (Clandinin et al., 1980a, b; Martinez, 1992; Makrides et al., 1994; Cunnane et al., 2000). It was initially shown that the differentiation (Bourre et al., 1983) of brain cells in culture required omega‐3 fatty acids and omega‐6 fatty acids. A lack of dietary ALA disturbs the composition of brain cell membranes (Bourre et al., 1984). Chemical, physicochemical, biochemical, and enzymological analyses, as well as toxicological, physiological, electrophysiological and behavioral studies on ALA provided the first experimental evidence (on the same animals) that a dietary component could influence the structure and function of the brain (Bourre et al., 1989a). Studies on rats showed a dose–effect relationship between the amount of ALA in a mother’s diet and its accumulation in the pup’s brain (Bourre et al., 1989b). It was subsequently shown that the omega‐3 fatty acids in modified baby formula influence the visual and cerebral (including intellectual) capacities of the child, as measured by its neurodevelopment, intellectual quotient, and motor index (Uauy et al., 2003). Hence, all baby formulas now contain at least ALA, in quantities equivalent to those in mother’s milk. The main relevance of the omega‐3 fatty acids to health is in the prevention and treatment of cardiovascular disease. However, the omega‐3 fatty acids are also valuable because they are indispensable for the construction, maintenance, and function of the brain. Thus the omega‐3 fatty acids are major brain components, important for higher functions (Bourre, 2004a, b). It is hardly surprising, then, that psychiatrists are interested in omega‐3 fatty acids. The high content of these fatty acids in the brain is one reason. Another is the direct consequence of studies on experimental animals showing that a lack of dietary ALA results in behavioral and cognitive defects, particularly problems of learning (Bourre et al., 1989a; Yamamoto et al., 1991; Lim and Suzuki, 2001; Salem et al., 2001;

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Diet, brain lipids, and brain functions: polyunsaturated fatty acids, mainly omega‐3 fatty acids

. Figure 17‐1 Omega‐3 fatty acids. Fatty acids are directly involved in the structure of most lipids, providing their chemical and biological characteristics and identity. A lipid is polyunsaturated if a major fraction of its fatty acids is polyunsaturated. There are two families of polyunsaturated fatty acids, the omega‐6 fatty acids, such as linoleic acid (LA; C18, 18:2 v6), and the omega‐3 fatty acids, such as alpha linolenic acid (ALA—also C18, but with a different number and distribution of double bonds, 18:3 v3). LA and ALA are essential nutrients as the human body cannot synthesize them or convert one to the other. They were both collectively formerly designed as vitamin F, before their chemical structure had been determined. The term ‘‘omega‐3 fatty acids’’ is plural as there are four main ones that have increasing numbers of double bonds and carbon atoms. Alpha linolenic acid, with three double bonds 18:3 v3, (ALA), is the precursor of stearidonic acid (18 carbon atoms, C18, and four double bonds, 18:4 v3, SA), which gives rise to eicosapentaenoic acid (EPA, C20, five double bonds, 20:5 v3; found in tuna fish). All omega‐3 fatty acids have their first double bond on C3, counting from the methylated, biochemically inert end (‘‘omega’’ as it is at the far end from the chemically reactive part of the molecule). This can also be written as 18:3 (n 3), but the term ‘‘omega’’ is widely used. A lipid is said to be saturated when most of its fatty acids are saturated; the main saturated fatty acids are palmitic (C16) and stearic (C18) acids, the main fatty acid in myelin is lignoceric acid (C24). A lipid is deemed to be monounsaturated if it contains mainly monounsaturated fatty acids such as oleic acid (C18, one double bond, 18:1 omega‐9). Myelin has a high content of nervonic acid (C24:1)

Wainwright, 2002), memory and habits (Frances et al.,1996a, 2000a) and reactions to morphine (Frances et al., 1996b). A lack of ALA results in the abnormal metabolism of neuromediators (Chalon et al., 2001; Kodas et al., 2002a). Several of these problems can be overcome by feeding an appropriate diet (Carrie et al., 2000a, b, 2002; Ikemoto et al., 2001: Kodas et al., 2002b). DHA is absolutely essential for the development of the human brain (Crawford et al., 2001). This review focuses on fatty acids and the influence of nutrition on their actions. Consequently, lipid signaling (Galli and Petroni, 1990), although extremely important (Chen and Bazan, 2005), and the regulation of synaptic function and dysfunction by lipids (Bazan 2005) are outside its scope. DHA is the precursor of neuroprotectin (Bazan, 2006), as docosanoids have been detected in the nervous system (Bazan, 1989; Marcheselli et al., 2003).

Diet, brain lipids, and brain functions: polyunsaturated fatty acids, mainly omega‐3 fatty acids

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Brain Development

2.1 ALA: Animal Experiments Most studies have used diets containing ALA from rapeseed oil (about 10% ALA), others have used soybean oil (8% ALA) or linseed oil (25–55% ALA, depending on the variety); Japanese studies have used perilla oil, but walnut oil (‘‘noix de Grenoble’’ oil, 11% ALA) and hemp oil have not been used. The usual models are newborn rats and mice, but some work has also used monkeys, hamsters, and pigs. The mothers of the newborn rats or mice were often fed a diet lacking the appropriated element for 2–3 weeks before mating; other studies have used the offspring of two generations of deficient parents. The perinatal period studied includes gestation–lactation, as glial cell development is greatest during the first 5 days postpartum, and myelination occurs around day 18. The great lack of DHA in animals fed a diet lacking ALA (made with sunflower or peanut oil) is generally offset by an excess of docosapentaenoic acid (22:5 6) in the brain (Galli et al., 1972; Tinoco et al., 1978); and in a variety of its cells, including the neurons, astrocytes, and oligodendrocytes and organelles such as the myelin and nerve terminals (Bourre et al., 1984). The total quantity of polyunsaturated fatty acid is thus almost unchanged, as are those of monounsaturated and saturated fatty acids. Dietary ALA is conserved, and its very long‐chain derivatives are reused; a 21‐fold reduction in dietary ALA results in only a fivefold reduction in DHA concentration in the organs, and only a twofold drop in DHA in neurons (Bourre et al., 1984; > Figure 17‐2). This preservation of DHA is owing to recycling triggered by deacylation and reacylation, which are reduced only by 30–70%, whereas the transfer between the blood and the brain is reduced 40‐fold (Contreras et al., 2000). This may be why there is only a 50% drop in the DHA in neurons (Bourre et al., 1984). Analysis of the fatty acid composition of total phospholipids in 11 regions of the brains of mice has shown that the DHA concentration is normally highest in the frontal cortex (> Figure 17‐3). A lack of

. Figure 17‐2 Effect of a lack of dietary ALA and DHA during pregnancy and lactation on the cells and organelles of the nervous system and other organs of rat pups. The experimental diet contained 3% ALA. From Bourre et al. (1989a)

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. Figure 17‐3 DHA and 22: 5(n 6) in regions of the brains of mice fed with an ALA‐deficient diet. From Carrie et al., (2000a)

dietary ALA does not alter the concentrations of DHA in all the brain regions to the same extent. The hypophysis, frontal cortex, and striatum are most severely affected (Carrie et al., 2000a), whereas the loss of DHA from the hippocampus is associated with the appearance of smaller neurons (Ahmad et al., 2002), although their number is unchanged, and there is a quantitative loss of phosphatidylserine (Murthy et al., 2002). There are also changes in the relative proportions and compositions of several phospholipids. For example, there is an increase in the DHA in phosphatidylethanolamine in the gray matter of monkeys, where it replaces ARA (Kanazawa et al., 1995). A lack of dietary ALA alters the phospholipids, melatonin, and lipoxygenase products in the pineal gland (Gazzah et al., 1993; Zaouali‐Ajina et al., 1999; Zhang et al., 1998). Thus, a lack of ALA leads to reduced phospholipid synthesis (Gazzah et al., 1995) and distorts protein turnover by increasing their synthesis (Giaume et al., 1994). There is a dose–effect relationship between the amount of ALA in the diet during pregnancy and lactation and the DHA content of the brains of the offspring (Bourre et al., 1989a), which persists when they become adults (Bourre et al., 1993). Dietary ALA must provide a minimum of 0.4% of an animal’s caloric intake (the French recommended daily intake is 0.8% of calories, as the brain weight/body weight ratio is very high in humans). The DHA concentration in cells and subcellular structures, including brain microvessels, recovers very slowly after a lack of dietary ALA (Youyou et al., 1986; Bourre et al., 1989a; Moriguchi et al., 2001; Moriguchi and Salem 2003), even though these blood vessels are in intimate contact with plasma lipoproteins having a normal composition because the liver rapidly recovers its capacity to synthesize them (Homayoun et al., 1988). In contrast, a single injection of ALA into the amnion of a pregnant rat lacking dietary omega‐3 fatty acids will correct the lack of omega‐3 fatty acids in the brains of the fetuses within 24 h (Schiefermeier and Yavin, 2002). A lack of dietary ALA disturbs the fluidity of nerve terminal membranes because of a lack of DHA and an increase in 22:5 omega‐6 fatty acids (Zerouga et al., 1991), resulting in lower activities of ATPase and 50 nucleotidase in the endoplasmic reticulum and of myelin CNPase; some microsomal (Barzanti et al., 1990), mitochondrial (Barzanti et al., 1994), and nuclear (Ammouche et al., 1994) enzymes are also affected.

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The DHA content of the membranes of various tissues is correlated with its ATPase activity (Turner et al., 2003). Animals fed a diet lacking ALA have only 50% of the normal ATPase activity in their nerve terminals (Bourre et al., 1989a) and the activities of ATPase isoforms in the brain are altered (Gerbi et al., 1993). Such alterations probably lead to biochemical and functional abnormalities, at least because Na‐K‐ATPase produces 60% of the brain energy, equivalent to 10% of an adult’s dietary carbohydrate intake, and this figure is probably much higher in newborns. Furthermore, a lack of omega‐3 fatty acids can alter energy metabolism in the brain by upsetting glucose transport (Ximenes et al., 2002), undoubtedly linked to the altered endothelial cells of the brain microvessels (Homayoun et al., 1988). A lack of dietary ALA does not severely alter neuromuscular function (Frances et al., 1995), but it affects learning (Bourre et al., 1989a) and behavior. These changes are best revealed by tests like the cross labyrinth, Morris swimming pool, high and low rotating rod, and ‘‘shuttle box’’ tests (Wainwright et al., 1994; Frances et al., 1996a). Those activities associated with memory and habituations are particularly affected (Frances et al., 1996b, 2000a). These findings have been confirmed in many studies (Coscina et al., 1986; Yamamoto et al., 1988; Carlson 2000; Salem et al., 2001; Wainwright, 2002). The suggested relationship between the changes in learning performance and the composition of the microsomal membrane (Yoshida et al., 1997) seems to correlate well with the cognitive and behavioral deficits seen in rats fed a low‐ALA diet and the defects in their monoaminergic nerve transmission, particularly in the frontal cortex (Chalon et al., 2001; Levant et al., 2004). However, neither the density nor the functions of dopamine transporters are affected (Kodas et al., 2002a). In agreement with fatty acid composition of nerve cells (Bourre et al., 1989a, b), dopaminergic neurotransmission is only partially restored in the frontal cortex and other areas by restoring dietary ALA (Kodas et al., 2002b). Thus the slow incorporation of fatty acids into the brain is correlated with a slow improvement in learning performance (Lim and Suzuki, 2001). There is a clear relationship between polyunsaturated fatty acids and neurotransmission, and hence between them and behavior (Chalon et al., 2001; Takeuchi et al., 2002). Both cholinergic and serotoninergic neurotransmissions (Aid et al., 2003; Kodas et al., 2004) are also influenced by a lack of dietary ALA. A PET scan study on conscious monkeys showed that the modulation of cholinergic nerve transmission by DHA involves cerebral blood flow, in addition to brain structures themselves (Tsukada et al., 2000). A lack of dietary ALA also results in abnormal electroencephalograms (Takeuchi et al., 2002). Finally, a reduced amount of DHA in the brain makes the cell membranes of an animal more sensitive to toxic agents (Bourre et al., 1989a) and to trans fatty acids, as described later. The studies quoted in this review concern only the results obtained in mammals; studies on birds and fish have not been cited, although their brain development has been monitored (Cherian and Sim, 1991; Henderson et al., 1996; Ajuyah et al., 2003). A diet rich in omega‐3 fatty acids improves the development of chickens and the production of eggs that are nutritionally valuable to humans (Simopoulos and Salem, 1992); their lipids are even used to prepare baby formula rich in ALA, DHA, and ARA (Chirouze et al., 1994; Koletzko et al., 1995; Decsi et al., 1995).

2.2 ALA: Trials on Newborn Infants Holman first reported treating a little girl suffering from neurological signs due to a lack of ALA with dietary ALA (Holman, 1986). These findings were confirmed by studies on another girl aged 7 years who was treated with linseed oil (55% ALA) and cod liver oil (Bjerve et al., 1988). Later studies were done on adults and older people. The importance of dietary ALA for the growth and development of the brain and retina, particularly in premature babies, has been emphasized since 1992 (Uauy et al., 1992). Most workers in the area now agree that the erythrocyte and blood plasma phospholipid fatty acid profiles are a reflection of that of the brain. Cholesterol esters are also important indicators of the fatty acid status of a newborn baby (Babin et al., 2000). The objective of the dietary intervention was thus to improve the blood plasma omega‐3 fatty acid status using baby formulas enriched with rapeseed oil (Crastes de Paulet et al., 1994; Billeaud et al., 1997). The effect depends on the amount of ALA in the formula, any increase in ALA content corrects the lack of omega‐3 fatty acids in the newborn plasma and erythrocytes (> Figure 17‐4) (Chirouze et al., 1994). Baby formula enriched in ALA can correct the abnormalities seen in the electroretinograms (ERGs)

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. Figure 17‐4 Effect of 2 ALA concentrations in milk on the omega‐3 fatty acids in phospholipids and erythrocytes of newborn infants. From Chirouze et al. (1994)

of premature babies (Uauy et al., 1992); and ensure the better development of the brains of full‐term, normal birth‐weight babies aged up to 1 year (Voigt et al., 2002) (> Table 17‐1). The LA/ALA ratio must not be less than 4 (Clark et al., 1992).

2.3 DHA and EPA: Animal Experiments ALA alone does not seem to be sufficient to ensure that the brain has an optimal concentration of DHA (Bowena and Clandinin, 2000), although an adequate equilibrium between LA and ALA could ensure the optimal synthesis of DHA (Cheon et al., 2000). Moreover, ALA did not contribute appreciably to DHA within the brains of adult rats fed a diet enriched in DHA (Demar et al., 2005). The authors proposed using a transgenic rapeseed oil (canola oil) with a high gamma linolenic acid (GLA) content (containing both 18:3 omega‐3 fatty acid and 18:3 omega‐6 fatty acid) to maintain the ratio between the two families of fatty acids (Wainwright et al., 2003), the equivalent could be a mixture of rapeseed (canola) and borage oils. A given amount of dietary DHA is more effective than the same amount of ALA (Abedin et al., 1999). This is logical as DHA is derived from ALA, and the diet normally contains much more ALA than DHA. Moreover, there is an uneven regional distribution of fatty acids in the brain, so that the impact of a lack of dietary ALA is region‐specific (Carrie et al., 2000a, b). A diet containing egg phospholipids or pig brain extract (containing DHA and ARA) is more effective than triglycerides containing ALA at restoring the fatty acids in all tissues of an animal lacking ALA, except the frontal cortex (Bourre and Dumont, 2002). Fish oil controls the activity of certain isoforms of ATPase (Gerbi et al., 1994), whereas a mixture of DHA and ARA increases the ATPase activity in nerve terminals (Bowen and Clandinin, 2002). The brain seems to use preformed very long‐chain omega‐3 polyunsaturated (and omega‐6) fatty acids most effectively under normal circumstances (Cunnane et al., 2001), which implies that they are synthesized in the liver from ALA (and LA) when they are not obtained directly from the diet. DHA injected into the blood stream is taken up by the brain (Bazan and Scott, 1990). Supplementing the diet of animals with DHA (and EPA) improves certain aspects of their cognitive performance (Carlson, 2000). This led to a comparison of diets without ALA and those containing either ALA (rapeseed and linseed oils) or DHA (seaweed extracts, fish oils, and purified products), or ALA and DHA (algae, brain or egg extracts, mixtures of vegetable, and fish oils).

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. Table 17‐1 Effect of ALA concentration on blood parameters and mental performance of newborn infants ALA in milk Plasma phospholipid ALA Erythrocyte phospholipid ALA Erythrocyte phospholipid DHA Bayley PDI Bayley MDI

0.40 0.10 0.05 0.83 88.80 94.80

0.95 0.16 0.09 1.55 95.50 96.30

1.70 0.23 0.12 1.72 93.70 101.00

3.20 0.46 0.23 2.52 96.40 100.50

Note: MDI, mental developmental index; PI, psychomotor developmental index. From Voigt et al. (2002)

Diets containing only very long‐chain omega‐3 fatty acids (EPA and DHA) are not satisfactory: a diet rich in fish oil alters the composition of brain membranes by increasing the DHA content but considerably reducing the ARA content (Bourre et al., 1990). This favors arousal and learning in young mice, but reduces motor activity and learning in older mice (Carrie et al., 2000c). The omega‐6 fatty acids must therefore be in balance and accompanied by omega‐3 fatty acids. Consequently, a diet supplemented with both DHA and ARA prevents the decrease in dopaminergic and serotoninergic neurotransmitters in the frontal cortex of piglets caused by a lack of ALA (de la Presa and Innis, 1999). Tests on the motor and neurological development of monkeys (Champoux et al., 2002) have given rise to discussions about the amounts of DHA that should be added to baby formulas (Sarkadi‐Nagy et al., 2004).

2.4 EPA and DHA Alone in Baby Formulas DHA is clearly required by babies, including full‐term infants (Gibson et al., 1996). For example, analysis of the frontal cortex of dead infants has shown that babies fed with formula lacking DHA have a lower‐than‐ normal DHA content (Makrides et al., 1994; Byard et al., 1995). The brain of a child takes up 48% of the 10mg of DHA incorporated each day. The brains of breast‐fed babies (mother’s milk provides about 60 mg/ day) contain about 1g DHA, whereas the brains of bottle‐fed babies contain only 0.6 g DHA (Cunnane et al., 2000). Fetal accretion of omega‐3 fatty acids was estimated at 67 mg/day during the third trimester (Clandinin et al., 1980a, b) In practice, a combination of ALA and DHA should be taken into account when preparing formulas for premature babies (Rodriguez et al., 2003), although there has yet to be a clear demonstration of the absolute need for EPA and DHA (Simmer and Patole, 2004). The negative effect of a lack of omega‐3 fatty acids on the development of the nervous system (and vision) led initially to the addition of omega‐3 fatty acids alone (both EPA and DHA, no omega‐6 fatty acids), generally as fish oil. This improved visual acuity (Carlson et al., 1993), but this type of supplementation does not favor the overall development of the infant, in terms of body weight, height, and head circumference (Carlson et al., 1992). This is not a great surprise as fish oil capsules are known to reduce hypertriglyceridemia, which is probably not good for the lipid metabolism of newborn infants. A diet supplemented with an oil rich in DHA but poor in EPA for up to 4 months improves the DHA status, but may be detrimental for omega‐6 fatty acid status (Lapillonne et al., 2000a). However, a daily intake by a pregnant woman of 500–1000 mg of a DHA/EPA mixture, either as a milk supplement or as a capsule, improves the omega‐3 fatty acid status of the fetus without influencing its omega‐6 fatty acid status (Velzing‐Aarts et al., 2001).

2.5 DHA and EPA: Trials on Newborn Infants, the Importance of ARA and Breast‐Feeding Baby formula must contain both DHA and ARA to ensure an overall balance, and to maintain the omega‐3/ omega‐6 fatty acid ratio (Koletzko, 1992; Ghebremeskel et al., 1995; Makrides et al., 2000). A formula

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containing only ALA and LA, but no DHA or ARA, is inferior to mother’s milk, even for full‐term infants (Decsi et al., 1995). The fatty acid contents of erythrocyte membranes are an index of the nutritional status of brain tissue, and the DHA and ARA concentrations in these membranes drop after birth if a child is fed a formula containing only LA and ALA (Ghebremeskel et al., 2000), showing the importance of DHA and ARA. Breast‐fed babies have a better brain and body DHA status than those fed baby formula lacking DHA (Cunnane et al., 2000). Hence, formulas should be supplemented with both DHA and ARA (Lapillonne et al., 2000b). Premature babies have special dietary needs (Salle, 2002). A combination of fish oils and borage oil has been suggested for them to initiate omega‐6 fatty acid chains with gamma linolenic acid, which avoids the potentially negative effects of its precursor LA (Fewtrell et al., 2004). Extracts of eggs enriched in omega‐3 and omega‐6 fatty acids have been used in baby formulas for several years (Chirouze et al., 1994; Koletzko et al., 1995). Phospholipids extracted from pig brain have also been given to underweight infants (Ramirez et al., 2001). Although the biochemical and physicochemical results are quite conclusive, the behavioral results are somewhat less clear. An analysis of eight published trials carried out in 2001 (Simmer, 2001) was inconclusive. A second analysis of 14 published studies was carried out 3 years later; it compared baby formulas with and without long‐chain polyunsaturated fatty acids and was more positive. The 14 studies included 7 on the effects on cognitive development; the results took into account the fact that brain DHA is derived from both ALA (with a 10% conversion rate) and preformed DHA (Uauy et al., 2003). This speculated conversion rate is probably much too high, as discussed later. A study by Agostoni and group (1995) showed a benefit at 4 months, as assessed by the development quotient (Brunet‐Lezine test), but when these children were examined later, at the age of 4–2 years, there was no longer any difference (Agostoni et al., 1997; > Table 17‐2) . Another study found that adding ALA and DHA to the formula improved cognitive performance at 4 months and that the improvement persisted (Willatts et al., 1998; Willatts, 2002). Thus, an effect at 4 months does not predict an effect at 24 months. However, there is still a close correlation between the long‐chain fatty acids in erythrocytes and neurological development (Agostoni et al., 1997). The feeding habits of families undoubtedly vary considerably, so that the DHA status of an infant will be more influenced by the dietary pattern than by the nature of the baby formula/breast milk consumed during its initial months of life. This is probably why no difference was found in infants aged 39 months (Auestad et al., 2003), nor any association between the status at birth and performance at age 7 years (Bakker et al., 2003). Another study found a statistically insignificant improvement in children fed a formula containing DHA and ARA (Wezel‐Meijler van et al., 2002). A more detailed study, in which a restricted number of children were monitored from before birth, showed that very long‐ chain polyunsaturated fatty acids are closely associated with an increased IQ at the age of 6.5 years (Gustafsson et al., 2004). The children of women who take cod liver oil during pregnancy and lactation have improved mental performance measured at 4‐year old (Helland et al., 2003). Other authors (Birch et al., 2002) suggest that babies should be given supplemented formula for at least 6 weeks to ensure maturation of the cerebral cortex and optimal visual performance in later years. These studies used multiple tests, and it is quite possible that only some of them are influenced by dietary omega‐3 fatty acids, as has

. Table 17‐2 DHA in milk, and phospholipids of blood plasma and erythrocytes and the neurodevelopment quotient DHA in milk (g/100 g fat) DHA in plasma phospholipids (%) DHA in erythrocyte phospholipids (%) Neurodevelopment quotient

Formula 1 0.3 2.7 4.1 105

Formula 2 0 0.9 1.8 96

Mother’s milk 01–0.6 2.8 4.1 102

Note: From Agostoni et al. (1997, 1994). Formula 1: milk enriched in long‐chain fatty acids. Formula 2: standard milk. The ALA contents of the 3 milks was equivalent to 0.7 g/10g fat

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been shown in animal studies (Frances et al., 1996a, b). This could emphasize the advantage of breast‐ feeding over a formula containing DHA or ARA. Several studies have shown the importance of breast‐feeding and some have justified it by the amount of polyunsaturated fatty acids, particularly omega‐3 fatty acids, in mother’s milk. It improves the neurological development of newborns, especially if continued for over 6 weeks, as evaluated by motor activity (Bouwstra et al., 2003), and this effect can be measured in infants up to 1‐year old. Some believe that this is due to the presence of omega‐3 fatty acids, especially DHA (Agostoni et al., 2001b). Others have found that babies thus fed have an IQ 8.3 points higher at 18 months (Lucas et al., 1992). One study using the Bailey test showed that breast‐feeding had a positive effect, measured at 1 year, on baby boys, but not on girls (Paine et al., 1999). The effect on neurological development is measurable in children up to the age of 7 or 8 (Horwood et al., 2001) or even 9 years (Lanting et al., 1994). However, others have found that the effect cannot be measured at 13 months or 5 years (Angelsen et al., 2001). These widely differing results may be due to differences in tests used. Some studies indicate that a vegetarian diet is linked to an increased number of premature births and more frequent Cesarian sections. There is also evidence (Reddy et al., 1994) that pregnancy is shorter by 5.6 days and that the body weight and head circumference of the newborn are slightly reduced, once other factors such as the duration of pregnancy, the mother’s size, the number of children, and smoking are allowed for. These children also have a below‐normal arterial DHA content.

2.6 Specific Actions on Sensory Organs Omega‐3 fatty acids have a direct influence on vision as the retina has a very high DHA content (Anderson et al., 1992; Bazan and Rodriguez de Turco, 1994; Bazan et al., 1994). DHA is concentrated in the specialized membrane that makes up the photoreceptor outer segments. It contributes 35–65% of the fatty acids in the phosphatidylethanolamine and phosphatidylserine in the outer segment of photoreceptors in species ranging from frogs to humans. DHA occurs almost entirely in the sn‐2 position of the phospholipids and in retinal phosphatidylethanolamine it accounts for 75% of the fatty acid in this position. Approximately 60 molecules of phospholipid surround each molecule of visual pigment. Thus, the retinal photoreceptor membrane contains the body’s highest concentration of DHA. In fact, DHA is extremely important for vision, involving both the brain and the retina: photoreceptors, neurotransmission, rhodopsin activation, cone and rod development, neuron and synapse development, and maturation (Neuringer, 2000; Uauy et al., 2001). Rat that lacks ALA in its diet has an altered fatty acid distribution in the membranes of retina cells, and this alters the amplitude of the ‘‘a’’ and ‘‘b’’ peaks of the electroretinogram (Bourre et al., 1989a). The ‘‘a’’ wave is generated primarily by the photoreceptors, whereas the ‘‘b’’ wave, a later cornea‐positive component, is generated by the inner retina. Studies have measured the effects of omega‐3 fatty acids on the retina and vision in rats (Acar et al., 2002a), mice (Carrie et al., 2002), guinea pigs (Abedin et al., 1999; Weisinger et al., 1999), monkeys (Lin et al., 1994; Diau et al., 2003), and pigs (Hrboticky et al., 1991). Injected DHA is actively taken up by the retina (Bazan and Scott 1990). Supplementing the diet with phospholipids with a high DHA content (together with ARA) improves visual function in old mice, as it does in young mice lacking omega‐3 fatty acids (Carrie et al., 2002). An increase in plasma DHA parallels an increase in DHA in the retina, but the concentration of DHA in the retina remains unchanged when the plasma concentration rises above a given level. Lowering the LA/ALA ratio does not improve visual performance (Jensen et al., 1997), so this ratio alone is not a sufficient definition. This raises the question of whether DHA itself is essential during early childhood (Makrides et al., 1995). Electrophysiological studies on newborns have been used to measure the impact of dietary omega‐3 fatty acids. They confirmed the findings of animal studies. As stated earlier, the negative effect of a lack of omega‐3 fatty acids on the development of the nervous system (and vision) led initially to the addition of only omega‐3 fatty acids (both EPA and DHA, no omega‐6 fatty acids), generally as fish oil. This improved visual acuity (Carlson et al., 1992), but this type of supplementation does not favor the overall development of the infant, in terms of body weight, height, and head circumference (Carlson et al., 1992). However, fish oil

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increases the visual acuity of premature infants by improving the DHA status (Carlson et al., 1993). Hence vision is greatly improved by dietary omega‐3 fatty acids (Neuringer, 2000; Agostoni and Giovannini, 2001a; Neuringer and Jeffrey, 2003; Hall, 2003). Several of the studies cited earlier have also looked at t behavior. Similar results have been obtained for the effect of DHA on olfaction. Changes in behavioral tests based on this sense are not due to poorer olfaction, but due to altered brain structures (Catalan et al., 2002). The olfactory bulb is very sensitive to a lack of dietary ALA, as it leads to a considerable reduction in its DHA content (Greiner et al., 2001). A lack of omega‐3 fatty acids also affects taste. For example, animals lacking dietary ALA taste sweetness only in response to higher than normal sugar concentrations (Frances et al., 1996b). Hearing is also affected by a lack of omega‐3 fatty acids, especially the brain response. A lack of ALA also leads to accelerated or early aging of the auditory nervous system in rats (Bourre et al., 1999). The amount of DHA in the milk of lactating rats influences the auditory system of their offspring (Haubner et al., 2002). The auditory systems of breast‐fed infants or those fed formula supplemented with long‐chain polyunsaturated fatty acids for their initial 16 weeks of life mature more rapidly (Unay et al., 2004). This is probably important in terms of the chronology of organ development (Collet et al., 1988). Thus fatty acids are important for the responses of sensory receptors and even more so for the brain sensory systems (Bourre, 2004b). More or less half of the human cortex is involved in some aspects of visual processing. The cortex is dramatically affected by a lack of dietary ALA, as described earlier.

2.7 Secondary Benefits Children suffering from hyperphenylalaninemia absorb DHA and ARA poorly because of the dietary restrictions imposed by this disorder. DHA supplements improve their DHA status and their visual acuity (Agostoni et al., 2003b; Agostoni and Haschke, 2003a). Severe protein‐energy malnutrition (Kwashiorkor) is accompanied by a reduction in DHA, perhaps because of reduced elongase activity or increased peroxidation (Leichsenring et al., 1995). As a general rule, malnourished infants should be given dietary supplements of DHA and ARA (Marin et al., 2000). A restricted intrauterine blood supply alters the polyunsaturated fatty acid status of both experimental animals (Morand et al., 1981) and low‐birth‐weight infants (Cetin et al., 2002). This justifies supplementation, especially because of the long‐term consequences (Salle et al., 2001). Giving both DHA and ARA should improve the neurological and sensory status of low‐ birth‐weight infants (Crawford et al., 2003).

2.8 The Special Case of Trans Fatty Acids Trans monounsaturated fatty acids are mainly found in the diet and can be incorporated in nervous system membranes, including those of the peripheral nervous system (Bourre et al., 1982). The influence of trans monounsaturated fatty acids on brain composition and function remains open to question (Wauben et al., 2001). The trans polyunsaturated fatty acids derived from ALA are taken up by the brain (Grandgirard et al., 1994). Animal studies indicate that they reduce the concentrations of neurotransmitters in the developing brain (Acar et al., 2002b) and may alter the normal, optimal course of pregnancy (Carlson et al., 1997). They are present in human milk (Chardigny et al., 1995), where their concentration is inversely proportional to the amounts of omega‐3 and omega‐6 fatty acids (Innis and King, 1999). It is wise to limit their intake, although the brain seems to be better protected than other organs (Larque et al., 2001). The concentration of 22:5 omega‐6 fatty acids (from erythrocyte phosphatidyl ethanolamines) is not a completely relevant biochemical indicator of a lack of omega‐3 fatty acids, at least in infants less than 18 months old, as a high dietary intake of trans fatty acids or LA may inhibit the incorporation of DHA into membrane phospholipids (Innis et al., 2004). Although the prenatal neurological state of the mother/infant seems to be positively influenced by an optimal DHA, AA, and general polyunsaturated fatty acid status, the trans fatty acids (18C) and LA have negative effects (Dijck‐Brouwer et al., 2005). The role of conjugated linoleic acid (CLA), especially rumenic acid (cis‐9, trans‐11), is still a matter for speculation.

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Psychiatric Diseases

3.1 Mood and Pain There have been considerable speculation about the relationship between fatty acids (particularly omega‐3 fatty acids) and mood, but there have been few studies on the subject. An Australian study reported that regular normal meals (including omega‐3 fatty acids), including an adequate breakfast, improved mood and cognitive performance (Lombard, 2000). A study performed in New Zealand on subjects aged more than 15 years found that the personal perception of good mental and physical health varied with the consumption of fish, thus of omega‐3 fatty acids, so that they were considered to stabilize mood (Silvers and Scott, 2002). The following year, a study performed in England found that the consumption of fish did not improve mood in people who were not depressed (Ness et al., 2003). Thus these fatty acids are essential components of normal general health and well‐being, mainly because of their biochemical properties. One attempt to treat chronic fatigue gave inconclusive results (Warren et al., 1999). The food consumption and nutrient intake of subjects suffering from depression, anxiety, and insomnia were studied and these parameters were taken as indices of compromised mental well‐being. Those subjects suffering from anxiety or depression had higher intake of omega‐3 fatty acids and omega‐6 fatty acids (Hakkarainen et al., 2004a). It is thus not yet possible to propose that omega‐3 fatty acids modulate mood. Omega‐3 fatty acids could also be directly implicated in reducing the neuronal and glial processes that generate inflammatory pain (Shapiro, 2003). Some common oils can have significant analgesic effect in rats suffering from persistent pain produced by partial nerve injury. This effect may be associated with the amount of omega‐3 fatty acids consumed (Perez et al., 2005). Ibuprofen and omega‐3 fatty acids were found to have equivalents effects on reducing arthritic pain. Omega‐3 fatty acids from fish oil (1200 mg/day EPAþDHA) appear to be an alternative to NSAIDs for treating nonsurgical neck or back pain (Maroon and Bost, 2006).

3.2 Stress Some assays were first performed using ALA (in combination with LA) given as capsules prepared from vegetable oils. The optimum dietary ratio between omega‐3 and omega‐6 fatty acids (LA: ALA) to counteract stress is 4:1. This treatment is particularly good for protecting against changes in the hippocampus, for instance, in response to excess cortisol and corticosteroids (Yehuda et al., 2000). EPA has been used to treat women with personality disorders (Zanarini and Frankenburg, 2003). Aggressive behavior was found to vary inversely with the consumption of fish (18% reduction) (Iribarren et al., 2004). Daily quantities of 1.5–1.8 g DHA in fish oil helped to reduce stress (Hamazaki et al., 1996, 2000), and decreased the aggressive tendencies of young adults (possibly by modulating stress); augmenting dietary DHA intake could prevent extraggression, perhaps due to the prevention of coronary heart disease by fish oils (Hamazaki et al., 1996). Similar doses given to subjects aged between 50 and 60 years for 2 months reduced their aggressive behavior by 30%, but doses of 150 mg/day were without effect (Hamazaki et al., 2002). A dietary supplement of omega‐3 fatty acid (300 mg EPA and 200 mg DHA per capsule) given for 1 month increases sympathetic nerve activity responses to physiological stress in human: muscle nerve sympathetic activity was not changed at rest, but sympathetic outflow was increased by physiological stress (Monahan et al., 2004). Some results on the role of essential fatty acids in chronic fatigue syndrome contrast sharply with those of previous studies. These new studies found that 85% of patients had clinically significant improved symptoms after taking Efamol marine for 3 months (Warren et al., 1999). These patients were given two 500 mg Efamol Marine capsules 4 times a day; these capsules contained 36 mg GLA (gamma‐linolenic acid), 1mg EPA, 11mg DHA, and 255 mg LA. The basic daily food consumption and nutrient intake of subjects self‐reporting depression, anxiety or insomnia, and all three or none of the symptoms were supplemented (Hakkarainen et al., 2004a, b) (> Table 17‐3).

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. Table 17‐3 Baseline daily food consumption and nutrient intake of subjects self‐reporting depression, anxiety or insomnia, and all three or none of the symptoms

Fish (g) Omega‐3 fatty acids (total) (g) Omega‐3 fatty acids (from fish) (g) Omega‐3 fatty acids (from vegetables) (g)

Depressed mood (n ¼ 4314) 39.3 2.21

Anxiety (n ¼ 6498) 39.9 2.24

Insomnia (n ¼ 5550) 40.1 2.16

All three symptoms (n ¼ 1670) 40.3 2.23

No symptoms (n ¼ 19116) 39.3 2.14

0.47

0.48

0.48

0.49

0.46

1.77

1.79

1.70

1.77

1.70

Note: From Hakkarainen et al. (2004b)

3.3 Dyslexia in Young and Adult Subjects and Hyperactive Children All polyunsaturated fatty acids, particularly omega‐3 fatty acids, could be involved in attention deficit, dyslexia, hyperactivity, and autism (Richardson and Ross, 2000c). The severity of dyslexic signs was found to vary with the deficiency of polyunsaturated fatty acids in boys, but not in girls (Richardson et al., 2000a). Fatty acids seem to be important for attention deficit associated with hyperactivity (Richardson and Puri, 2000b). The oxidative breakdown of omega‐3 fatty acids was found to be 50% above normal, whereas that of protein was unaltered (Ross et al., 2003). Supplementing the diets of 50 children with essential fatty acids (daily doses of 80 mg EPA, 480 mg DHA, 96 mg gamma‐linolenic acid, 40 mg arachidonic acid, and 24 mg tocopherol acetate) improved their symptoms (Stevens et al., 2003). But another study, on hyperactive children suffering from attention deficit, found that dietary supplements did not improved their clinical symptoms, although it had a positive influence on their blood parameters (Voigt et al., 2001). Finally, a study on 135 adults (74 men) found that dyslexia was accompanied by a lack of polyunsaturated fatty acids (Taylor et al., 2000). Adults suffering from hyperactivity may respond better to a high fish diet rather than high doses of flax oil.

3.4 Drug Addiction A diet‐related change in the membranes of certain regions of the brain or neurons could perhaps make some people more unstable, so that they become more predisposed to drug addiction in response to other impulses. Drug addicts often have poor diets, and this dietary deficiency can aggravate their condition and interfere with treating their addiction or make readdiction more likely. Animals lacking dietary ALA produce an altered response to morphine (Frances et al., 2000b). Polyunsaturated fatty acids may also be involved in drug addiction, both omega‐3 fatty acids and possibly omega‐6 fatty acids. Former cocain addicts whose diets lack polyunsaturated fatty acids are likely to more rapidly return to addiction (Buydens‐ Branchey et al., 2003a, b).

3.5 Depression Ecological and epidemiological studies indicate that the increased prevalence of depression over the past half century parallels fundamental changes in dietary habits, mainly a reduced intake of foods containing omega‐3 fatty acids, such as oily seafood (Colin et al., 2003). For instance, the frequency of depression in

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British Columbia increased in parallel with the loss of traditional fish eating, and then the frequency improved as these dietary seafoods were reintroduced (Bates, 1988). More isolated circumpolar populations do not yet suffer from great seasonal mood changes as do non‐Artic populations; they suffer less from SAD (seasonal affective disorders) and have unexpectedly low rates of depression in winter (McGrath‐Hanna et al., 2003), unlike other less isolated circumpolar populations. There is little doubt that omega‐3 fatty acids are important for treating seasonal affective disorder syndrome (SADS), as the disorder modifies certain activities of the hypophysis in animals, particularly melatonin secretion (Zaouali‐Ajina et al., 1999). This may be linked to a lack of seasonal mood changes in the Icelandic population (Magnuson et al., 2000). Pregnant women in a fishing community in Iceland consume considerable amounts of fish (Thorsdottir et al., 2004). Genetics alone cannot explain the rapid increase in mental distress (depression, SAD, suicides) in circumpolar peoples, but raises the question of diet: the contamination of traditional food sources is a great concern for circumpolar peoples and has led to a decrease in the use of their food resources (McGrath‐ Hanna et al., 2003). There is a close relationship between the drop in omega‐3 fatty acid consumption (in oily seafood) and the risk of depression: the incidence of the disorder varies from 1 to 50 per 1000 population, depending on the country, in parallel with fish consumption (Hibbeln and Salem, 1995; Tanskanen et al., 2001). However, this extremely interesting observation was not confirmed by another study (Hakkarainen et al., 2004b). A study in Crete showed that there is an inverse relationship between the risk of depression and the DHA concentration in adipose tissue (Mamalakis et al., 2002). Data from 41 published studies covering 23 countries showed that a fish‐poor diet led to a low DHA concentration in mothers’ milk (which is undesirable for newborns) and an increased risk that the mothers would suffer from postnatal depression; in contrast, there was no such relationship between dietary EPA or ARA (Hibbeln, 2002). Biochemical data indicate that the link between the severity of postnatal depression and the decreased plasma DHA concentration, together with a slower return to normal of the plasma DHA concentration, indicates the true importance of supplementing the maternal diet with EPA and DHA, especially during pregnancy and the period immediately after childbirth (De Vriese et al., 2003; Otto et al., 2003). An increase in the blood plasma ARA/EPA ratio is a good indicator of increased risk of depression (Adams et al., 1996); similar relationships have been found for plasma phospholipids and cholesterol esters as in major depressions (Maes et al., 1996). The severity of depression varies with the drop in the omega‐3 fatty acid concentration in erythrocyte membranes, whatever the dietary caloric intake may be (Edwards et al., 1998). This could be linked to oxidative damage (Peet et al., 1998). Some clinical studies have used 2 g per day of EPA (as the ethyl ester) to successfully treat cases of depression that responded only partially to classical psychiatric treatment (Nemets et al., 2002; Peet and Horrobin, 2002). A patient given EPA together with conventional treatment showed improved clinical signs (suicidal tendencies, social phobias) and also was found to undergo changes in brain morphology (reduced volume of the lateral ventricles) (Puri et al., 2001). DHA has also been used successfully to treat minor depression (Mischoulon and Fava, 2000). A patient suffering from postnatal depression was successfully treated with omega‐3 fatty acids (Chiu et al., 2003), but, another study that used fish oil (2.7 g per day, EPA/DHA ratio¼1.4) starting at the 34– 36th week of pregnancy and continuing until 12 weeks after birth showed no effects (Marangell et al., 2004). EPA may increase the action of antidepressant drugs (Murck et al., 2004). A dietary supplement of 200 mg DHA per day for 4 months after childbirth prevented the drop in plasma DHA, but did not alter the patients’ self‐evaluated state of depression (Llorente et al., 2003). This could be because they measured mood changes rather than real depression and/or because the doses of DHA were too low. Treating patients suffering from major depression with 9.6 g omega‐3 fatty acid per day for 8 weeks gave positive results (Su et al., 2003). In contrast, DHA alone (2 g per day for 6 weeks) had no effect (Marangell et al., 2003). A reduction in omega‐3 fatty acid consumption increases the risk of depression and suicide, perhaps by increasing the serotoninergic activity in the brain and reducing impulsive and aggressive behavior (Brunner et al., 2002). A study of bipolar disorder patients (manic depressives) in 14 countries showed a correlation between the prevalence of the disorder and low fish consumption, with the threshold of vulnerability being 65 g per day (Noaghiul and Hibbeln, 2003); treatment with omega‐3 fatty acids could be useful under certain specific conditions (Stoll et al., 1999).

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But omega‐3 fatty acids are probably not involved (at least not directly) in most neurological depression disorders, such as brain tumors, cranial/cerebral trauma, Parkinson’s disease, temporal epilepsy, or in endocrine depressions such as those accompanying hypo‐ and hyperthyroid syndromes. Psychiatric signs of a lack of vitamin B12 (and folate and vitamin B9) precede a drop in its circulating concentration. Major depression is associated with lower omega‐3 fatty acid concentrations in patients with recent acute coronary syndromes, thus dietary, genetic, and hormonal factors may all influence depression and coronary heart disease (Frasure‐Smith et al., 2004). However, there is little convincing evidence of a relationship between an increased risk of depression with age and the changes in fatty acids, although there is a link between the plasma fatty acid composition and depression during aging (Tiemeier et al., 2003). Regular fish consumers in a community, elderly French people aged 65 years and above, had fewer depressive symptoms and scored higher on a Mini Mental Status Examination. Similarly, regular fish consumers (at least weakly) were found to be better educated (OR 1.19–1.65) and have a higher income (1.37–1.89.) (Barberger‐Gateau et al., 2005). Classical treatment with any of the antidepressants does not seem to normalize membrane omega‐3 fatty acids (Coppen and Bailey, 2000).

3.6 Dementias and Alzheimer’s Disease A 1/4 mixture of ALA and LA, given in capsules containing vegetable oil, improved the quality of life for people suffering from Alzheimer’s disease, as measured by tests of mood, appetite, spatial orientation, cooperation, sleep and hallucinations, short‐ and long‐term memory (Yehuda et al., 1996). Recent ecological and epidemiological studies have shown that dietary EPA and DHA may be important in the prevention of dementia. The Rotterdam study showed that the risk of dementia (with vascular features) was negatively correlated with the consumption of omega‐3 fatty acid‐rich fish and positively correlated with the consumption of saturated fatty acids (Kalmijn et al., 1997a). However, this result was not confirmed in another study (Engelhart et al., 2002). Morris et al. (2003b) found that a diet rich in unsaturated fatty acids and unhydrogenated fat protected against Alzheimer’s disease, in contrast to a diet rich in saturated fatty acids and trans fatty acids. The consumption of meat is poorly correlated with an increased risk of dementia in France, whereas the consumption of fish has a protective effect. This was found by following participants in the PAQUID study, which lasted 7 years and involved 1,416 subjects aged more than 67 years living in the south west of France. The population included 170 cases of dementia, 135 of which were Alzheimer’s. Those subjects who ate fish at least once a week were 34% less likely to develop any form of dementia, and 31% less likely to suffer from Alzheimer’s disease. The effect was still present when socioeconomic factors were taken into account, as these factors are linked to both the reduced risk of Alzheimer’s disease and the fish consumption (Barberger‐Gateau et al., 2002). Similarly, a study carried out in the USA found that Alzheimer’s disease was 60% less common in people who consumed about 60 mg DHA per day (at least one seafood meal a week) than in people who ate very little (Morris et al., 2003a, b). The findings were similar in Japan (Otsuka, 2000). A low plasma concentration of omega‐3 fatty acids (including DHA) is an indication of risk of cognitive deficiencies and various types of dementia, including Alzheimer’s disease (Conquer et al., 2000). However, one study found that plasma omega‐3 fatty acid concentrations were unaltered (Laurin et al., 2003). DHA attenuates amyloid‐beta (Abeta) secretion by human neural cells under cytokine stress, and this effect is accompanied by the formation of NPD1, a novel DHA‐derived 10,17S‐docosatriene (Lukiw et al., 2005). Vascular dementias and Alzheimer’s disease have nutritional factors in common: an excess of omega‐6 fatty acids and a lack of omega‐3 fatty acids; this leads to changes in the microvasculature, chronic inflammation, platelet aggregation, and endothelial dysfunction (Otsuka et al., 2002). These changes provide at least a partial explanation of why the cognitive disorders that occur in very elderly people are positively correlated with the consumption of LA, and negatively correlated with the consumption of fish. However, there has been no published report on the use of omega‐3 fatty acids to prevent dementia. The cardiovascular risk increases the risk of dementia, particularly vascular dementia (Kalmijn et al., 2000).

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Inflammatory processes may well be implicated in all these disorders (Simopoulos, 2002). Nevertheless, the influence of omega‐3 fatty acids on vascular components and the cerebral parenchyma are not yet completely clear. There appears to be a relationship between dementia and cardiovascular risk, as there is a lack of omega‐3 FA in both cases, together with an increase in homocysteine concentration (Severus et al., 2001). Elevated plasma homocysteine is also a risk factor for psychiatric disorders during aging (Reutens and Sachdev, 2002). Curiously, there have been apparently no studies on the implication of omega‐3 fatty acids in alcoholism and alcoholic dementia, although animal experiments have shown that dietary ALA modulates the effects of alcohol on various membranes, including nerve endings (Zerouga et al., 1991). There may be a correlation between dementia and overweight among women (Gustafson et al., 2003).

3.7 Schizophrenia Schizophrenic patients who eat large amounts of seafood have less severe clinical signs (Peet et al., 1996). There could well be subgroups of patients for whom the omega‐3 fatty acids are particularly important, which would explain why clinicians have found divergent results in which the DHA content was elevated, depressed, or normal. Some have found subnormal concentrations of DHA and EPA in the erythrocyte membranes of schizophrenics, but there was a discontinuity between the severity of the clinical disease and the change in the fatty acid profile. The differences did not depend on sex, hormonal status, or cannabis use (Assies et al., 2001). Although tobacco smoking seemed to be a discriminating factor, another study did not find these results, (Hibbeln et al., 2003). The turnover of phospholipids could be altered in schizophrenics (Fenton et al., 2000). Changes in phospholipid metabolism are believed to be implicated in the cause of schizophrenia because of the amounts of omega‐3 fatty acid‐rich phospholipids in the developing human brain (Horrobin, 1998). Treatment with 10 g fish oil per day for 6 weeks seems to improve these symptoms (Laugharne et al., 1996), as does a combination of 120 mg EPA, 150 mg DHA, 500 mg vitamin C and 400 IU vitamin E given twice a day for 4 months, but the improvement is relatively modest (Arvindakshan et al., 2003). Symptoms were improved in a schizophrenic patient given EPA alone; brain atrophy had receded after treatment for 6 months, and the turnover of brain phospholipids, measured by 31P NMR, returned to normal (Puri et al., 2000). Patients were treated and stabilized by a course of EPA for 3 months (Peet et al., 2001); it was also used to supplement a 6‐month treatment with antipsychotic drugs, but it left residual symptoms (Emsley et al., 2002). One author reported that treatment with 3 g EPA per day produced no results (Fenton et al., 2001), perhaps because the patients concerned were nonresponders or because the dose was wrong (Horrobin, 2003). There have been several proposals as to how omega‐3 fatty acids are involved in schizophrenia (Peet, 2003), including the modulation of neurotransmission, particularly dopaminergic transmission, but there is as yet no hard evidence (Peet et al., 2001). DHA is a ligand for the retinoid X receptor in the mouse brain (de Urquiza et al., 2000) and can reduce haloperidol‐induced dyskinesia in mice. As this involves retinoid receptors (Ethier et al., 2004) it offers a possible treatment of the symptoms of schizophrenia. Typical antipsychotic drugs are partly effective, but the induction of extrapyramidal symptoms represents a serious handicap, which considerably limits their usefulness. Fatty acids affect binding of serotonin to the serotonin receptor and transporter in the rat brain, which may be important in neuropsychiatric diseases such as schizophrenia, where there are links between altered fatty acids and the serotoninergic system (du Bois et al., 2006).

3.8 Autism Bell et al. (2000) found that the phospholipids in the erythrocytes of autistic children were 70% below normal. Similarly, the DHA in the plasma phospholipids of autistic children is 23% subnormal, with the

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total omega‐3 fatty acid concentration being decreased by 20% and the omega‐6 fatty acids being unchanged (Vancassel et al., 2001). An autistic boy aged 11 years was successfully treated with fish oil (540 mg EPA per day) for 4 weeks (Johnson and Hollander, 2003). The transmission of signals involving phospholipids is nevertheless normal in these subjects (Puri and Singh, 2002). However, there are very few studies for any conclusion to be reached.

4

Aging

4.1 Biochemical Changes and Diet Increasing age results in a reduction in the arachidonic acid incorporated into the brain and a reduction in its turnover (Gatti et al., 1986). Nutrition plays an important role in myelination as myelin membranes are extremely rich in lipids (Svennerholm et al., 1992), consequently, and severe malnutrition and a lack of essential fatty acids cause severe hypomyelination (Di Biase and Salvati, 1997) and reduced myelin lipid turnover during aging (Ando et al., 2003). A study on the composition of the human cortex between the ages of 2 years and 88 years (Carver et al., 2001) found that DHA and monounsaturated fatty acid concentrations increased up to the age of 18 years. The concentrations of polyunsaturated fatty acids, especially ARA, decreased with aging, whereas that of ALA increased. The polyunsaturated fatty acid content of phosphatidylethanolamine is markedly decreased in Alzheimer’s disease (Edlund et al., 1992). Major changes occur in the polyunsaturated fatty acid composition of the rat cortex and hippocampus, and these especially affect the DHA content of phosphatidylethanolamine plasmalogens (Favrelere et al., 2000). These age‐related changes are associated with altered participation in membrane structures resulting in reduced cognitive performance, neurotransmission, and antioxidant function (Brosche and Platt, 1998). The ARA, 22:4o6 and DHA concentrations in the cortex and cerebellum decrease with age, whereas the concentrations of the 18:1o9 and 20:1o9 FA increase mainly in phosphatidylethanolamines and phosphatidylserines (Lopez et al., 1995; Giusto et al., 2002). These defects can be corrected by an appropriate diet (McGahon et al., 1999a, b). The exchanges of choline and serine bases are also modified during aging (Ilincheta de Boschero et al., 2000). The microsomes synthesize less phosphatidylethanolamines and phosphatidylserines with age (Montanini et al., 1983), and the activity of the enzyme phosphatidylserine decarboxylase decreases, reducing the production of phosphatidylethanolamine (Salvador et al., 2002). A lack of dietary ALA causes the selective reduction of phosphatidylserine and an increase in MAO‐B activity in the hippocampus of aging rats, but has no effect on the serotonin and noradrenaline contents (Delion et al., 1997). The cerebral concentration of the proinflammatory cytokine (interleukin‐1‐beta) increases with age (Martin et al., 2002a), and this may be responsible for some deterioration of certain cellular functions, especially as the binding of interleukin‐1 to its receptor inhibits the release of glutamate from the hippocampal nerve endings of young rats, but not from those of older rats (McGahon et al., 1998). Reducing cerebral glucose levels leads to modifications of cerebral metabolism, resulting in peroxidation in aging rats (Benzi et al., 1987). The turnover of phospholipids (especially phosphatidylcholine and phosphatidylethanolamine) and cholesterol in synaptic membranes slows during aging (Ando et al., 2002). The lipid composition of the mitochondria in synapses is also affected, especially their LA content (Ruggiero et al., 1992). Thus, relationship between changes in fatty acid composition during aging and diet is poorly documented, even in animals.

4.2 Cognitive Alterations During Aging The changes that occur with advancing age are complex in both animals and humans. Omega‐3 fatty acids may be directly or indirectly involved, depending on the part of the body, the structure, cells and organelles,

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or lipids concerned (Bourre, 2004a). Peroxysome metabolism is also implicated in controlling brain fatty acids during aging, especially for polyunsaturated fatty acids (Perichon et al., 1998). A recent French study showed that age‐related cognitive deficit is linked to a reduction in the omega‐3:omega‐6 fatty acid ratio in erythrocytes (Heude et al., 2003); an excess of nutritional linoleic acid has also been linked to a decline in cognitive performance, whereas the reverse is true for fish oils (Kalmijn et al., 1997a, b). Fish consumption may be associated with a slower cognitive decline with age: the rate was 10% slower among persons who consumed one fish meal per week, and 13% slower in persons who consumed two or more fish meals per week (Morris et al., 2005). An increased oxidative stress during aging, due to reduced protection against free radicals, can result in a lower omega‐3 fatty acid concentration in the nervous system. Hence a diet rich in EPA could have antioxidant properties that would help counteract the effects of aging (Martin et al., 2002b). A lower concentration of polyunsaturated fatty acids in brain structures could result in poorer movement of solutes across the blood–brain barrier because of poorer incorporation into membranes, or reduced activities of the enzymes delta‐6 and delta‐9 desaturase, together with increased production of free radicals due to oxidative stress. All these factors reduce membrane fluidity (Yehuda et al., 2002). Phosphatidylcholine improves the memory, learning, concentration, vocabulary recall, and mood in elderly people suffering from cognitive loss (Kidd, 1999). Phosphatidylcholine, together with vitamin B12, improves learning in aging mice (Hung et al., 2001). There appears to be no question but that an adequate intake of omega‐3 fatty acids ensures the turnover of membranes, so helping to protect against brain aging. However, a dietary supplement of high concentrations of omega‐3 fatty acids produces behavioral changes that vary with age of the individual, improving learning in young animals, but reducing learning and motor activity in older ones (Carrie et al., 2000c). This should be borne in mind when considering dietary supplements. Several studies have been carried out on humans and experimental animals to identify the abnormalities of lipid metabolism, particularly phospholipids that are associated with aging, together with a range of neurological and psychological disorders. The tissues examined have included the brain and the skin, where lipid messengers are involved. As these have no direct link with diet they are not included here, but can be found in Bourre (2004a).

5

Miscellaneous: Neurological Disorders and Handicaps

Although supported by only a few studies, it has long been proposed that fatty acids are important for preventing (and eventually treating some clinical aspects of) multiple sclerosis. Perhaps they just supply the polyunsaturated fatty acids needed for maintenance and reconstruction of the myelin sheath (Bates, 1990), but a dietary supplement helps to reduce the severity of the disease and prevent recurrence, at least for 2 years (Nordvik et al., 2000), whereas LA in the plasma erythrocytes is decreased (Di Biase and Salvati, 1997). Giving patients 0.9 g omega‐3 fatty acid per day (as fish oil) for a total of 2 years may decrease the occurrence and severity of disease incidents, perhaps by modulating cytokines (Calder, 1997). The effect of omega‐3 fatty acids on other neurological diseases is poorly supported by convincing results. No association was found for dietary saturated fat, cholesterol, or trans fatty acids in sufferers from Parkinson’s disease, but a high intake of unsaturated fatty acids might protect against the disease (de Lau et al., 2005). The changes seen in some patients, such as those suffering from infantile neuronal ceroid lipofuscinosis associated with dementia, are side effects (Vallat et al., 1985). The successful treatment of Zellweger peroxysomal disease with DHA simply compensates for the metabolic defect (Martinez, 2001). Neurologically handicapped children do not seem to consume enough omega‐3 fatty acids, as indicated by the presence of marker such as 20:3 omega‐9 and 22:5 omega‐6 fatty acids in their blood plasma. This deficiency does not facilitate the renewal of their damaged brain structures (Hals et al., 2000).

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Desaturases and the Relationship between Omega‐3 Fatty Acids and Antioxidants

Dietary ALA is probably elongated and unsaturated in the liver to give longer carbon chains. These fatty acids are essential for the brain, as shown in studies on fetal brain cells; such cells multiply, take up, and release neurotransmitters only if there is ARA or DHA in the medium, and not if the medium contains only LA or ALA (Bourre et al., 1983; Tixier‐Vidal et al., 1986). The activities of desaturases, particularly delta‐6‐ desaturase (the first enzyme in the synthesis of long‐chain, more unsaturated fatty acids it acts on ALA and LA), are still to be evaluated, but they are very much less active immediately after birth, and essentially zero in the brains of animals; the liver activities are greatly reduced during aging (Hrelia et al., 1989; Bourre and Piciotti, 1992). As a result, DHA comes from the hepatic transformation of dietary ALA or directly from the diet. However, the potential of astrocytes should not be neglected (Williard et al., 2001). Little is synthesized at the blood–brain barrier but some may be produced at the choroid plexus as it has a high delta‐6‐ desaturase activity (Bourre et al., 1997a). Nevertheless, mother’s milk provides most of the ARA and DHA ingested by the newborn infant (Larque et al., 2002). ALA is 7–10 times more effective, mole for mole, than DHA (Abedin et al., 1999) for the construction and maintenance of nervous system membranes (Su et al., 1999). Dietary ALA and one‐tenth as much DHA result in identical DHA concentrations in the brains of developing animals (Valenzuela et al., 2004). Moreover, DHA is more biologically available than ALA (Poumes‐Ballihaut et al., 2001). A newborn infant can transform ALA into DHA and LA into ARA, as shown by studies with stable isotopes. But this capacity is limited in newborns, as in any other age of the life: activities are low in humans, with some variations according sex, age, and physiopathological conditions (Williams and Budge, 2006; Goyens et al., 2006). Hence, DHA is itself also considered to be an essential nutrient (Muskiet et al., 2004). Vegetarians need much more ALA than the general population because of their restricted dietary intake of DHA; ALA is converted to DHA relatively poorly and there is active competition from omega‐6 fatty acids (Davis and Kris‐Etherton, 2003). ALA is undoubtedly an essential nutrient, unlike the other omega‐3 fatty acids and their derivatives; however, it must be combined with DHA from oily fish. Any ALA or DHA added to the diet must be protected from oxidation in both the food itself and in the tissues into which it is incorporated. Animal experiments have shown that only alpha‐D‐tocopherol is integrated into the nervous system membranes and not, for example, gamma tocopherol (Clement et al., 1995; Clement and Bourre, 1997). The metabolic system of an infant discriminates between natural vitamin E and the synthetic vitamin (Stone et al., 2003). Although the ingestion of large amounts of fish oil modifies the fatty acid profile of the brains of animals without causing peroxidation (Chaudie`re et al., 1987), any diet of newborn infants that supplements their intake of very long‐chain polyunsaturated fatty acids must include vitamin E (Koletzko et al., 1995; Kaempf‐Rotzoll et al., 2003). The activities of desaturases are modified by vitamin E (Despret et al., 1992).

7

Conclusions

The timetable of cerebral development means that any short‐term perturbation can result in long‐term changes in the biochemistry, physiology, and function of the brain, and the possibilities of recovery are relatively restricted, because of the genetically programmed brain development. A dietary lack of omega‐3 fatty acids results in their replacement by others. Dietary omega‐3 fatty acids contribute to the construction and maintenance of the brain. They are involved in many membrane‐based reactions (Bourre, 2004a), and even in the regulation of gene expression in the brain (Kitajka et al., 2004; Lapillonne et al., 2004). A lack of ALA leads to abnormal activities of certain nuclear membrane enzymes (Ammouche et al., 1994) and this polyunsaturated fatty acid can protect against neurone death and other deleterious processes (Crawford et al., 2003). Peroxisomal oxidation (Bourre and Piciotti, 1997) and the recycling of polyunsaturated fatty acids are important, and will undoubted be the subject of future studies (Cunnane et al., 1999). A part of the dietary

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ALA is used to synthesize lipids (Menard et al., 1998), and is thus not subject to desaturation and chain lengthening (Cunnane et al., 2001; Cunnane et al., 2003), or oxidation (Yang et al., 1994). Little is known about how fatty acids cross the blood–brain barrier, although phospholipid synthesis (Magret et al., 1996), sphingolipids (Homayoun et al., 1989), and lysophosphatidylcholine (Bernoud et al., 1999) may all be involved. Brain microvessels, which control the supply of nutrients to the brain and are largely responsible for the blood–brain barrier, are sensitive to omega‐3 fatty acids (Homayoun et al., 1988; Ziylan et al., 1992). In vivo approaches have been developed for quantifying and imaging brain fatty acids, including ARA and DHA (Rapoport, 2003). This review of omega‐3 fatty acids focuses on the importance of the balance between omega‐3 and omega‐6 fatty acids. However, monounsaturated fatty aids, particularly omega‐9 fatty acids, must not be ignored, as animal experiments have shown that the endogenous supply is not adequate during development. They are conditionally essential during this period of life (Bourre et al., 1997b). Certain saturated fatty acids, particularly the very long‐chain fatty acids in myelin, like lignoceric acid (C24:0) may also be required (Bourre et al., 1977; Cinti et al., 1992), which raises the serious question of how essential are certain dietary saturated fatty acids (Beare‐Rogers, 1995). The enzymes involved in the synthesis of myelin components appear to all be activated simultaneously, rather than sequentially (Tennekoon et al., 1980). There are therefore two fundamental concepts of how fatty acids influence brain physiology. One is the effects of dietary fatty acids, especially omega‐3 fatty acids, on the production and maintenance of brain structures, and hence on their function. The other involves the physiological mechanisms in which fatty acids take part. Thus, omega‐3 fatty acids have two broad actions, one long term and the other short term. Their long‐term actions are on membrane composition and function. This is supported by studies on brain development and probably the role of dietary omega‐3 fatty acids in the prevention of dementia, including Alzheimer’s disease. Their short‐term actions could involve phospholipids metabolism, and hence the modulation of signal transduction. The evidence for this includes the effect of EPA on depression, schizophrenia, and autism. In addition, the two types of action can occur simultaneously, as in inflammation during Alzheimer’s disease. However, it is too early to state that omega‐3 fatty acids prevent depression by treating inflammation, even though there are good indications of inflammation occurring during depression. Although there is clear evidence that omega‐3 fatty acids do, to some extent, prevent and reduce these symptoms, these relationships are not necessarily causal. The effects of omega‐3 fatty acids on brain structures and functions are clear, whereas most experiments that have focused on the prevention of psychological problems (like depression, dementias, and other disorders) have concerned by consuming oily seafood. Their main characteristic is their high omega‐3 fatty acid content; however, other components could be involved, directly or indirectly: iodine has an important effect on the brain biochemistry and function, selenium may also have positive effects, as well as cobalamin. Clinical trials have yet to provide proof that capsules containing omega‐3 fatty acids as fish oil extracts are effective. The same applies to purified omega‐3 fatty acids, generally supplied as their ethyl esters, as not nearly enough experiments or psychiatric clinical trials have been done to provide conclusive, convincing evidence. Moreover, the bioavailability of omega‐3 in fish oil, especially DHA, is much higher than in capsules (Visioli et al., 2003, 2005). The major sources of ALA in the normal diet is rapeseed (canola) oil, walnut oil (which is less readily available; perilla oil in Japan), and soybean oil (but this also contains a large amount of LA). Walnuts are a good source. The main sources of EPA and DHA are seafood, especially oily fish (both wild and farmed), the farmed fish must be correctly fed, as discussed in a recent issue (Bourre, 2004b) and ‘‘omega‐3 eggs.’’ Transformation of ALA into longer chains by the human body seems to be relatively slow, and differs for men and women and with age and physiological condition (Williams and Burdge, 2006). The availability of omega‐3 fatty acids, especially DHA, in the diet has probably had a major influence on the evolution of the human brain (Crawford et al., 2001). Thus, the diet must provide both ALA and DHA. Although an increased dietary intake of omega‐3 fatty acids (specifically ALA) is fundamental, it is not sufficient. Omega‐6 fatty acids must also be taken into account, to ensure that the omega‐6/omega‐3 ratio is not more than 5. This ratio is too high in the diet of most western people. It can only be effectively reduced by increasing the intake of omega‐3 fatty acids rather than by decreasing the intake of omega‐6 fatty acids.

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Choline and Its Products Acetylcholine and Phosphatidylcholine

R. J. Wurtman . M. Cansev . I. H. Ulus

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 445

2 2.1 2.1.1 2.1.2 2.1.3 2.2 2.2.1 2.3

Choline in the Blood . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 446 Sources of Plasma Choline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 446 Dietary Choline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 447 Endogenous Choline Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 449 Choline‐Containing Membrane Phospholipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 452 Fates of Circulating Choline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 454 Choline as a Source of Methyl Groups (Choline Oxidase System) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 456 Effects of Physiologic or Pathologic States on Plasma Choline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 457

3 3.1 3.1.1 3.1.2 3.1.3 3.1.4

Choline in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 458 Sources of Brain Choline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 458 Uptake of Circulating Choline into the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 459 Liberation from Membrane PC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 459 Reutilization of Choline Formed from Hydrolysis of Acetylcholine . . . . . . . . . . . . . . . . . . . . . . . . . . . . 460 De Novo Synthesis of Phosphatidylcholine and Choline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 460

4 4.1 4.1.1 4.1.2 4.2 4.2.1 4.2.2 4.2.3 4.2.4 4.3

Brain Proteins that Interact with Choline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 460 Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 461 Choline Acetyltransferase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 461 Choline Kinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 462 Transport Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 462 Facilitated‐Diffusion Carrier at Blood–Brain Barrier . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 462 Choroid Plexus Choline Transporter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 463 High‐Affinity Uptake Protein in Cholinergic Terminals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 463 Low‐Affinity Cellular Uptake Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 464 Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 465

5 5.1 5.2 5.2.1 5.2.2 5.2.3 5.2.4

Utilization of Choline in Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 466 Biosynthesis of Acetylcholine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 467 Biosynthesis of Choline‐Containing Phosphatides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 467 CTP:Phosphocholine Cytidylyltransferase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 471 CDP‐Choline:1,2‐Diacylglycerol Cholinephosphotransferase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 472 Uptake of Uridine and Cytidine into Brain Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 473 Phosphorylation of Uridine and Cytidine to UTP and CTP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 473

6 6.1

Physiological and Behavioral Effects of Choline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 474 Blood Pressure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 474

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_18, # Springer ScienceþBusiness Media, LLC 2009

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Choline and its products acetylcholine and phosphatidylcholine

6.2 6.3 6.4 6.5 6.6 6.7 6.8

Body Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475 Pain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475 Neuroendocrine Effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475 Peripheral Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475 Behavior . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 476 Drug Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 476 Neuroprotective and Cytoprotective Effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 477

7 7.1 7.2 7.3 7.4 7.5

Effects of Exogenous CDP‐Choline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 477 Hypoxia and Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 478 Head Trauma (Cranio‐Cervical Trauma) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 479 Induced Lesions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 479 Other Effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 479 Clinical Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 480

8

Choline in Autonomic and Motor Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 480

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Abstract: Choline, a quaternary amine obtained largely from the diet but also synthesized in the brain and, especially, liver, is an essential precursor of the neurotransmitter acetylcholine (ACh) and of the major membrane constituent phosphatidylcholine (PC). Plasma choline concentrations can vary over a fivefold range depending on the choline contents of the foods being digested. Since choline readily crosses the blood–brain barrier (BBB) through an unsaturated facilitated‐diffusion system, these plasma changes can produce parallel changes in brain choline levels. In addition, since the enzymes that convert choline to ACh [choline acetyltransferase (ChAT)] and PC’s precursor phosphocholine [choline kinase (CK)] are also poorly saturated with their choline substrate, increases in plasma choline can enhance the formation of ACh and phosphocholine, and the release of ACh. The subsequent conversion of phosphocholine to PC is increased if PC’s other circulating precursors (uridine and omega‐3 fatty acids) are provided. This leads to an increase in the levels of synaptic membrane within the brain. Choline is principally metabolized in the liver to betaine, which provides a source of methyl groups for the regeneration of methionine and S‐adenosylmethionine. List of Abbreviations: 5-methyl-THF, 5-methyl-tetrahydrofolate; 5,10-MTHF, 5,10-methylene-tetrahydrofolate; AA, arachidonic acid; ACh, acetylcholine; AChE, acetylcholinesterase; ACTH, adrenocorticotropic hormone; AI, adequate intake; AMPA, DL-amino-3-hydroxy-5-methyl-isoxazole-4-propionic acid; APP, amyloid precursor protein; BBB, blood-brain barrier; BuChE, butyrylcholinesterase; CaM-kinase, Ca+2/calmoduline kinase; CDP, cytidine-50 -diphosphate; CDP-choline, cytidine-50 -diphosphocholine; ChAT, choline acetyltransferase; CHT, high-affinity choline transporter; CK, choline kinase; CMP, cytidine-50 -monophosphate; CNS, central nervous system; CNT, concentrative nucleoside transporter; CP, choroid plexus; CPT, CDP-choline:1,2-diacylglycerol cholinephosphotransferase; CT, CTP:phosphocholine cytidylyl transferase; CTL, choline-transporter-like protein; CTP, cytidine-50 -triphosphate; DAG, diacylglycerol; DHA, docosahexaenoic acid; ECF, extracellular fluid; ENT, equilibrative nucleoside transporter; EPA, eicosapentaenoic acid; EPT, CDP-choline:1,2-diacylglycerol ethanolaminephosphotransferase; FAD, flavine adenine dinucleotide; FNB, Food and Nutrition Board; GH, growth hormone; GPC, glycerophosphocholine; GPC-CPD, Glycerophosphocholine cholinephosphodiesterase; GPC-PD, glycerophosphocholine phosphodiesterase; GRAS, generally regarded as safe; HC3, hemicholinium-3; IP3, inositol triphosphate; LH, luteotropic hormone; LHRH, luteinizing hormone-releasing hormone; LPCAT, lysophosphatidylcohline acyltransferase; LTP, long-term potentiation; mAChR, muscarinic acetylcholine receptor; MAPK, mitogen-activated protein kinase; nAChR, nicotinic acetylcholine receptor; NAD, nicotinamide adenine dinucleotide; NDPK, nucleoside diphosphate kinase; NGF, nerve growth factor; NMDA, N-methyl-D-aspartate; OCT, organic cation transporter; PAF, platelet-activating factor; PC, phosphatidylcholine; PCho, phosphocholine; PE, phosphatidylethanolamine; PEMT1, phosphatidylethanolamine-Nmethyltransferase; PEMT2, phosphatidyl-N-methylethanolamine-N-methyltransferase; PK, protein kinase; PKA, protein kinase A; PLA1, phospholipase A1; PLA2, phospholipase A2; PLC, phospholipase C; PLD, phospholipase D; PS, phosphatidylserine; PUFA, polyunsaturated fatty acid; RDA, recommended daily allowance; SAH, S-adenosylhomocysteine; SAM, S-adenosylmethionine; SM, sphingomyelin; THF, tetrahydrofolate; TRH, thyrotropin-releasing hormone; TSH, thyroid-stimulating hormone; UL, upper limit; UCK, uridine-cytidine kinase; UMP, uridine-50 -monophosphate; UDP, uridine-50 -diphosphate; UTP, uridine-50 -triphosphate

1

Introduction

Choline (2‐hydroxyethyl‐trimethyl‐ammonium), a simple but unusual compound, consists of a 2‐carbon chain in which one carbon is attached to a hydroxyl group and the other to an amine nitrogen (> Figure 18‐1). The particularly unusual quality of choline is that this amine nitrogen bonds with a total of four hydrogen or carbon atoms instead of with the usual three, and thus carries a partial positive charge. Though choline is not an amino acid, it shares with that family of compounds the property of being present in cells both in free form and—like the amino acids in proteins—within subunits (e.g., PC) of a macromolecule (biologic membranes). Moreover, like tyrosine, tryptophan (Cansev and Wurtman, 2006),

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Choline and its products acetylcholine and phosphatidylcholine

. Figure 18‐1 Structure of choline

and histidine, choline is the precursor of a neurotransmitter (ACh), and also like tyrosine and histidine, choline must be obtained from both endogenous synthesis and dietary sources. This chapter describes the choline in the blood and, particularly, in the brain, —its sources; its uses to make membrane phospholipids and ACh; and its other biologic effects.

2

Choline in the Blood

Choline is a normal constituent of the plasma (> Table 18‐1), present as the free base (Cohen and Wurtman, 1975; Hirsch et al., 1978; Savendahl et al., 1997); as a constituent of phospholipids [including PC; sphingomyelin (SM); lyso‐PC; choline‐containing plasmalogens; and the platelet‐activating factor (PAF)] and as PC’s water‐soluble metabolites [principally phosphocholine and glycerophosphocholine (GPC) (Hirsch et al., 1978; Klein et al., 1993; Ilcol et al., 2005a)]. Free choline is also found in other biological fluids (> Table 18‐1). In blood, choline is concentrated within erythrocytes (10–150 mM;

. Table 18‐1 Free choline concentrations in human body fluids Body fluids Plasma Serum Urine Cerebrospinal fluid Amniotic fluid Colostrum Breast milk Semen Peritoneal dialysate

Free choline (mM) 7.0–13.0 9.0–13.3 15.5  8.8 0.7–2.5 22.8–24.5 132  21 110–300 17,000–24,000 14–28

References Holm et al. (2003) Holm et al. (2003); Ilcol et al. (2002b, 2005a) Buchman et al. (1999) Flentge et al. (1984); Ikeda et al. (1990); Toghi et al. (1996) Ilcol et al. (2002e) Ilcol et al. (2005a) Holmes et al. (2000); Ilcol et al. (2005a) Takatori et al. (1984); Manabe et al. (1991) Hjelle et al. (1993); Ilcol et al. (2002b)

Note: Values are given as the range of the means from the cited references, or as the mean  SEM

Jope et al., 1980; Stoll et al., 1991) through the action of an uptake molecule that is unsaturated (Km ¼5– 10 mM; Riley et al., 1997) at normal plasma choline concentrations.

2.1 Sources of Plasma Choline Plasma choline derives from three sources—dietary choline, consumed as the free base or as a constituent of phospholipid molecules; endogenous synthesis, principally in liver; and liberation from its reservoir within the membrane phosphatides of all mammalian cells.

Choline and its products acetylcholine and phosphatidylcholine

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2.1.1 Dietary Choline Choline is present within many foods (> Table 18‐2), principally as the free molecule or as phosphatides, and its plasma levels can rapidly increase severalfold after ingestion of choline‐rich foods (Hirsch et al., 1978). For example, the consumption of a 5‐egg omelet (containing about 1.3 g of choline) by humans increased these levels from 9.8 mM to 36.6 mM within 4 h (Hirsch et al., 1978). Prolonged fasting reduced human plasma choline levels from 9.5 mM to 7.8 mM after 7 days (Savendahl et al., 1997). Similarly, removal of all choline‐containing foods from the diet for 17–19 days gradually lowered plasma choline, from 10.6 mM to 8.4 mM in humans (Zeisel et al., 1991; Zeisel, 2000) and from 12.1 mM to 6.3 mM in rats (Klein et al., 1998), indicating that plasma choline can be partially but not fully sustained by release from endogenous stores. Dietary PC is deacylated within the gut to form lyso‐PC. About half of this product is further degraded to free choline within the gut or liver. The remainder is reacylated to regenerate PC (Houtsmuller, 1979), which is then absorbed into the lymphatic circulation (Fox et al., 1979) and eventually enters the blood stream. Much of the dietary choline that reaches the liver through the portal circulation is destroyed by oxidation to betaine, as described later (> Figure 18‐2), ultimately providing methyl groups that can be used to regenerate S‐adenosylmethionine (SAM) from homocysteine. The rest of the choline in portal venous blood passes into the systemic circulation (Fox et al., 1979; Houtsmuller, 1979). In 1998, the Food and Nutrition Board (FNB) of the US Institute of Medicine established a dietary reference intake (DRI) for choline. Since the FNB did not believe that existing scientific evidence allowed calculation of a recommended daily allowance (RDA) for choline, it instead set an adequate (daily) intake level (AI), and an upper (daily) limit (UL) that should not be exceeded (> Table 18‐3). The main criteria for determining the AI and UL were, respectively, the amount of choline needed to prevent liver damage, and the choline intake associated with choline’s most sensitive adverse effect, i.e., hypotension (see Dietary Reference Intakes, Institute of Medicine, National Academy of Sciences USA, 1998). Subsequent studies have shown that the enzymes (described later) that synthesize and metabolize choline can be affected by

. Table 18‐2 Choline contents of common foods A. High Food Egg yolk Beef liver Chicken liver Cereals, ready to eat or wheat germ Pork or bacon Cake, chocolate, without frosting Coffee, instant, decaffeinated Cauliflower B. Low Food Olive oil Kale Iced tea Egg white Apple juice Coffee, brewed from grounds

Choline 1 61 69 69 12 5 93 24.5

GPC 1 83 5 33 18 61 8 0.7

PCho 1 11 6 4 3 1 0 1.8

PC 634 245 213 44 86 58 0 12.1

SM 45 24 14 0 10 2 0 0

Total Choline 682 424 307 150 129 127 101 39.1

Choline 0 0.1 0.4 0.2 0.7 1.9

GPC 0.3 0 0 0.6 0.7 0.7

PCho 0 0 0 0 0 0

PC 0 0.3 0 0.3 0.4 0.4

SM 0 0 0 0 0 0

Total Choline 0.3 0.4 0.4 1.1 1.8 2.6

Note: Data are given as mg choline moiety/100 g of food. Foods are grouped based on having relatively high or low choline contents. Data from USDA Database for the Choline Content of Common Foods, 2004 GPC, glycerophosphocholine; PCho, phosphocholine; PC, phosphatidylcholine; SM, sphingomyelin

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. Figure 18‐2 Metabolism of choline to betaine, methionine and, S‐adenosylmethionine (SAM). THF, Tetrahydrofolate; 5,10‐MTHF, 5,10‐methylene‐tetrahydrofolate; 5‐methyl‐THF, 5‐methyl‐tetrahydrofolate; B12, Vitamin B12; SAM, S‐adenosylmethionine; SAH, S‐adenosylhomocysteine

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. Table 18‐3 Proposed adequate (i.e., minimum) daily choline intake, and upper limit (which should not be exceeded) A. Adequate Intakes (AI) Age

Gender

AI (mg/day)

0–6 months 7–12 months 1–3 years 4–8 years 9–13 years 14–18 years 9–13 years 14–18 years Above 19 years Above 19 years Pregnancy Lactation

Both Both Both Both Male Male Female Female Male Female

125 150 200 250 375 550 375 400 550 425 450 550

B. Upper Allowable Intakes (UL, Upper Limit) Life stage

Age

UL (g/day)

Infancy Childhood

0–12 months 1–8 years 9–13 years 14–18 years 14–18 years 19 years and older 14–18 years 19 years and older

N/Aa 1 2 3 3 3.5 3 3.5

Adolescence Pregnancy Lactation a

Not possible to establish; sources of intake should only be mother’s milk and infant formulas

common genetic polymorphisms, which cause important person‐to‐person variations in dietary choline needs (da Costa et al., 2006). For further details about dietary reference intakes and the choline contents of various foods, the reader is referred to the official websites of the Institute of Medicine (http://www. nap.edu/catalog/6015.html#toc) and the USDA (http://www.nal.usda.gov/fnic/foodcomp/Data/Choline/ Choline.html).

2.1.2 Endogenous Choline Synthesis Endogenous choline is produced, principally in liver (Bremer and Greenberg, 1960) but also to a small extent within brain (Blusztajn et al., 1979; Crews et al., 1980), by the sequential addition of three methyl groups to the amine nitrogen of phosphatidylethanolamine (PE); this forms PC, which can then be broken down to liberate the choline (> Figure 18‐3). The methylation reactions are catalyzed by two enzymes, phosphatidylethanolamine‐N‐methyltransferase (PEMT1; EC: 2.1.1.17), which converts PE to its monomethyl derivative, and phosphatidyl‐N‐methylethanolamine‐N‐methyltransferase (PEMT2; EC: 2.1.1.71), which adds the second and third methyl groups (A single enzyme may catalyze all three methylations in liver). Both enzymes use SAM as the methyl donor (Bremer and Greenberg, 1960; Hirata et al., 1978). Their Kms for SAM are 2–4  10 6 M and 20–110  10 6 M, respectively (Crews et al., 1980; Blusztajn et al., 1982; Hitzemann, 1982; Percy et al., 1982), whereas brain SAM concentrations are 10–17 mg/g wet weight

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. Figure 18‐3 Sequential methylation of phosphatidylethanolamine (PE) to phosphatidylcholine (PC). PEMT‐I, Phosphatidylethanolamine‐N‐methyltransferase; PEMT‐II, phosphatidyl‐N‐methylethanolamine‐N‐methyltransferase; SAM, S‐adenosylmethionine; SAH, S‐adenosylhomocysteine

Choline and its products acetylcholine and phosphatidylcholine

18

[50–85 mM assuming about 50% of the brain mass is aqueous (Wurtman and Rose, 1970; Ordonez and Wurtman, 1974)]. Hence, PEMT1 is probably fully saturated with SAM whereas PEMT2 is not. The gene for human PEMT2 has been localized to chromosome 17p11.2 (Walkey et al., 1999), and cDNA for PEMT2 has been cloned from rat liver and expressed in COS‐1 cells (Cui et al., 1993). PEMT activity was identified in membranous fractions from homogenates of rat and bovine brain (Blusztajn et al., 1979; Mozzi and Porcellati, 1979; Crews et al., 1980); highest specific activies were present in synaptosomes (Blusztajn et al., 1979; Crews et al., 1980) suggesting that nerve terminals are able to synthesize choline. In the course of these transmethylations, the phosphatide intermediates ‘‘flip’’ from the membrane’s cytoplasmic side, where most of the less‐polar PE and phosphatidylserine (PS) are found, to the more polar external leaf (Hirata and Axelrod, 1978). PE itself can be formed in liver, kidney, or brain from free ethanolamine, via the CDP‐ethanolamine cycle (or ‘‘Kennedy Cycle’’; > Figure 18‐4) described later, or from the decarboxylation of PS (Kennedy, 1956; Borkenhagen et al., 1961). PS is produced, in nerve terminals (Holbrook and Wurtman, 1988) and elsewhere, by the process of ‘‘base‐exchange,’’ in which a serine molecule substitutes for the ethanolamine in PE or the choline in PC. Free choline is liberated from newly synthesized PC and from PC molecules formed from preexisting choline, by a family of enzymes, the phospholipases (> Figure 18‐5). Phospholipase D (PLD) acts directly on the choline/phosphate bond of PC to generate choline and phosphatidic acid (> Figure 18‐5a). Phospholipase A2 (PLA2) acts on the bond connecting a fatty acid to the hydroxyl group on PC’s 2‐carbon to yield that fatty acid [often arachidonic acid (AA) or docosahexaenoic acid (DHA)] and lyso‐PC (> Figure 18‐5b). This lyso‐PC can then be further metabolized to choline, either directly, through the action of a phosphodiesterase, or first to GPC, by phospholipase A1 (PLA1), and then to choline by a phosphatase. Phospholipase C (PLC) acts on the bond connecting the phosphate and the hydroxyl group on PC’s 3‐carbon to yield diacylglycerol (DAG) and phosphocholine; the phosphocholine can then be metabolized to free choline through the action of a phosphatase (> Figure 18‐5c). . Figure 18‐4 Biosynthesis of phosphatidylcholine (PC) from preexisting choline through the CDP‐choline cycle (‘‘Kennedy cycle’’). Synthesis of PC is shown here. Synthesis of phosphatidylethanolamine (PE) is similar except that it uses ethanolamine instead of choline. Boxes indicate compounds most or all of which must be taken up into the brain from the circulation. Uridine is the principal circulating precursor of the CTP (cytidine‐50 ‐triphosphate) needed for PC and PE synthesis through the Kennedy pathway, and exogenous cytidine is rapidly deaminated to uridine in humans (Wurtman et al., 2000). CK, Choline Kinase; CT: CTP: phosphocholine cytidylyltransferase; CPT: CDP‐choline, 1,2‐diacylglycerol cholinephosphotransferase; PUFA, Polyunsaturated fatty acid; DHA, Docosahexaenoic acid; EPA, Eicosapentaenoic acid; AA, Arachidonic acid

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It is estimated that, on average, about 15% of the free choline that enters the human blood stream derives from endogenous synthesis, the rest coming principally from dietary sources (Zeisel, 1981). Acute or chronic liver disease or deficiencies in methionine, folic acid, or vitamin B12 intake could thus lower plasma choline levels by impairing hepatic PC synthesis (> Table 18‐4).

2.1.3 Choline‐Containing Membrane Phospholipids Cellular membranes contain most of the choline in the body, principally in the form of the phosphatide PC, but also as PC’s products SM (> Figure 18‐6) and lyso‐PC (> Figure 18‐5b), or as less‐abundant choline‐ containing phospholipids like the PAF (1‐O‐alkyl‐2‐acetyl‐sn‐glycero‐3‐phosphocholine). Membranes also contain the phosphatides PS, PE, and phosphatidylinositol (PI), as well as specific proteins, cholesterol, and various minor lipids. The quantities of choline present in brain as PC (2–2.5 mmoles/g) or as SM (0.25 mmoles/g) are orders of magnitude greater than those of free choline (30–60 mM). The proportion of any membrane’s phospholipids represented by PC can vary depending on the species and age of the animal, the particular brain region or cell type being studied, and the membrane’s function within the cell (e.g., nuclear membrane and plasma membrane) (Suzuki, 1981). In the gray matter of human brain, PC constitutes 42% of total phospholipids and SM 10%; in white matter, the proportions of PC and SM are 33% and 15%, respectively (Suzuki, 1981). Moreover, ‘‘PC’’ is highly heterogeneous, actually representing a family of compounds with differing fatty acid compositions (> Figure 18‐3; Lee and Hajra, 1991) and, consequently, differing chemical and physical properties. The fatty acid in the C‐1 position of PC tends most often to be saturated, e.g., stearic or palmitic acid, whereas that in position C‐2 is more likely to be monounsaturated (oleic acid) or polyunsaturated [e.g., the omega‐3 fatty acids (DHA; 22:6) and eicosapentenoic acid (EPA; 20:5) or the omega‐6 fatty

. Figure 18‐5 Phospholipases that metabolize phosphatidylcholine (PC). (a) Phospholipase D; (b) Phospholipase A2; (c) Phospholipase C. Boxes surrounding the portions of the PC molecule differentiate the glycerol, fatty acid (R1 and R2), and choline moieties. Lyso‐PC, Lyso‐Phosphatidylcholine; LPCAT, Lyso‐Phosphatidylcholine acyltransferase; GPC‐PD, Glycerophosphocholine phosphodiesterase; GPC‐CPD, Glycerophosphocholine cholinephosphodiesterase; DAG, Diacylglycerol

Choline and its products acetylcholine and phosphatidylcholine . Figure 18‐5 (continued)

18

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Choline and its products acetylcholine and phosphatidylcholine

. Figure 18‐5 (continued)

acid (AA; 20:4)]. Newly synthesized phosphatide molecules contain relatively larger quantities of polyunsaturated fatty acids (PUFA) than the phosphatide molecules present at steady state (Tacconi and Wurtman, 1985). This reflects either faster turnover of PUFA‐containing phosphatides, or their rapid deacylation followed by reacylation with more saturated fatty acid species (Houtsmuller, 1979), or both. Membranes of retinal and brain cells are especially rich in PUFA, particularly DHA [which comprises about 20% of the total fatty acids in retinal phospholipids (Futterman and Andrews, 1964; Martinez, 1992) and about 7% of those in brain phospholipids (Martinez, 1992), respectively]. As described later, administration of supplemental DHA accelerates PC synthesis and increases brain levels of PC and other phosphatides (Wurtman et al., 2006).

2.2 Fates of Circulating Choline Plasma choline can be taken up into the brain and other tissues for further metabolism, or, as discussed later, oxidized—principally in liver and kidney—to form betaine, a source of methyl groups. This latter process involves two enzymes, choline dehydrogenase and betaine dehydrogenase, usually collectively termed ‘‘choline oxidase’’ (> Figure 18‐2). Negligible choline dehydrogenase activity ( Table 18‐1) by glomerular filtration followed by partial renal tubular reabsorbtion. Dietary choline or choline secreted into the gut can be broken down by intestinal bacteria to form trimethylamine and related amine products (de la Huerga and Popper, 1951; Neill et al., 1978). This process is responsible for the ‘‘fishy odor’’ sometimes detected in people taking large doses of choline supplements (Rehman, 1999).

2.2.1 Choline as a Source of Methyl Groups (Choline Oxidase System) The choline oxidase system in mammals is composed of two enzymes; in microorganisms a single enzyme, also termed ‘‘choline oxidase’’ (EC 1.1.3.17) converts choline to betaine. In mammals, the choline is first oxidized to betaine aldehyde by choline dehydrogenase (EC 1.1.99.1) (> Figure 18‐2), an enzyme located at (or bound to) the inner membrane of mitochondria (Streumer‐Svobodova and Drahota, 1977; Lin and Wu, 1986). This enzyme can also convert the aldehyde to betaine; however, unlike the choline oxidase of microorganisms, its affinity for betaine aldehyde is very low (only about 5% its affinity for choline), so choline dehydrogenase has only a minor effect on net betaine synthesis (Tsuge et al., 1980). This enzyme is a monomeric flavoprotein with a molecular weight of 61.000 Da (Lin and Wu, 1986); its activity requires FAD (Rothschild et al., 1954) and molecular oxygen serves as the primary electron acceptor (Zhang et al., 1992). One molecule of choline oxidized through the respiratory chain yields two molecules of mitochondrial ATP (Lin and Wu, 1986). Choline dehydrogenase has been cloned from rat liver mitochondria using a cDNA formed from the enzyme’s terminal amino acid sequence (Huang and Lin, 2003). The enzyme is most active in liver and kidney (Bernheim and Bernheim, 1933; Mann and Quastel, 1937), and, as discussed earlier, only negligible choline dehydrogenase activity is observed in brain (Haubrich et al., 1979; Haubrich and Gerber, 1981). Recent estimates of choline dehydrogenase’s Km for choline—140–270 mM (Zhang et al., 1992)— suggest values that are substantially lower than those estimated earlier (5–7 mM; Rendina and Singer, 1959; Tsuge et al., 1980; Haubrich and Gerber, 1981). However, this revised Km is still high compared with actual liver choline concentrations (60–230 mM; Sundler et al., 1972; Haubrich and Gerber, 1981), suggesting that treatments that increase hepatic choline levels also increase its rate of degradation. Very high portal venous choline concentrations (2.5 mM; Zeisel et al., 1980b) produced experimentally in studies on isolated perfused livers can fully saturate the enzyme. Betaine aldehyde dehydrogenase (EC 1.2.1.8) further oxidizes betaine aldehyde to betaine (> Figure 18‐2). This enzyme, found both in cytoplasm and mitochondria (Wilken et al., 1970; Pietruszko and

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Chern, 2001), uses NAD as a cofactor (Wilken et al., 1970). Its Km for betaine aldehyde (118 mM in rat liver mitochondria, Chern and Pietruszko, 1995; 123 mM in rat liver cytoplasm, Vaz et al., 2000; 182 mM in human liver cytoplasm, Vaz et al., 2000) is probably substantially higher than actual in vivo concentrations of the aldehyde; hence, the betaine aldehyde formed when hepatic choline levels rise is rapidly metabolized. Choline’s oxidation occurs through two irreversible reactions; hence, betaine cannot be reduced back to choline. Choline administration enhances betaine formation both in vivo (Haubrich et al., 1975; Zeisel and Wurtman, 1981) and in vitro (Wilken et al., 1970; Zeisel et al., 1980b; Zhang et al., 1992). In isolated perfused liver, betaine accounts for 60% of the labeled metabolites of [methyl‐14C]choline at concentrations of 5–125 mM (Zeisel et al., 1980b). Betaine is used by liver as a source of methyl groups for the generation of methionine from homocysteine, a process catalyzed by the enzyme betaine homocysteine methyltransferase (EC 2.1.1.5; Klee et al., 1961). The importance of this pathway (and thus of choline as a source of methyl groups) is probably increased when other methylating pathways have been compromised by ethanol ingestion, drugs, or nutritional deficiencies affecting folic acid, pyridoxine, or vitamin B12 (Barak and Tuma, 1983). Thus, treatment with betaine could be beneficial in early‐stage alcoholic liver injury (Barak et al., 1996; Kharbanda et al., 2005).

2.3 Effects of Physiologic or Pathologic States on Plasma Choline In adult humans, plasma (or serum) concentrations of free choline are maintained around 10 mM after a 6–24 h fast (> Table 18‐4), and increased by up to 50 mM postprandially, depending on the choline contents of the consumed food (Hirsch et al., 1978; Zeisel et al., 1980c). Basal plasma concentrations of free choline decrease by about 25–30% in human subjects undergoing a 1‐week fast (Savendahl et al., 1997; da Costa et al., 2004) or consuming a choline‐deficient diet ( Table 18‐4). These concentrations are much higher in newborns (about 40 mM) than in adult humans (Zeisel et al., 1980a; Zeisel and Wurtman, 1981; Buchman et al., 2001; Ilcol et al., 2002e, 2005a); a similar age relationship has been described in rats (Zeisel et al., 1980a; Zeisel and Wurtman, 1981) and rabbits (Zeisel et al., 1980a). In rats, the high postnatal choline concentrations fall to adult levels during the first 3 weeks of postnatal life (Zeisel and Wurtman, 1981); in humans, this occurs by 2 years of age (Ilcol et al., 2005a). In humans, serum‐free choline levels rise gradually during pregnancy (Ilcol et al., 2002e; Velzing‐Aarts et al., 2005) reaching 15–20 mM at term (Ulus et al., 1998; Ilcol et al., 2002e, 2005a); then they decrease by 35– 40% during the 12–20 h after delivery (Ulus et al., 1998; Ilcol et al., 2002e). Like pregnancy, breast‐feeding is associated with elevated serum‐free choline levels, in the mothers, rising to 15–20 mM during a 180‐day period of lactation. In both pregnancy and lactation, considerable amounts of free choline are transferred from the mother to the fetus or breast‐fed infant, through the placenta or the breastmilk, and used for growth and development (e.g., as a precursor of membrane phosphatides; Zeisel, 2006). The high‐serum‐free choline concentrations observed in pregnant and lactating women may reflect a process for promoting fetal and infant growth at the expense of depleting the mother’s choline stores (Zeisel et al., 1995). In patients with end‐stage renal disease, plasma‐free choline levels are several‐folds higher than those in control subjects (Rennick et al., 1976; Buchman et al., 2000b; Ilcol et al., 2002a, b) or in patients who have successfully undergone renal transplantation (Ilcol et al., 2002a). Considerable amounts of choline are lost into hemodialysates (Rennick et al., 1976; Buchman et al., 2000b; Ilcol et al., 2002a) or peritoneal dialysates (Ilcol et al., 2002b), but plasma‐free choline levels decrease only slightly during hemodialysis (Rennick et al., 1976; Buchman et al., 2000b; Ilcol et al., 2002a). Several studies have shown that plasma‐free choline concentrations decrease significantly by about 25–40% after prolonged exercise, e.g., running a marathon (Conlay et al., 1986, 1992; Buchman et al., 1999, 2000a), and remain depressed for at least 48 h after the race (Buchman et al., 1999).

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Serum‐free choline concentrations decrease by 20–45% during (Ulus et al., 1998; Ilcol et al., 2002d) and after surgery, in humans (Ulus et al., 1998; Ilcol et al., 2002d, 2004, 2006) or dogs (Ilcol et al., 2003b). This phenomenon is a response to surgical stress and is inversely correlated with the stress‐induced elevations in serum cortisol, adrenocorticotropic hormone (ACTH), prolactin, and b‐endorphin (Ilcol et al., 2002a). The magnitude and duration of surgery‐induced declines in serum choline depend on the severity and the type of surgery (Ulus et al., 1998; Ilcol et al., 2002d, 2003b, 2005b). Thus, free choline concentrations return to presurgical values within 24 or 48 h after a cesarean section (Ilcol et al., 2002e), or a transurethral prostatectomy (Ulus et al., 1998), but require 72 h to do so after abdominal surgery (Ilcol et al., 2003b) or 96 h after coronary artery bypass surgery (Ilcol et al., 2004, 2006) or removal of a brain tumor (Ilcol et al., 2004). The decline in serum‐free choline concentration associated with surgery can be mimicked in dogs by the administration of methylprednisolone (Ilcol et al., 2003b). Perhaps paradoxically, plasma and whole blood choline concentrations reportedly increase significantly in patients with acute coronary syndromes (Danne et al., 2003, 2005).

3

Choline in the Brain

Because choline is, by virtue of its quaternary nitrogen atom, relatively polar, it had generally been assumed (Ansell and Spanner, 1971; Diamond, 1971) that plasma choline was unavailable to the brain. Moreover, as brain cells were also thought to be incapable of synthesizing choline de novo, the ability of cholinergic neurons to maintain the intracellular choline concentrations needed for ACh synthesis was usually attributed either to an extraordinarily effective reuptake mechanism, described later, for reutilizing virtually all the choline formed from the hydrolysis of ACh, or, less likely, to the uptake into brain of circulating PC or lyso‐ PC (Illingworth and Portman, 1972; Kuhar and Murrin, 1978). In addition, since the poor affinity of ChAT, the enzyme that catalyzes choline’s conversion to ACh for choline made it likely that intracellular choline concentrations would control brain ACh synthesis; it was broadly conjectured that choline’s high‐affinity uptake from the synaptic cleft controlled ACh synthesis (cf., Taylor and Brown, 2006). It is no longer held that the brain choline levels are sustained solely by the high‐affinity uptake of free choline from synapses, or that variations in this uptake are normally responsible for observed variations in brain choline levels. Choline molecules (but not those of PC or lyso‐PC; Pardridge et al., 1979) do readily cross the BBB (Cornford et al., 1978), and brain cells do indeed synthesize choline de novo (Blusztajn and Wurtman, 1981). Physiological variations do occur in choline levels within brain neurons; however, these result principally from changes in plasma choline concentrations after eating choline‐rich foods, or from choline’s metabolism. It is possible in laboratory studies to make the reuptake of intrasynaptic choline become the limiting factor controlling ACh synthesis, for example, by giving a drug‐like hemicholinium‐3 (HC3), which blocks the reuptake process. However, no food constituents or endogenous compounds have ever been shown to share this ability. It is possible that the density of choline‐reuptake sites in nerve terminals may be modulated by the rate of ACh release (Taylor and Brown, 2006); however, variations in the rate of ACh release have not been demonstrated to affect the efficiency of choline reuptake. Mammalian brains contain choline as the free base; as such water‐soluble phosphorylated metabolites as phosphocholine and GPC (Nitsch et al., 1992), and as constituents of membrane phospholipids including PC, SM, and lyso‐PC. Free choline levels in the brains of humans and rats reportedly vary between 36–44 mM (Ross et al., 1997) and 30–60 mM (Stavinoha and Weintraub, 1974; Klein et al., 1993), whereas PC and SM levels are orders of magnitude higher (2000–2500 mM and 250 mM, respectively; Marshall et al., 1996). These high levels reflect the ubiquity of phospholipids, and the numerous essential roles they mediate when they form membranes. Membrane phospholipids also serve as reservoirs for choline and for such ‘‘second messenger’’ molecules as DAG, AA, inositol trisphosphate (IP3), and phosphatidic acid.

3.1 Sources of Brain Choline Free choline molecules in the brain derive from four known sources such as uptake from the plasma; liberation from the PC in brain membranes; high‐affinity uptake from the synaptic cleft after ACh released

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from a cholinergic terminal has been hydrolyzed; and, probably to a minor extent, the breakdown of newly synthesized PC formed from the methylation of PE.

3.1.1 Uptake of Circulating Choline into the Brain The brain can obtain circulating choline and various other circulating nutrients (e.g., neutral and basic amino acids, glucose, adenine, or adenosine; Pardridge and Oldendorf, 1977; Pardridge, 1986) via two routes: Small amounts can pass from the blood to the cerebrospinal fluid through the action of a specific transport protein, organic cation transporter 2 (OCT2), which is present in cells that line the choroid plexus (CP) (Sweet et al., 2001). However, orders of magnitude more pass bidirectionally between the blood and the brain’s extracellular fluid (ECF) by facilitated diffusion. This process is catalyzed by a different transport protein, not yet cloned, which is localized within the endothelial cells that line the brain’s capillaries (Oldendorf and Braun, 1976; Pardridge and Oldendorf, 1977; Cornford et al., 1978). Its action is independent of sodium, and can be blocked by HC3. Studies using an in situ brain perfusion technique, or a cell line of immortalized endothelial microvessels from rat brain (RBE4), have demonstrated the existence of a transport protein with a relatively low Km for choline [(39–42 mM; Allen and Smith, 2001) or (20 mM; Friedrich et al., 2001)] which could mediate choline’s bidirectional flux across the BBB. Other investigators using other experimental systems had proposed substantially higher Kms for endothelial choline transport, i.e., 220–450 mM (Oldendorf and Braun, 1976; Pardridge and Oldendorf, 1977; Cornford et al., 1978; Mooradian, 1988). The differences among the affinities noted in these studies might, as discussed later, reflect the different methods used for their measurement. However, in any case, the protein would still be unsaturated at physiological plasma choline concentrations, and its net activity is still affected by variations in these concentrations. It might constitute a kind of pore through which choline can pass in either direction, based on the gradient between its blood and brain levels (Klein et al., 1990). Hence when plasma choline levels have been elevated, for example by eating a choline‐rich meal (e.g., to 50 mM in the rat; Zeisel et al., 1980a), choline tends to enter the brain, but when plasma choline levels are low its flux is in the opposite direction. It has been estimated that the plasma choline concentration in rats required in order for the net choline flux to be from blood to brain is about 15 mM; below this concentration, net choline flux presumably is from brain to blood (Klein et al., 1990). No endogenous circulating compound has been shown to compete effectively with choline for facilitated diffusion across the BBB. Very high concentrations of carnitine and spermidine, compared with those in the blood, can reduce brain uptake of choline by 20–25% (Cornford et al., 1978). One drug, diethylaminoethanol, apparently does block BBB choline uptake, and has been used to lower brain choline levels and thereby suppress ACh synthesis (Cornford et al., 1978; Millington et al., 1978). Lithium ion, given acutely (Cornford et al., 1978) or chronically (Millington et al., 1978), may also block BBB choline uptake. However, lithium also blocks choline’s efflux from brain to blood, thus producing a net increase in brain choline levels (Millington et al., 1978). Once circulating choline has entered the brain’s ECF, it can be taken up into all cells by a low‐affinity transport protein (Km ¼30–100 mM), or into cholinergic nerve terminals by a high‐affinity uptake protein (Km ¼0.1–10 mM) (Haga and Noda, 1973; Yamamura and Snyder, 1973; Blusztajn and Wurtman, 1983). Both of these are described later. The high‐affinity process, unlike the passage of choline across the BBB, is energy‐ and sodium‐dependent.

3.1.2 Liberation from Membrane PC The choline in membrane PC can be liberated through the actions of the phospholipase enzymes, described earlier, which catalyze the hydrolysis of various bonds between PC’s three oxygen molecules and fatty acids or its phosphate moiety (> Figure 18‐5). The activation of each of these enzymes is tightly regulated and, in general, initiated by the interaction of a neurotransmitter or other biologic signal with a receptor coupled to a G‐protein. For example, both the PLC enzymes (which act on PC to yield DAG and phosphocholine, or on PI) and PLD (which acts on PC to yield phosphatidic acid and choline) are activated when ACh attaches to M1 or M3 muscarinic receptors (Sandmann and Wurtman, 1990, 1991; Sandmann et al., 1991).

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The DAG generated by PLC activates a family of protein kinase (PK) enzymes that phosphorylate various proteins, including those that control the metabolism of the amyloid precursor protein (APP) to form either soluble APP or the A‐beta peptides (Hung et al., 1993; Nitsch et al., 1994; Slack et al., 1997; Slack and Wurtman, 2006). The release of choline from PC can also be enhanced, and its reincorporation into PC is diminished by sustained neuronal depolarization (Farber et al., 1996). This process has been termed ‘‘autocannibalism’’ when some of the choline is diverted for the synthesis of ACh (Blusztajn et al., 1986; Ulus et al., 1989). Autocannibalism may, by decreasing the quantities of phosphatide molecules, and thus of neuronal membranes, underlie the particular vulnerability of cholinergic neurons in certain diseases (Blusztajn et al., 1986; Ulus et al., 1989). It is not known whether the accelerated breakdown of PC associated with sustained neuronal depolarization results from changes in ion flux or requires the release of local neurotransmitters and activation of particular receptors. The depletion of membrane PC and other phosphatides—including those not containing choline—caused by frequent or sustained depolarizations can be diminished or blocked entirely, and the release of ACh is enhanced by providing the brain with supplemental choline (Ulus et al., 1989).

3.1.3 Reutilization of Choline Formed from Hydrolysis of Acetylcholine ACh released into synapses is rapidly hydrolyzed to free choline and acetate. This process terminates the neurotransmitter’s physiologic actions, i.e., its ability to combine with and activate its pre‐ or postsynaptic muscarinic or nicotinic receptors. (The inactivation of ACh differs from that of other aminergic transmitters, e.g., dopamine and serotonin, in which it involves a chemical change in the neurotransmitter molecule, and not simply physical removal of that molecule from the synaptic cleft by reuptake into its nerve terminal of origin.) The enzymes that catalyze ACh hydrolysis, the acetylcholinesterases (EC 3.1.1.7; AChE), are particularly abundant within the cholinergic synapse; they are synthesized in the cholinergic neuron and secreted into the synapse, along with ACh, when the neuron is depolarized. A related enzyme, butyrylcholinesterase (EC 3.1.1.8; BuChE), synthesized in the liver and present in plasma, probably functions to inactivate potentially toxic dietary esters but not intrasynaptic ACh: It is active in the nervous system during development, but not thereafter, and mutant animals lacking the BuChE gene—in contrast to those lacking AChE—apparently fail to exhibit neurologic symptoms (Taylor and Radic, 1994; Giacobini, 2003). Most of the free choline liberated by the intrasynaptic hydrolysis of ACh is taken back up into its nerve terminal of origin by the high‐affinity choline transporter (CHT) described later, and either reacetylated to form ACh or phosphorylated for ultimate conversion to membrane PC (Ulus et al., 1989).

3.1.4 De Novo Synthesis of Phosphatidylcholine and Choline As described earlier, brain cells—including nerve terminals (Holbrook and Wurtman, 1988)—contain all the enzymes needed to synthesize PC from ethanolamine (> Figure 18‐3) or from PS. These include the Kennedy cycle enzymes that convert ethanolamine to PE (Spanner and Ansell, 1979), PS decarboxylase, which forms PE from PS (Butler and Morell, 1983), and the enzymes (PEMT1 and PEMT2), which methylate PE (Crews et al., 1980).

4

Brain Proteins that Interact with Choline

Free choline is known to interact with two brain enzymes and four transport proteins, as well as various receptors for ACh.

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The two enzymes are ChAT and CK—which catalyze, respectively, the transformations of choline to ACh within cholinergic terminals, and to phosphocholine within all cells. The four transport proteins include two that move choline across the BBB, i.e., the facilitated‐diffusion site in brain capillaries through which choline passes, bidirectionally, between the plasma and the brain’s ECF, and the organic cation transporter that carries plasma choline across the CP and into the cerebrospinal fluid; and two that enable the choline in brain ECF to enter cells, i.e., the low‐affinity uptake site that catalyzes choline’s uptake into all brain cells, and the high‐affinity uptake site that transports intrasynaptic choline into the presynaptic terminals of cholinergic neurons. The cholinergic receptors, which choline can also activate, include both nicotinic and muscarinic varieties. This section describes the properties of each of these proteins, and the consequences of their interactions with choline.

4.1 Enzymes 4.1.1 Choline Acetyltransferase ChAT (acetyl‐CoA: choline‐O‐acetyltransferase, EC 2.3.1.6) mediates a single reaction, the transfer of an acetyl group from acetyl‐coenzyme A (acetyl‐coA) to choline, which thereby generates the neurotransmitter ACh in cholinergic neurons. ChAT, a single‐stranded globular protein, is encoded by a single gene with, in humans, six distinct transcripts formed from the alternative splicing of five noncoding exons (Misawa et al., 1992, 1997; Oda et al., 1992; Robert and Quirin‐Stricker, 2001) Polymorphism among these transcripts is apparently limited to their 50 ‐untranslated regions. In humans, four of the six transcripts (designated as H, R, N1, and N2) translate to the same 69‐kD protein (Misawa et al., 1992, 1997; Oda et al., 1992; Robert and Quirin‐Stricker, 2001). The fifth and sixth transcripts, designated as M and S, have two translation sites and yield, besides the 69‐kD enzyme, 82‐kD and 74‐kD forms of ChAT, respectively. The 82‐kD ChAT differs from the 69‐kD form in that it has an aminoterminal extension with 118 amino acid residue (Oda et al., 1992; Misawa et al., 1997). Physiological roles for the 74‐kD and 82‐kD forms of ChAT remain to be elucidated, and indeed it is not clear that these larger forms of human ChAT actually are synthesized in vivo (Oda, 1999). ChAT probably exists in at least two forms within cholinergic nerve terminals—a soluble form (80–90% of the total enzyme activity) and a membrane‐associated form (10–20%; Benishin and Carroll, 1981; Salem et al., 1994; Pahud et al., 1998). These two forms exhibit different physicochemical and biochemical properties (Benishin and Carroll, 1983; Eder‐Colli et al., 1986; Pahud et al., 2003). The soluble form is hydrophilic, and the membrane‐bound form is amphiphilic (Benishin and Carroll, 1983; Eder‐Colli et al., 1986; Pahud et al., 2003). Soluble ChAT has higher affinities for both of its substrates, choline and acetyl‐ CoA, when assayed at low ionic strength (Km for choline 350 mM; for acetyl‐CoA 2.5 mM) than that when assayed at higher ionic strengths (Km for choline 6700 mM; for acetyl‐CoA 77 mM; Rossier, 1977). The activity of ChAT in crude synaptosomal preparations (presumably representing a mixture of the soluble and membrane‐bound forms) also varies with ionic strength; the affinity of synaptosomal ChAT for choline appears to be greater than that of soluble ChAT (Km ¼22 mM at low ionic strength and 540 mM at high ionic strength; Rossier, 1977). In any case, ChAT is invariably unsaturated with choline at the choline concentrations that could exist within nerve terminals (Tucek, 1990), indicating that ChAT is in kinetic excess (Hersh, 1982; Tucek, 1990), and that its substrate‐saturation, not its levels, is rate limiting in ACh synthesis. Both choline and acetyl‐CoA (Rossier, 1977; Hersh, 1982; Tucek, 1990) levels can affect the rate at which ACh is produced. There is also evidence that phosphorylation and dephosphorylation of ChAT can alter its catalytic activity, subcellular distribution, and interactions with other cellular proteins (see review of Dobransky and Rylett, 2005). ChAT is a substrate for multiple PKs; 69 kDa ChAT is phosphorylated by PK‐C, PK‐CK2, and a Ca2þ/calmodulin‐dependent PK‐II (CaM‐kinase) but not by PK‐A, whereas 82 kDa ChAT is phosphorylated by PK‐C and CaM‐kinase (Dobransky et al., 2000, 2001). ChAT is differentially phosphorylated by PK‐C isoforms on four of its serine residues (Ser‐440, Ser‐346, Ser‐347, and Ser‐476) and one threonine

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residue (Thr‐255); this phosphorylation is hierarchical, such that phosphorylation at Ser‐476 is required in order for the other serines to become phosphorylated (Dobransky et al., 2004). Phosphorylation at some but not all of those sites (Ser‐476 with Ser‐440 and Ser‐346/347; Dobransky et al., 2004) affects basal ChAT activity. Phosphorylation of ChAT by PK‐C alone can double the enzyme’s activity, whereas coordinated phosphorylation of ChAT at threonine 456 (by CaM‐kinase II) and serine 440 (by PK‐C) can treble ChAT activity (Dobransky et al., 2003). Whether the phosphorylation and dephosphorylation of ChAT also alter the enzyme’s affinities for choline or acetyl‐CoA in intact cells is clear.

4.1.2 Choline Kinase CK (ATP:choline phosphotransferase; EC 2.7.1.32) catalyzes the first phosphorylation reaction in the Kennedy cycle of PC synthesis (> Figure 18‐4); ATP is the phosphate donor and the presence of Mgþ2 is required (Wittenberg and Kornberg, 1953). CK can also catalyze the phosphorylation of ethanolamine, as well as N‐monomethylethanolamine and N,N‐dimethylethanolamine (Ishidate et al., 1985; Porter and Kent, 1990; Uchida and Yamashita, 1990); however, a separate ethanolamine kinase enzyme exists, demonstrated by cloning cDNA from human liver (Lykidis et al., 2001). CK is mainly cytosolic but is also associated with particulate (membrane‐bound) fractions of rat striatum (Reinhardt and Wecker, 1983). The enzyme has been purified to homogeneity from various rat tissues (Ishidate et al., 1985; Porter and Kent, 1990), including brain (Uchida and Yamashita, 1990). HC3 (Ansell and Spanner, 1974) and ADP (Burt and Brody, 1975) inhibit CK activity in vitro, whereas high concentrations of the polyamines spermine and spermidine (Uchida and Yamashita, 1990) enhance its activity. Several isoforms of CK exist in brain and other tissues, differentiable by their subunit masses (Porter and Kent, 1990; Uchida and Yamashita, 1990) and by cloning and expression studies (Uchida and Yamashita, 1992; Uchida, 1994; Aoyama et al., 1998). At least as three isoforms (CK‐a1, CK‐a2, and CK‐b), encoded by two separate genes termed ck‐a and ck‐b (Aoyama et al., 1998, 2000, 2004), are now recognized. The latter resides on chromosome 22q13 in humans (Froguel and McGarry, 1997); the locus of the former awaits determination. These isoforms may not be active as monomers, but become active on forming dimeric or oligomeric structures (Aoyama et al., 2004). In some circumstances, CK activity may be rate limiting in PC synthesis; for example, a 3.5‐fold increase in CK activity in livers of rats deficient in essential fatty acids was accompanied by a parallel increase in PC synthesis (Infante and Kinsella, 1978). Similar relationships have been described in livers of estrogen‐treated roosters (Vigo and Vance, 1981) or quiescent murine 3T3 cells in culture (Warden and Friedkin, 1985). However, it is probably not the activity of CK per se, but rather its degree of substrate saturation that affects the rate of PC synthesis. The Km of CK for choline in rat brain is 14–134 mM (Uchida and Yamashita, 1990; Cao and Kanfer, 1995); this value is 32–310 mM in rabbit brain (Haubrich, 1973). An even higher Km value (2.6 mM) was described by Spanner and Ansell (1979) who assayed the enzyme at a more physiological pH (7.5) than that customarily used (pH ¼ 9.0); this allowed phosphocholine, CK’s reaction product, to be assayed without first being hydrolyzed. Hence, CK is unsaturated with choline at normal brain choline concentrations (30–60 mM), and the production of phosphocholine through CK, like that of ACh by ChAT, is controlled principally by brain choline levels (Millington and Wurtman, 1982; Cohen et al., 1995).

4.2 Transport Proteins 4.2.1 Facilitated‐Diffusion Carrier at Blood–Brain Barrier A transport protein that mediates the bidirectional facilitated‐diffusion of choline at the BBB has been identified (Oldendorf and Braun, 1976; Pardridge and Oldendorf, 1977; Cornford et al., 1978). This protein does not require metabolic energy or sodium flux and cannot maintain a concentration gradient. The CHT at the BBB, as discussed earlier, might allow bidirectional passage of choline based on the gradient between its blood and brain concentrations (Klein et al., 1990). A blood choline concentration of

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15 mM has been estimated to be required for choline influx to the rat’s brain to predominate; below this concentration, choline efflux predominates (Klein et al., 1990). As discussed earlier, the BBB CHT’s Km for choline in vivo is estimated as 220–450 mM (Oldendorf and Braun, 1976; Pardridge and Oldendorf, 1977; Cornford et al., 1978; Mooradian, 1988). Hence, it is unsaturated at physiologic plasma choline concentrations (10 mM). The Km of BBB CHT for choline has been demonstrated as 39–42 mM in a recent study using an in situ brain perfusion technique (Allen and Smith, 2001); the higher affinity of the BBB transporter for choline found in this study, compared with previous data, probably reflects methodological differences. In vitro assays, using cell lines of immortalized rat (RBE4; Friedrich et al., 2001) and mouse (MBE4; Sawada et al., 1999) brain endothelial microvessels, have demonstrated the existence of a transport protein with a relatively low Km for choline (20 mM in both studies). However, these studies investigated uptake only at the luminal side of the endothelial cells, and not at both luminal and abluminal sides as was investigated in previous in vivo studies. The CHT at the BBB has not yet been cloned and awaits further characterization. A perhaps‐related transporter for carnitine, the organic cation/carnitine transporter OCTN2 (Tamai et al., 1998), has been cloned from RBE4 cells (Friedrich et al., 2003). Carnitine uptake through this transporter is not blocked by choline (Tamai et al., 1998).

4.2.2 Choroid Plexus Choline Transporter Much less blood choline is transported through the CP epithelium to the cerebrospinal fluid than through the BBB carrier to the brain’s ECF, because the surface area of the CP epithelium is much smaller than that of the BBB epithelium (Pardridge, 2001). Ventricular choline transport, in the rat, is mediated by one of the OCT proteins, OCT2 (Sweet et al., 2001). The three subtypes of OCTs (OCT1–3) have been isolated from rat (Grundemann et al., 1994; Okuda et al., 1996; Kekuda et al., 1998), mouse (Schweifer and Barlow, 1996; Mooslehner and Allen, 1999), and human tissues (Gorboulev et al., 1997; Zhang et al., 1997; Grundemann et al., 1998). OCTs are transmembrane proteins with 12 membrane‐spanning domains (Koepsell et al., 2003). Transport of a cation by an OCT protein is electrogenic, Naþ‐independent, and reversible with respect to direction (Koepsell and Endou, 2004). In the human, genes encoding OCT1–3 have been found on chromosome 6 (6q26–6q27; Koehler et al., 1997, 2003). The hOCT2 protein is expressed in kidney (Gorboulev et al., 1997) and brain (Busch et al., 1998); however, its localization in human brain ventricles, and its possible transport activity remain to be established.

4.2.3 High‐Affinity Uptake Protein in Cholinergic Terminals A saturable, sodium‐and energy‐dependent, HC3‐sensitive, high‐affinity CHT has been demonstrated in synaptosomes (Yamamura and Snyder, 1972, 1973; Guyenet et al., 1973; Haga and Noda, 1973). The Km of this transporter for choline is 0.1–10 mM (Guyenet et al., 1973; Haga and Noda, 1973; Yamamura and Snyder, 1973; Blusztajn and Wurtman, 1983). Choline uptake through the CHT is competitively inhibited by nanomolar concentrations of HC3 (Ki: 10–100 nM; Yamamura and Snyder, 1972; Haga and Noda, 1973; Kuhar and Murrin, 1978). The high‐affinity CHT protein, made up of two polypeptides with molecular masses of 58 and 35 kDa, has been identified and partially purified from rat corpus striatum (Rylett et al., 1996). cDNAs from rat (rCHT1; Okuda et al., 2000), mouse (mCHT1; Apparsundaram et al., 2001), and human (hCHT1; Apparsundaram et al., 2000) have also been isolated, cloned and expressed. These studies have shown that CHT1 does not belong to the neurotransmitter transporter family, but rather to the sodium‐dependent glucose transporter family (SLC5, in which CHT1 is designated as SLC5A7) (Apparsundaram et al., 2000; Okuda et al., 2000; Okuda and Haga, 2003). CHT1 protein has 13 transmembrane domains (Apparsundaram et al., 2000; Okuda et al., 2000). The human CHT1 gene is localized on chromosome 2q12 (Apparsundaram et al., 2000). High‐affinity choline transport occurs predominantly into terminals of cholinergic neurons (Misawa et al., 2001). Using antibodies raised against CHT1, CHT‐immunoreactive cells have been shown to be

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widely distributed throughout the rat, primate, and human central nervous systems (Misawa et al., 2001; Kus et al., 2003). Primate cerebellums contain numerous CHT‐immunoreactive cells (Kus et al., 2003), and mouse cerebellum expresses CHT1 mRNA, particularly during development (Berse et al., 2005). CHT1 is also present in terminals and in those of peripheral motor neurons (Lips et al., 2002; Nakata et al., 2004) and of parasympathetic neurons to the tongue (Lips et al., 2002). CHT1 is not expressed in glial cells (Inazu et al., 2005) but, contrary to what had been believed, is expressed in such nonneuronal cells as rat trachea (Pfeil et al., 2003), rat and human arteries (Lips et al., 2003), and skin (Haberberger et al., 2002). The function of CHT1 in these tissues awaits determination. Within neurons, CHT immunoreactivity is detectable in cell soma, proximal dendrites, axons and, particularly, axon terminals (Kus et al., 2003). Within the terminals, the CHT1 protein is especially abundant in plasma membrane, synaptic vesicles, and endosomal vesicles (Ferguson et al., 2003; Ribeiro et al., 2003; Ferguson and Blakely, 2004). In motor neurons of diaphragm, the CHT is mainly (>90%) concentrated within synaptic vesicles, rather than in the presynaptic membrane itself (Nakata et al., 2004). These vesicles may store CHT1 in the resting state, and the protein may migrate to the synaptic membrane during depolarization (Ribeiro et al., 2003, 2005). Activity‐dependent modulation of CHT1 has been described in studies using several different experimental systems, for example electrical or pharmacological stimulation of cholinergic neurons in vitro (reviewed in Ferguson and Blakely, 2004). The capacity and density of CHTs are apparently increased in medial prefrontal cortices of rats performing attentional tasks (Apparsundaram et al., 2005). Neuronal activity per se (Ferguson et al., 2003) might, by altering the phosphorylation state of CHT1 protein (Gates et al., 2004), enhance the transfer of CHT1 into and out of vesicles, thus modulating its activity. NGF, which can upregulate CHT1 through a PI3K‐dependent process, might similarly influence its activity (Berse et al., 2005).

4.2.4 Low‐Affinity Cellular Uptake Protein A nonsaturable, Naþ‐independent, high‐capacity, and low‐affinity CHT has also been identified (Haga and Noda, 1973; Yamamura and Snyder, 1973). Not surprisingly—since all cells need choline for phospholipid synthesis—it appears to be ubiquitous in mammals and is found, in, for example, kidney (Bevan and Kinne, 1990), liver (Zeisel et al., 1980), and placenta (Grassl, 1994) as well as in brain synaptosomes (Ferguson et al., 1991). The Km of the low‐affinity transport protein for choline varies between 30 and 100 mM (Dowdall and Simon, 1973; Haga and Noda, 1973; Yamamura and Snyder, 1973); it also is inhibited by HC3 with Kis of about 40–50 mM (Haga and Noda, 1973) in brain and 100 mM in human placenta (Grassl, 1994). Low‐affinity choline transport has been suggested to be a carrier‐mediated process (Ferguson et al., 1991; Inazu et al., 2005). As discussed later, CTL1, a member of the family of choline‐transporter‐like proteins (Traiffort et al., 2005), has been proposed as mediating the low‐affinity transport of choline into rat cortical astrocytes (Inazu et al., 2005) and mouse cortical neurons (Fujita et al., 2006). The choline‐transporter‐like proteins CTL1–CTL5 are encoded by five different genes, also labeled from CTL1 to CTL5 (Traiffort et al., 2005). CTL1, a transmembrane protein with 10 transmembrane domains, has been cloned and characterized from rat (rCTL1; O’Regan et al., 2000), human (hCTL1; Wille et al., 2001), and mouse (mCTL1; Yuan et al., 2004) tissues. The human gene is located on chromosome 9q31.2 (Wille et al., 2001), and its protein product is expressed as two polypeptides, of 50 and 23 kDa, which have been found in such tissues as brain, heart, small intestine, kidney, liver, lung, skeletal muscle, pancreas, spleen, ovary, and testis (Yuan et al., 2006). Another choline‐transporting system, the OCT proteins (members of the solute carrier family SLC22; Koepsell et al., 2003; Koepsell and Endou, 2004), have also been implicated in low‐affinity choline uptake. For example, rat OCT1 (rOCT1), cloned from renal proximal tubule epithelial cells or hepatocytes and expressed in Xenopus oocytes, can mediate low‐affinity choline uptake (Km ¼1.1 mM; Busch et al., 1996), and human OCT1 (hOCT1) and human OCT2 (hOCT2) can mediate, respectively, hepatic and renal choline transport (Km ¼210 mM; Gorboulev et al., 1997). It has not yet been determined whether the low‐affinity CHT is one of the OCT proteins; CTL1 (perhaps more likely in brain; Inazu et al., 2005; Fujita et al., 2006); or even a different protein.

Choline and its products acetylcholine and phosphatidylcholine

18

4.3 Receptors Choline in sufficiently high concentrations can directly activate both muscarinic (mAChRs) and the nicotinic (nAChRs) acetylcholine receptors. The five muscarinic receptors (M1–M5) mediate slow metabolic responses to ACh, and the nicotinic receptors, which are ligand‐gated ion channels, implement fast, ACh‐mediated synaptic transmission in the CNS, ganglia, and neuromuscular synapses. The M1, M3, and M5 muscarinic receptors activate phospholipase C, thereby generating the second messengers IP3 and DAG (Caulfield and Birdsall, 1998); the M2 and M4 muscarinic receptors inhibit adenylate cyclase activity, thus reducing intracellular cAMP levels, or can enhance the flux of potassium and other ions through nonselective ion channels. The nicotinic receptors, pentameric structures made up of combinations of 17 known individual subunits, increase the flux of sodium into postsynaptic cells, thus increasing the likelihood of the cells’ depolarization. Free choline concentrations in synaptic fluid following neuronal depolarization apparently have not been measured, and may or may not attain levels sufficient to activate cholinergic receptors under physiological circumstances. Much higher concentrations, produced experimentally, are readily shown to activate the receptors in vitro. Many years ago, it was noted that choline can produce ‘‘muscarine‐like’’ or ‘‘nicotine‐like’’ effects in various peripheral tissues (Dale, 1914; Chang and Gaddum, 1933), voluntary muscles (Bacq and Brown, 1937; Hutter, 1952; Del Castillo and Katz, 1957), and autonomic ganglia (Feldberg and Vartiainen, 1934; Kosterlitz et al., 1968; Krstic, 1972), with 1/20,000 to 1/714th the potency of ACh (Chang and Gaddum, 1933). Functional (Pomeroy and Raper, 1972; Ulus et al., 1979, 1988a; Holz and Senter, 1981) and receptor binding studies (Speth and Yamamura, 1979; Palacios and Kuhar, 1979; Costa and Murphy, 1984; Ulus et al., 1988), in which the choline presumably had not first been acetylated to authentic ACh, identified the effective choline concentrations needed to bind to AChRs and/or to produce ACh‐like biological responses. Choline, acting as a direct muscarinic agonist, excited cortical neurons (Krnjevic and Reinhardt, 1979); contracted isolated smooth muscle in rat stomach fundus (EC50¼0.41 mM), rat trachea (EC50¼1.7 mM), rat urinary bladder (EC50¼10.9 mM) (Ulus et al., 1988), and guinea pig ileum (EC¼0.6 mM; Pomeroy and Raper, 1972 or EC50¼0.20 mM; Ulus et al., 1979); and reduced the frequency at which isolated rat or guinea pig right atrium beat spontaneously (Ulus et al., 1979, 1988). It also inhibited ACh release from myenteric plexus‐longitudinal muscle preparations of guinea pig ileum (EC50¼0.3 mM) (Kilbinger and Kruel, 1981), and inhibited [3H]‐quinuclidinyl benzilate binding to rat brain membranes (Placios and Kuhar, 1979; Speth and Yamamura, 1979; Costa and Murphy, 1984; Ulus et al., 1988) and rat peripheral tissues (Ulus et al., 1988). Choline’s potency for inhibiting [3H]‐quinuclidinyl benzilate binding was found to vary among brain regions (Ki ¼0.46–3.5 mM) (Palacios and Kuhar, 1979; Speth and Yamamura, 1979; Costa and Murphy, 1984; Ulus et al., 1988) and also in peripheral tissues (Ki ¼0.28–1.17 mM) (Ulus et al., 1988). The wide range of variations in the muscarinic potency of choline and in its relative tissue selectivity (Ulus et al., 1988) may result from its varying affinities for mAChRs subtypes (M1–M5). Choline acts as a full agonist on human mutant M1 receptors to stimulate phosphoinositide hydrolysis (EC50¼0.2 mM; Huang et al., 1998), and on cloned human M1 receptor to stimulate nitric oxide synthesis and elevate intracellular Caþ2, at 0.1–1.0 mM concentrations (Carriere and El‐Fakanay, 2000). By activating nicotinic receptors as a full agonist (Ulus et al., 1988) and/or a partial agonist (Holz and Senter, 1981), choline stimulates catecholamine secretion from the vascularly perfused adrenal gland (EC50¼2.1 mM; Ulus et al., 1988), and from primary cultures of bovine adrenal chromaffin cells (at 1–10 mM; Holz and Senter, 1981). It also competes with L‐[3H]‐nicotine for binding to membrane preparations of rat brain (Costa and Murphy, 1984; Ulus et al., 1988) and peripheral tissues (Ulus et al., 1988). The potency of choline in displacing L‐[3H]‐nicotine from brain nicotinic receptors varies within a threefold range (Ki ¼ 379–1167 mM), and within a 1.5‐fold range for peripheral tissues (Ki ¼ 575–805 mM; Ulus et al., 1988). Patch‐clamp studies have shown that choline interacts with nAChRs in a ‘‘subtype selective’’ and concentration‐dependent manner. At concentrations of 0.1–10 mM, choline acts as a full agonist on a7 nAChR (Mandelzys et al., 1995; Papke et al., 1996, 2000; Alkondon et al., 1997, 2000; Albuquerque et al., 1998; Cuevas et al., 2000; Papke and Papke, 2002; Alkondon and Albuquerque, 2006; Gonzales‐Rubio et al., 2006) or a partial agonist on a3b4 nAChRs (Mandelzys et al., 1995; Papke et al., 1996; Albuquerque et al., 1998), a3b4*

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nAChRs (Seddik et al., 2003), and a4b4 nAChRs (Zwart and Vijverberg, 2000). It desensitizes a7nAChRs at 10–100 mM concentrations (Mandelzys et al., 1995; Papke et al., 1996, 2000, 2002; Albuquerque et al., 1997; Alkondon et al., 1997), inhibits a3b4* and a4b2* nAChRs at 10–1000 mM concentrations (Alkondon and Albuquerque, 2006), and potentiates or inhibits a4b4 nAChR‐mediated ACh currents at 10–300 mM or 1–30 mM concentrations, respectively (Zwart and Vijverberg, 2000).

5

Utilization of Choline in Brain

All cells use choline to produce the PC and SM in their membranes. Cholinergic neurons also use choline for an additional purpose, synthesis of their neurotransmitter, ACh. Both the PC and the ACh are ultimately broken down to regenerate free choline, thus both of these compounds can also be considered ‘‘reservoirs’’ for free choline. The synthesis of PC (> Figure 18‐4) is initiated by the phosphorylation of choline, catalyzed by an enzyme, CK, which forms phosphocholine by transferring a monophosphate group from ATP to the hydroxyl oxygen of the choline. As described later, this phosphocholine then combines with cytidine‐50 ‐triphosphate (CTP) to form cytidine‐50 ‐diphosphocholine (CDP‐choline), which, in turn, combines with DAG to yield PC. The synthesis of ACh, catalyzed by the enzyme ChAT, involves a single reaction, the transfer of an acetyl group from acetyl‐CoA, also to the hydroxyl oxygen of the choline. The ACh is then stored, largely within synaptic vesicles, for future release. Both CK and ChAT have low affinities for their choline substrate: Their Kms in brain, which describe the choline concentrations at which the enzymes operate at only half‐maximal velocity, may be as high as 2.6 mM (Spanner and Ansell, 1979) and 540 mM (Rossier, 1977), respectively, whereas brain choline levels, as noted earlier, are only about 30–60 mM, and thus well below the concentrations that would probably be needed to enable either enzyme to operate at maximal velocity. Hence, both of the enzymes are highly responsive to treatments that raise or lower brain choline levels. The ability of choline administration to increase the syntheses and brain levels of phosphocholine and ACh was first noted in 1982 (Millington and Wurtman, 1982) and 1975 (Cohen and Wurtman, 1975; Haubrich et al., 1975). It had previously been shown that the synthesis and levels of another brain neurotransmitter, serotonin, were increased if animals were given physiologic doses of its circulating precursor, tryptophan (Fernstrom and Wurtman, 1971; Cansev and Wurtman, 2006). This was because tryptophan hydroxylase, the enzyme that determines the overall rate at which tryptophan is converted to serotonin, has a low affinity for this substrate. Moreover, since ChAT’s affinity for choline had also been shown, in vitro, to be low, it seemed reasonable to enquire into whether choline’s ChAT‐mediated conversion to ACh also was precursor‐dependent. Once this relationship was affirmed, experiments soon followed demonstrating the precursor‐dependence of phosphocholine synthesis (Millington and Wurtman, 1982). Even though brain choline concentrations shared with those of tryptophan the ability to control the rates at which the precursor is of used for neurotransmitter synthesis, the two precursors differed in an important respect: Although tryptophan and choline are both used by certain neurons for two purposes: tryptophan for conversion to serotonin and incorporation into proteins, and choline for conversion to ACh and incorporation into phospholipids, in the case of tryptophan these two processes are segregated into different parts of the neuron—the nerve terminal and perikaryon, respectively—whereas for choline both can take place within the nerve terminal, inasmuch as that structure contains both ChAT and CK. Hence, the acetylation and phosphorylation of choline sometimes compete for available substrate (Farber et al., 1996; Ulus et al., 2006): When cholinergic neurons are forced to fire frequently and to sustain the release of ACh, choline’s incorporation into PC decreases (Farber et al., 1996) and the breakdown of membrane PC increases (‘‘autocannibalism’’), liberating additional choline for ACh synthesis (Maire and Wurtman, 1985; Blusztajn et al., 1986; Ulus et al., 1989). When the utilization of choline to form PC is increased (by providing supplemental uridine and an omega‐3 fatty acid; see later), ACh synthesis is not diminished, probably because so little choline is used for phosphatide formation compared with the amount used for ACh synthesis (Ulus et al., 2006).

Choline and its products acetylcholine and phosphatidylcholine

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5.1 Biosynthesis of Acetylcholine ACh is synthesized in cholinergic neurons—principally their terminals—by the ChAT‐mediated acetylation of free choline. Since, as described earlier, ChAT’s affinity for choline is low compared with brain choline levels, local choline concentrations normally control the rate of ACh synthesis (Blusztajn and Wurtman, 1983), and treatments which increase brain choline (e.g., administering choline; Cohen and Wurtman, 1975) or PC (Magil et al., 1981), or consuming choline‐rich foods (Cohen and Wurtman, 1976) rapidly cause parallel changes in brain ACh levels; in the amounts of ACh released when neurons fire (Maire and Wurtman, 1985; Jackson et al., 1995); and in postsynaptic ACh‐dependent functions like the control of rat striatal (Ulus and Wurtman, 1976) and adrenomedullary (Ulus et al., 1977a, b, c) tyrosine hydroxylase activities. The affinity of ChAT for its other substrate, acetyl‐CoA—formed from glucose in mitochondria—is substantially greater (Km ¼ 77 mM; Rossier, 1977) than that for choline (Km ¼ 540 mM), however actual acetyl‐CoA concentrations in the vicinity of ChAT may still be insufficient to saturate the enzyme, and thus might also affect the rate of ACh synthesis. In support of this possibility, administration of glucose has been found to stimulate ACh synthesis (Dolezal and Tucek, 1982), and to attenutate the depletion of brain ACh induced by giving a muscarinic antagonist (Ricny et al., 1992). In microdialysis studies, glucose enhanced the rise in ACh output produced by scopolamine (Ragozzino et al., 1994; Ragozzino and Gold, 1995). Systemic administration of glucose also increased hippocampal ACh release (Ragozzino et al., 1996, 1998; Kopf et al., 2001). If choline levels in nerve terminals are reduced pharmacologically by administering a drug, HC3 that blocks the reuptake of free choline from the synapse, the synthesis and release of ACh also decline in parallel (Maire and Wurtman, 1985). Although such experiments confirm the importance of choline availability in controlling ACh synthesis, they do not necessarily allow it to be concluded that high‐affinity choline uptake is the rate‐limiting factor controlling intracellular choline levels or ACh biosynthesis. This synthesis is affected by any process that modifies the neuron’s concentration of free choline, and these levels vary considerably as a function of plasma choline concentrations in addition, possibly, to changes in reuptake efficiency. Moreover, the choline that enters the neuron via high‐affinity uptake apparently is not selectively used for acetylation as opposed to phosphorylation (Kessler and Marchbanks, 1979; Jope and Jenden, 1981). As discussed earlier, it is possible, but not yet clearly demonstrated, that the density or activity of high‐affinity choline uptake sites in presynaptic membranes is affected by phosphorylation, neuronal firing, or the rate at which ACh is being released (Simon and Kuhar, 1975; Ferguson et al., 2003; Gates et al., 2004).

5.2 Biosynthesis of Choline‐Containing Phosphatides All cells use choline as an essential component of phospholipid subunits which, when aggregated, form all of their membranes. The principal subunit, the phosphatide PC, is synthesized from choline by the CDP‐ choline cycle (or ‘‘Kennedy Cycle’’; Kennedy and Weiss, 1956) (> Figure 18‐4); PC, in turn, provides the phosphocholine moiety for the synthesis of SM, the other major choline‐containing phospholipid (> Figure 18‐6). The CDP‐choline cycle involves three sequential enzymatic reactions (> Figure 18‐4): In the first (described earlier), catalyzed by CK, a monophosphate is transferred from ATP to the hydroxyl oxygen of the choline, yielding phosphocholine. The second, catalyzed by CTP:phosphocholine cytidylyltransferase (CT), transfers cytidylylmonophosphate (CMP) from CTP to the phosphorus of phosphocholine, yielding cytidylyldiphosphocholine (CDP‐choline). The third and last reaction, catalyzed by CDP‐choline:1,2‐DAG choline phosphotransferase (CPT), bonds the phosphocholine of CDP‐choline to the hydroxyl group on the 3‐carbon of DAG, yielding the PC. All these steps use compounds that the brain must obtain entirely or in part from the circulation, i.e., choline; a pyrimidine‐like uridine for conversion to CTP; a polyunsaturated fatty acid‐like DHA, and all three steps can also affect the overall rate of PC synthesis in brain (Cansev et al., 2005; Wurtman et al., 2006). Thus, choline administration increases brain phosphocholine levels in rats (Millington and Wurtman, 1982) and humans (Babb et al., 2004), because CK’s Km for choline (2.6 mM; Spanner and Ansell, 1979) is much

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higher than usual brain choline levels (35–60 mM). Most commonly, the second CT‐catalyzed reaction is most rate limiting, either because not all of the CT is fully activated by being attached to a cellular membrane (Vance and Pelech, 1984) or because local CTP concentrations are insufficient to saturate the CT (Ross et al., 1997). As described later, when brain CTP levels are increased by giving animals uridine, CTP’s circulating precursor in humans (Cansev et al., 2005), PC synthesis is accelerated. The activity of CPT and the extent to which this enzyme is saturated with DAG can also control the overall rate of PC synthesis, for example, in PC12 cells extending neurites after exposure to the nerve growth factor (NGF) (Araki and Wurtman, 1997). If rodents are given a diet that contains both choline and uridine (as its monophosphate, UMP) and, by gavage, a PUFA (particularly the omega‐3 fatty acids DHA or EPA), brain PC synthesis rapidly increases (Cansev et al., 2006; Wurtman et al., 2006), and absolute levels of PC per cell (DNA) or per mg protein increase substantially (e.g., by 40–50% after 4 weeks of daily treatment (Wurtman et al., 2006; > Table 18‐5). These treatments produce parallel or greater increases in each of the other membrane phosphatides, as well as in proteins localized within synaptic membranes, like synapsin‐1, PSD‐95 (> Figure 18‐7), and syntaxin‐3 (Fujita and Kurachi, 2000; Ferreira and Rapoport, 2002;

. Table 18‐5 Effects of UMP‐supplemented diet and/or DHA on brain phospholipid levels in gerbils Treatments Control diet þ Vehicle One week UMP þ DHA Three weeks UMP þ DHA Four Weeks Control diet þ Vehicle UMP diet þ Vehicle Control diet þ DHA UMP diet þ DHA

Total PL 403  23

PC 155  8

PE 69  3

SM 47  3

PS 34  1

PI 20  2

436  15

188  8a

79  6

57  6

47  1c

23  1

502  12c

217  5c

102  4c

73  5b

41  1a

27  1a

351  8 367  22 392  20 442  24c

152  6 171  8a 185  12a 220  12c

65  4 84  8a 78  5a 113  6c

45  2 52  5 56  3a 73  4c

33  3 35  3 39  3 46  6c

21 2 31 2b 32  2b 36  3c

Note: Groups of eight gerbils were given either a control diet and DHA’s vehicle (5% gum Arabic solution, by gavage) or a UMP‐containing (0.5%) diet and DHA (300 mg/kg; in 5% gum Arabic solution, by gavage) for 1 or 3 weeks. In another set of experiments, groups of eight gerbils were given either a control or a UMP‐containing (0.5%) diet, and received orally (by gavage) DHA (300 mg/kg; in 5% gum Arabic solution) or just its vehicle for 4 weeks. At the end of each supplementation period, the gerbils’ brains were harvested and assayed for phospholipids. Data are presented as nmol/mg protein. Data from 1‐and 3‐week‐treated control diet and vehicle groups were pooled as there were no significant differences among these groups a P Figure 18‐4). Uridine‐cytidine kinase (UCK) (ATP:uridine‐50 ‐phosphotransferase, EC 2.7.1.48), the first enzyme in this cascade, catalyzes the phosphorylations of uridine and cytidine to form UMP and cytidine‐50 ‐monophosphate (CMP), respectively (Canellakis, 1957; Skold, 1960; Orengo, 1969). Several different forms of UCK exist, possibly as isoenzymes (Krystal and Webb, 1971; Absil et al., 1980). Humans have two such isoenzymes, UCK1 and UCK2, which have now been cloned (Koizumi et al., 2001; van Rompay et al., 2001). The next enzyme in this sequence, which phosphorylates UMP and CMP to form uridine‐50 ‐diphosphate (UDP) and CDP, respectively, is UMP–CMPK (ATP:CMP phosphotransferase, EC 2.7.4.14) (Hurwitz, 1959; Sugino et al., 1966; Ruffner and Anderson, 1969). UDP and CDP are further phosphorylated to UTP and CTP, by nucleoside diphosphate kinases (NDPK) (Nucleoside triphosphate:Nucleoside diphosphate phosphotransferase, EC 2.7.4.6) (Berg and Joklik, 1954; Parks and Agarwal, 1973).

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The mRNAs for UCK1 (van Rompay et al., 2001) and UMP–CMPK (van Rompay et al., 1999) as well as NDPK activity have been described in brain (Langen et al., 1999; Kim et al., 2002). The interconversions of uridine and cytidine, and of their respective nucleotides, are also observed in mammalian cells. Cytidine and CMP can be deaminated to uridine and UMP (Wang et al., 1950), whereas UTP is aminated to CTP by CTP synthase [UTP:ammonia ligase (ADP‐forming), E.C. 6.3.4.2] (Lieberman, 1956; Hurlbert and Kammen, 1960). This enzyme acts by transferring an amide nitrogen from glutamine to the C‐4 position of UTP, thus forming CTP (Zalkin, 1985). CTP synthase activity has been demonstrated in rat brain (Genchev and Mandel, 1974). All the enzymes described earlier apparently are unsaturated with their respective nucleosides or nucleotides in brain and other tissues. For example, Kms for uridine and cytidine of UCK prepared from various tissues varied between 33 and 270 mM (Skold, 1960; Orengo, 1969; Anderson, 1973; Greenberg et al., 1977), and the Km for uridine of recombinant enzyme cloned from mouse brain was 40 mM (Ropp and Traut, 1996, 1998). Brain uridine and cytidine levels are about 22–46 pmol/mg wet weight (Mascia et al., 1999; Cansev et al., 2005) and 6–43 pmol/mg wet weight (Peters et al., 1987; Cansev et al., 2005), respectively. Hence, the syntheses of UTP and CTP, and the subsequent syntheses of brain PC and PE via the Kennedy pathway, depend on the availability of their pyrimidine substrates. Indeed, an increase in the supply of uridine or cytidine to neuronal cells, in vitro (Savci and Wurtman, 1995; Richardson et al., 2003; Pooler et al., 2005) or in vivo (Cansev et al., 2005; Cansev and Wurtman, 2005), enhanced the phosphorylation of uridine and cytidine, elevating the levels of UTP, CTP, and CDP‐choline.

6

Physiological and Behavioral Effects of Choline

Administration of choline by direct placement into the CNS, orally or by injection, can produce numerous physiological or behavioral effects. Some of these are readily attributable to enhanced ACh release; others may be mediated by phospholipid metabolism or, conceivably, by direct actions of the choline on cholinergic receptors, as discussed earlier. Some remain unexplained.

6.1 Blood Pressure Intravenous choline administration lowers blood pressure in both humans and animals (Mendel et al., 1912; Steigmann et al., 1952; Anton, 1954; Kapp et al., 1970; Singh, 1973; Savci et al., 2003). Intramuscular administration of choline (20 mg/kg/day for 3 days) to rats attenuates the fall in blood pressure induced by acute hemorrhage, and increases survival rate (Altura, 1978). Intraperitoneal choline (60 mg/kg) partially restores blood pressure after the induction of hypotension by acute hemorrhage (Ulus et al., 1995); in contrast intravenous choline (54 mg/kg) further decreases blood pressure and can cause death in hemorrhaged rats (Savci et al., 2003). Oral choline fails to affect cardiovascular function in rats (Ulus et al., 1979) but reportedly lowered blood pressure slightly in some patients with Alzheimer’s disease (Boyd et al., 1977). Intracerebroventricular choline (8 mg/dog) produced a biphasic blood pressure response in anesthetized dogs, an immediate and short‐lasting (about 10–15 min) blood pressure rise (by 60–90 mm Hg) followed by a longer‐lasting (about 60 min) fall (by 15–20 mm Hg; Srimal et al., 1969). In rats, intracisternal choline (12.5–50 mg; Kubo and Misu, 1981a) or its microinjection (1–3 mg/site; Kubo and Misu, 1981b) into the dorsal medulla lowered blood pressure, whereas intracerebroventicular choline (50–150 mg/rat) raised blood pressure and decreased heart rate (Caputi and Brezenoff, 1980; Arslan et al., 1991; Isbil‐Buyukcoskun et al., 2001; Li and Buccafusco, 2004) for 5–20 min. In rats, intracerebreventricular choline (25–150 mg/rat) restored normal blood pressure among animals made hypotensive by acute hemorrhage (Ulus et al., 1995; Savci et al., 2002b), endotoxin (Savci and Ulus, 1997), chemical sympathectomy (Gurun et al., 1997a), spinal cord transection (Savci and Ulus, 1998), autonomic ganglion blockade, or a‐adrenoceptor blockade (Savci and Ulus, 1996). At a dose of 180 nmol, choline also potentiated the pressor responses evoked by naloxone or glycyl‐glutamine (Gurun et al., 2003). In normotensive rats, the blood pressure responses to choline administered centrally involve local activation of both mAChRs and nAChRs (Arslan et al., 1991);

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including the a7 nAChR subtype (Li and Buccafusco, 2004). In hypotensive animals, these blood pressure responses involve presynaptic activation of central nAChRs (Ulus et al., 1995; Savci and Ulus, 1996, 1997, 1998; Gurun et al., 1997a; Savci et al., 2002b).

6.2 Body Temperature Intracerebroventricular (75–300 mg/rat), but not intraperitoneal (30–120 mg/kg), choline decreases body temperature (Unal et al., 1998). This hypothermia is mainly mediated by M1and M3 muscarinic receptors (Unal et al., 1998).

6.3 Pain Choline can alter responses to painful stimuli in experimental animals, and can modify the actions of analgesic drugs. Given intraperitoneally (15–60 mg/kg) to rats, it diminishes the analgesic actions of morphine (10 mg/kg; subcutaneously) as assessed using the hot‐plate test (Botticelli et al., 1977). In mice, subcutaneous (up to 500 mg/kg; Damaj et al., 2000) or intravenous (2–64 mg/kg; Wang et al., 2005b) choline fails to alter responses to thermal pain, but when given intracerebroventricularly (Damaj et al., 2000; Wang et al., 2005b) or intrathecally (Damaj et al., 2000) at doses of 30–120 mg/animal, it produces antinociception in the same pain model. This latter effect can be blocked by the nonselective mAChRs antagonist atropine or by antagonists of a7 nAChRs (e.g., metillycaconitine; a‐bungarotoxin). Intravenous choline (4–64 mg/kg) produced significant antinociception in mice in the late phase of inflammatory pain responses to subcutaneous injection of 5% formalin (Wang et al., 2005b). At a 2 mg/kg dose, it also enhanced the antinociceptive effects of aspirin (9.4 mg/kg; i.v.) and morphine (0.165 mg/kg; i.v.). This action was also blocked by the a7 nAChR antagonists metillycaconitine and a‐bungarotoxin, but not by atropine or naloxone (Wang et al., 2005b).

6.4 Neuroendocrine Effects Choline produces a variety of neuroendocrine responses when administered peripherally or centrally to rats. Its oral administration (20 mmol/kg by stomach tube) for 4 days increased urinary catecholamine output (Scally et al., 1978). Given intraperitoneally, choline (30–120 mg/kg) elevated plasma catecholamines (Ilcol et al., 2002c) and insulin (Ilcol et al., 2003a). Given intracerebroventricularly (50–150 mg/rat), it increased plasma concentrations of the catecholamines (Arslan et al., 1991; Ulus et al., 1995; Gurun et al., 2002); vasopressin (Arslan et al., 1991; Ulus et al., 1995; Savci and Ulus, 1996, 1998; Gurun et al., 1997a; Savci et al., 2003); ACTH (Savci et al., 1996); b endorphin (Savci et al., 1996); and prolactin (Gurun et al., 1997b). The increases in plasma vasopressin, ACTH, and b‐endorphin were found to be mediated by central nAChRs, and that of prolactin by mAChRs (Gurun et al., 1997b). The increases in plasma insulin after intraperitoneal choline involve both ganglionic nAChRs and the M1 and M3 subtypes of mAChRs (Ilcol et al., 2003a). Choline’s site of action in producing rest of these neuroendocrine effects is principally presynaptic, i.e., via enhancing ACh release (Scally et al., 1978; Savci et al., 1996, 1998, 2003; Ilcol et al., 2003a), and the effects are enhanced by hemorrhagic, hypotensive, or osmotic stresses (Scally et al., 1978; Ulus et al., 1995; Savci et al., 1996, 1998, 2003; Ilcol et al., 2003a).

6.5 Peripheral Metabolism Peripheral (40–120 mg/kg; intraperitoneal) or central (75–300 mg/rat; icv) administration of choline increases blood glucose levels in rats (Gurun et al., 2002; Ilcol et al., 2002c). This hyperglycemic response is prevented by blockade of ganglionic nAChRs or a‐adrenoceptors, as well as by bilateral adrenalectomy

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(Ilcol et al., 2002c), and apparently is mediated by the stimulation of adrenomedullary catecholamine release and subsequent activation of a‐adrenoceptors. It apparently also involves activation of central nAChRs affecting the sympathoadrenal system (Gurun et al., 2002). As decribed earlier, glucose is a source of acetyl‐CoA, which, like choline, is a limiting precursor for ACh. Hence, choline administration may enhance ACh synthesis by this additional mechanism.

6.6 Behavior Acute administration of choline to humans, as choline chloride or as PC, improved short‐term memory in some studies (Leathwood et al., 1982; Ladd et al., 1993) but not in others (Davis et al., 1980; Harris et al., 1983). Intraperitoneal choline (6–60 mg/kg) in combination with glucose (10–30 mg/kg) improved passive avoidance behavior in mice (Kopf et al., 2001). Chronic consumption of a choline‐rich diet by aged mice reportedly counteracted the age‐associated decline in learning and memory (Bartus et al., 1980; Golczewski et al., 1982; Leathwood et al., 1982). Supplementation of pregnant or lactating rats with choline chloride during the perinatal period (embryonic days 12–17 and/or postnatally 16–30 days) caused long‐lasting improvements in spatial memory, as illustrated using radial‐arm maze tests (Meck et al., 1988, 1989; Meck and Williams, 1997b, 1999; Williams et al., 1998) and the Morris water maze test (Schenk and Brandner, 1995; Tees, 1999a, b; Tees and Mohammadi, 1999). In adulthood, rats that had received supplemental choline (about four times dietary levels) perinatally exhibited increased memory capacity (Meck and Williams, 1997c) and precision (Meck and Williams, 1999), and performed more accurately on tests of spatial memory (Meck et al., 1988, 1989; Meck and Williams, 1997a, b, c, 1999; Williams et al., 1998). Moreover, perinatal supplementation with choline provided some protection against memory impairments usually associated with normal aging (Tees and Mohammadi, 1999), neonatal alcohol exposure (Thomas et al., 2004; Wagner and Hunt, 2006), or epileptic seizures (Yang et al., 2000; Holmes et al., 2002). Although the mechanism by which perinatal choline supplementation produces these long‐term effects on memory remains unclear, such supplementation is known also to cause enduring neuroanatomic and neurochemical changes in brain regions involved in memory. For example, choline supplementation increased hippocampal ChAT activity, mAChRs density (Meck et al., 1989), ACh release (Cermak et al., 1998), and the amplitude of ACh‐mediated excitatory potentials (Montoya et al., 2000), and it reduced acetylcholinesterase activity (Cermak et al., 1998, 1999). Moreover, basal forebrain cholinergic neurons projecting to the hippocampus were larger and more spherical among rats that had been supplemented perinatally with choline (Loy et al., 1991; Williams et al., 1998). Prenatal choline supplementation also reportedly enhanced MAPK and CREB activation (Mellott et al., 2004); N‐methyl‐D‐aspartate (NMDA) receptor‐mediated neurotransmission (Montoya and Swartzwelder, 2000); long‐term potentiation (LTP) (Pyapali et al., 1998); PLD activity (Holler et al., 1996); NGF levels (Sandstrom et al., 2002); and dendritic spine formation (Mervis, 1982) in rat hippocampus. It also decreased the rate of apoptosis in the hippocampus and basal forebrain of 18‐day old fetuses (Holmes‐McNary et al., 1997) and increased brain cell division (Albright et al., 1999a, b). Prenatal supplementation with choline also was associated with greater excitatory responsiveness, reduced slow afterhyperpolarizations, enhanced afterdepolarizing potentials, larger somata, and greater basal dendritic arborization in hippocampal CA1 pyramidal cells studied postnatally at 20–25 days of age (Li et al., 2003).

6.7 Drug Interactions Acute or chronic choline administration can modify the actions of some centrally active drugs. Chronic (28–35 days) treatment of rats with choline through the diet (about ten times more choline than in control diets) produced behavioral hyperactivity and attenuated the sedative/hypnotic and hypotermic effects of pentobarbital (Wecker et al., 1987). Chronic choline supplementation also increased the density of binding sites for nicotine (Coutcher et al., 1992) and a‐bungarotoxin (Morley and Garner, 1986) and produced

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tolerance to the convulsive and lethal actions of nicotine (Wecker et al., 1982). Chronic dietary choline supplementation to mice modulated benzodiazepine receptor binding and g‐aminobutyric acid receptor function (Miller et al., 1989), also decreasing seizure activity and the lethality of such seizure‐promoting drugs such as nicotine, paraoxon, strychnine, and pentylenetetrazol (Wecker et al., 1982). Acute intraperitoneal choline administration (100 mg/kg) to morphine‐dependent rats decreased withdrawal symptoms and the associated weight loss (Pinsky et al., 1973; Frederickson and Pinsky, 1975).

6.8 Neuroprotective and Cytoprotective Effects Choline exhibits cytoprotective and neuroprotective actions in vivo and in vitro: In vitro, choline itself or some of its analogs (i.e., mono‐, di‐, and triethylcholine and pyrrolidinecholine), can, at 1–10 mM concentrations, protect against manifestations of cytotoxicity in differentiated PC12 cells induced by growth factor deprivation (Jonnala et al., 2003). Choline (5–75 mM) suppresses the development of dark cell degeneration in Purkinje neurons following receptor activation with AMPA (DL‐amino‐3‐hydroxy‐5‐ methyl‐isoxazole‐4‐propionic acid; Strahlendorf et al., 2001). At concentrations of 1 mM or greater it also reduces NMDA toxicity in organotypic hippocampal slice cultures derived from neonatal rats (Mulholland et al., 2004), and reportedly it is cytoprotective in PC‐12 cells (Jonnala et al., 2003); Purkinje neurons (Strahlendorf et al., 2001); and hippocampal cells (Mulholland et al., 2004)—an effect mediated by the a7 subtype of nAChRs. Prenatal choline supplementation for 6 days during days E12–E17 of gestation protected against subsequent neurodegeneration in the posterior cingulate and retrosplenial cortices, induced in female adolescent rats by peripheral administration of dizocilpine (Guo‐Ross et al., 2002). In cultured neonatal cardiac ventricular cells, choline (0.1–1 mM) reduced H2O2‐induced apoptotic cell death, by acting via M3 mAChRs (Yang et al., 2005). In vivo, choline (20 mg/kg; i.v.) attenuated endotoxin‐ induced multiple organ injury (i.e., renal, hepatic, and cardiac injuries) in dogs (Ilcol et al., 2005b). Similarly, in rats, choline supplementation in the diet for 3 days attenuated endotoxin‐induced hepatic injury and improved survival (Rivera et al., 1998). Choline‐induced protection from the tissue injuries induced by endotoxin is associated with a reduction in serum levels of tumor necrosis factor‐a (TNF‐a) (Rivera et al., 1998; Ilcol et al., 2005b) and with the improvement in platelet counts and platelet closure times (Yilmaz et al., 2006a). Intravenous choline administration (5 mg/kg) protected rats (5 mg/kg; i.v.) from ischemic myocardial injuries by stimulating M3‐mAChRs (Yang et al., 2005).

7

Effects of Exogenous CDP‐Choline

Cytidine‐50 ‐diphosphocholine (CDP‐choline; citicoline), which is composed of choline and cytidine linked by a diphosphate bridge, is both an essential intermediate in the synthesis of endogenous PC through the Kennedy cycle (> Figure 18‐4), and a drug used in some countries to treat cerebral ischemia, traumatic brain injury, Parkinson’s disease, or stroke. When administered orally or parenterally, exogenous CDP‐choline is completely hydrolyzed, first to cytidine monophosphate and phosphocholine, and then to free cytidine and choline (Lopez G‐Coviella et al., 1987). In humans, the cytidine is further transformed to uridine, hence giving CDP‐choline to humans causes dose‐related increases in serum uridine and choline levels but not in serum cytidine (Wurtman et al., 2000; Cansev, 2006). In laboratory rodents, depending on species and on the activity of the hepatic enzyme cytidine deaminase (Chabot et al., 1983; Ku¨hn et al., 1993), which converts cytidine to uridine, CDP‐choline administration can principally elevate either serum uridine (e.g., in gerbils) or cytidine (e.g., in rats), besides choline. Moreover, as treatments that elevate plasma uridine (or cytidine) and choline thereby increase brain PC synthesis (Wurtman et al., 2000; Richardson et al., 2003; Cansev et al., 2005; Ulus et al., 2006) as well as steady‐state brain levels of PC and other membrane phosphatides (Lopez G‐Coviella et al., 1992, 1995; Wurtman et al., 2006), it is likely that some of CDP‐choline’s therapeutic actions result from changes that it produces in the quantities or the composition of brain membranes. Some other effects of exogenous CDP‐choline (Savci et al., 2002a, 2003; Cavun and Savci, 2004;

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Cavun et al., 2004) probably are mediated by increasing ACh release, secondary to the rise it produces in plasma and brain choline levels (Savci et al., 2002a, 2003). By increasing brain levels of endogenous CDP‐choline, exogenous CDP‐choline also increases the amounts of DAG that combine with this intermediate to form PC (and that combine with endogenous CDP‐ethanolamine to form PE) (Araki and Wurtman, 1998). CDP‐choline administration can also affect the brain by increasing the amount of free AA that is used to form DAG, which then combines with endogenous brain CDP‐choline. This causes a decrease in free AA levels, which might otherwise be neurotoxic, and thus it decreases the ultimate size of the brain damage following a stroke and brain injury (Warach et al., 2000). This reduction in AA may be the major mechanism underlying CDP‐choline’s acute therapeutic effects (Lopez G‐Coviella et al., 1998). It has also been suggested that exogenous CDP‐choline may decrease brain levels of free AA by directly inhibiting phospholipase A2 activity or by decreasing the formation of that enzyme protein (Adibhatla and Hatcher, 2003, 2005; Adibhatla et al., 2006).

7.1 Hypoxia and Ischemia Intracerebral CDP‐choline (0.6 mmoles) prevented the ischemia‐induced loss of radioactive choline from glycerophospholipids, and suppressed the increase in brain levels of free fatty acids in a global model of ischemia in rats (Dorman et al., 1983; Goldberg et al., 1985). In a rat model of transient cerebral ischemia, intraperitoneal CDP‐choline (250 mg/kg; twice a day for 4 days) improved neurological signs and attenuated the increases in glucose and pyruvate levels and the decrease in ACh synthesis from labeled glucose (Kakihana et al., 1988). In an ischemic and anoxic rat model, CDP‐choline (300 mg/kg, i.p.) decreased the incidence of neurological deficits (Yamamoto et al., 1990). In a chronic hypoxia rat model produced by placing animals in chambers in which the oxygen content was depressed (7–15%) for extended time periods, CDP‐choline (100 mg/kg in food) protected vigilance behavior (Hamdorf and Cervos‐ Navarro, 1990), reduced hypoxia‐induced behavioral deterioration (Hamdorf and Cervos‐Navarro, 1991), and increased survival time at 7% O2 (Hamdorf and Cervos‐Navarro, 1991; Hamdorf et al., 1992). In a model of rat experimental hypoxia induced by giving potassium cyanide, oral CDP‐choline given for 4 days before the induction of hypoxia increased survival time (Tornos et al., 1983a). Araki et al. (1988) also observed a neuroprotective effect of CDP‐choline in mice in which the cerebral ischemia was induced by decapitation or by potassium cyanide intoxication. CDP‐choline (500 mg/kg; i.p.) for 14 days delayed cell membrane damage and behavioral dysfunction in spontaneously hypertensive rats in which ischemia had been caused by artificially induced occlusion of the lateral middle cerebral artery (Aronowski et al., 1996). In a similar study, CDP‐choline (500 mg/kg; i.p.) decreased infarct volume and edema in a rat model of temporary focal ischemia (Schabitz et al., 1996). In mice with an intracerebral hemorrhage, CDP‐choline (500 mg/kg; i.p.) reduced the volume of ischemic injury surrounding the hematoma, and improved the behavioral outcome (Clark et al., 1998). In another study, CDP‐choline (400 mg/kg; i.p.) increased blood pressure, reduced infarct volume, and decreased the mortality rates of hypotensive rats with a experimental subarachnoid hemorrhage (Alkan et al., 2001). In rats with permanent occlusion of the middle cerebral artery, CDP‐choline inhibited MAP kinase signaling pathways (Krupinski et al., 2005). In a focal brain ischemia model in rats, CDP‐choline (0.5–2 g/kg; i.p.) reduced infarct size and inhibited ischemia‐induced decreases in cortical and striatal ATP levels (Hurtado et al., 2005). CDP‐choline produces synergistic neuroprotective effects when this treatment is combined with glutamate receptor antagonists (i.e., MK‐801; Onal et al., 1997 or lamotrigine; Ataus et al., 2004); thrombolytic agents (i.e., recombinant tPA; Andersen et al., 1999; De Lecinana et al., 2006 or urokinase; Shuaib et al., 2000); the calcium channel blocker, nimodipine (Sobrado et al., 2003); or basic fibroblast growth factor (Schabitz et al., 1999) using experimental ischemia models in rats. In a gerbil model in which brain ischemia was produced by bilateral ligation of the carotid arteries, intraventricular (0.6 mmol; Trovarelli et al., 1981) or intraperitoneal (150 mg/kg; Trovarelli et al., 1982) CDP‐choline partially prevented the ischemia‐induced increases in fatty acids and decreases in PC levels (Trovarelli et al., 1981, 1982). CDP‐choline reduced the dysfunctions of the BBB after reperfusion in gerbils (Rao et al., 1999), and reduced the cerebral edema, concurrently reducing the levations of AA levels and

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leukotriene C4 synthesis (Rao et al., 2000). In a transient cerebral ischemia model, CDP‐choline (500 mg/kg daily for 2 days; i.p.) restored the decreases in PC, SM, cardiolipin, and total glutathione levels induced by ischemia (Adibhatla et al., 2001). In cats undergoing brief periods of cerebral ischemia, CDP‐choline attenuated the depression in the cortical evoked potentials (Boismare et al., 1978).

7.2 Head Trauma (Cranio‐Cervical Trauma) Neuroprotective effects of CDP‐choline have been described in studies using various trauma models and experimental animals. In a weight‐drop concussive head injury model in mice, CDP‐choline (60–250 mg/kg) shortened the recovery time (Boismare et al., 1977). In a controlled lateral‐impact model in rat, CDP‐choline (100 mg/kg; i.p.) increased extracellular ACh levels, decreased cognitive deficits and attenuated the trauma‐induced increased sensitivity to the memory‐disrupting effects of scopolamine (Dixon et al., 1997). In a cortical impact model in rat, intraperitoneal CDP‐choline (400 mg/kg) reduced brain edema (Baskaya et al., 2000), and decreased neuronal loss in the hippocampus, and improved neurological recovery (Dempsey and Raghavendra Rao, 2003). In a rat experimental (weight‐drop) spinal cord injury model, CDP‐choline (400 mg/kg; i.p.) improved behavioral and neuroanatomic signs of recovery (Yucel et al., 2006).

7.3 Induced Lesions Neuroprotective actions of CDP‐choline have also been demonstrated in lesion studies. Oral administration of CDP‐choline, at a daily dose of 1 g/kg for 4 days, significantly extended survival time and increased the percentage of survivors from KCN‐induced toxicity (Tornos et al., 1983a). CDP‐choline administration (500 mg/kg, i.p.) for 7 days ameliorated functional behavior, as shown by reducing the number of apomorphine‐induced contralateral rotations. It also attenuated the loss of substantia nigra dopaminergic neurons and the decrease in tyrosine hydroxylase immunoreactivity, in the ipsilateral striatum in rats injected intrastriatally with the dopaminergic toxin, 6‐hydroxydopamine (Barrachina et al., 2003). CDP‐ choline (62.5–250 mg/kg, i.p.) protected hippocampal neurons against apoptosis and the degeneration induced by injecting beta‐amyloid into brains of rats also undergoing cerebral hypoperfusion (Alvarez et al., 1999). CDP‐choline (50 mg/kg) prevented mice and rats from an acrylamide‐induced neurological syndrome (Agut et al., 1983). In tissue culture studies, CDP‐choline protected the retinal ganglion cells (Oshitari et al., 2002) and prevented glutamate‐mediated cell death in cerebellar granule neurons (Mir et al., 2003).

7.4 Other Effects Oral or intraperitoneal administration of CDP‐choline (10–500 mg/kg, for 5–7 days) improved memory in rats with memory deficits induced by muscarinic AChR antagonists, by the a2‐adrenoceptor agonist clonidine, by electroconvulsive shock, or by hypoxia (Petkov et al., 1992, 1993). Dietary CDP‐choline supplementation protected rats against the development of memory deficits in aging (Teather and Wurtman, 2003), and prevented memory impairments caused by impoverished environmental conditions (Teather and Wurtman, 2005). In humans, CDP‐choline improved verbal memory in aging (Spiers et al., 1996) and benefitted memory in elderly subjects (Alvarez et al., 1997). In rat striatum, CDP‐choline activated tyrosine hydroxylase (Martinet et al., 1981), increased dopamine levels (Martinet et al., 1979; Shibuya et al., 1981), and enhanced Kþ‐evoked dopamine release (Agut et al., 2000) and haloperidol‐induced elevation in dopamine metabolites (Agut et al., 1984). Oral CDP‐choline increased the total urinary excretion of the noradrenaline metabolite 3‐methoxy‐4‐hydroxyphenylglycol, in rats and humans (Lopez G‐Coviella et al., 1986). Centrally-administered CDP‐choline (0.5–2.0 mmol)

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increased plasma vasopressin (Cavun et al., 2004), and ACTH concentrations and potentiated the release of GH, TSH, and LH (Cavun and Savci, 2004) stimulated by clonidine, TRH, and LHRH, respectively. Intravenously injected CDP‐choline (250 mg/kg) increased plasma concentrations of noradrenaline and adrenaline in rats (Savci et al., 2003). Oral administration of a single dose (2 g/kg) of CDP‐choline to mice decreased the intensity of the morphine withdrawal syndrome (Tornos et al., 1983b). In a single study, treatment with CDP‐choline was reported to affect some measures of craving in cocaine‐dependent human subjects (Renshaw et al., 1999). CDP‐Choline increased blood pressure, reversed hypotension in hemorrhagic shock (Savci et al., 2002a, 2003), and prolonged survival time (Yilmaz et al., 2006b) when given intravenously (100–500 mg/kg) or intracerebroventricularly (0.1–2.0 mmol) to rats. CDP‐choline decreased platelet reactivity to aggregating agents when given acutely (250 mg/kg) and increased the antiaggregatory activity of aortic walls when given chronically (250 mg/kg, i.p., 2 weeks) to rats. In dogs, CDP‐choline (70 mg/kg; i.v.) prevented the endotoxin‐induced decrease in circulating platelet counts and prolonged platelet closure times (Yilmaz et al., 2006a). Intracerebroventicular CDP‐choline (0.5–2.0 mmol) produced antinociception in three different acute pain models (i.e., thermal paw withdrawal tests, mechanical paw pressure test, and acetic acid writhing test) in rats (Hamurtekin and Gurun, 2006).

7.5 Clinical Studies CDP‐choline effects have been examined in studies involving numerous normal subjects and patients with cerebral ischemia, traumatic brain injury, hypoxia, Alzheimer’s and Parkinson’s diseases. To date sufficient evidence has not been accumulated regarding any such use to warrant its approval for drug status by the US Food and Drug Administration. It is, however, approved for sale in a few other countries and sold under its international nonproprietary name, citicoline. Clinical trials conducted in the USA, tested daily oral doses of 500, 1000, or 2000 mg/day, given for 6 weeks. In some such studies, the drug was administered within the first 48 h of an ischemic stroke (Clark et al., 1999); in others, it was first given to patients up to 14 days after the onset of the ischemic episode (Tazaki et al., 1988). Pooling of individual patients data from four USA trials yielded evidence that CDP‐ choline treatment could improve overall recovery at 12 weeks in acute ischemic stroke patients (Davalos et al., 2002). Pooled diffusion‐weighted magnetic resonance imaging data from two clinical trials showed a significant dose‐dependent reduction on percent change in lesion volume (Warach, 2002).

8

Choline in Autonomic and Motor Neurons

All nerves that leave the brain or spinal cord (i.e., axons of motor neurons, parasympathetic preganglionic neurons, and sympathetic preganglionic neurons), as well as all postganglionic parasympathetic neurons, release ACh as their neurotransmitter, and in all of them choline availability determines the rates at which ACh is synthesized and released. ACh is also present in the periphery in placenta, lymphocytes, the bladder, and tracheal epithelium; however, in these cells the effects of increasing choline availability on ACh synthesis have not yet been determined. More than 50 years ago, Hutter (1952) demonstrated that a low intravenous dose (7 mg/kg) of choline enhanced neuromuscular transmission, whereas a high dose (50 mg/kg) blocked this transmission. Hutter also demonstrated that choline, at doses of 3–60 mg/kg, could restore neuromuscular transmission in curarized cats. Based on these observations, he suggested that choline increased ACh output from motor nerve endings. More recently, using the isolated, vascularly perfused rat phrenic nerve‐hemidiaphragm preparation Bierkamper and Goldberg (1979, 1980) directly demonstrated that choline (at 30–60 mM concentrations) could increase ACh release at the neuromuscular junction. Effects of choline on ACh synthesis and release, and on cholinergic neurotransmission, at parasympathetic synapses have been demonstrated in vivo (Kuntscherova, 1972; Haubrich et al., 1974, 1975;

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Ilcol et al., 2003a) and in vitro, using isolated hearts (Dieterich et al., 1978), and atrial (Meyer and Baker, 1986) and pancreatic minces (Ilcol et al., 2003a). Choline infusion (10 mM) increases by 2–3‐fold ACh release evoked by electrical stimulation of the vagus nerve in chicken hearts, and by at least 23‐fold in cat heart (Dieterich et al., 1978). The presence of choline (10 mM) in the perfusion medium also increased, by—two to threefold, ACh release evoked by electrical field stimulation (at 20 Hz for 20 min) from isolated chicken, rat, cat, and guinea pig hearts (Dieterich et al., 1978). Subcutaneous choline administration (200 mg/kg) increased, by 34%, the ACh content of the atrium (Kuntscherova, 1972) and in atrial minces, choline (at 1–100 mM) increased ACh synthesis and release in a concentration‐dependent manner (Meyer and Baker, 1986). In rats, intraperitoneal choline (90 mg/kg) increased, by 45%, the ACh contents of pancreatic tissue; this was associated with increased cholinergic neurotransmission to insulin secreting b‐cells (Ilcol et al., 2003a). Choline (10–130 mM) also increased ACh synthesis and release from rat pancreatic minces (Ilcol et al., 2003a). Oral choline administration (20 mmol/kg) to rats increases adrenal ACh levels by more than twofold for 8 h (Ulus et al., 1977a) and tyrosine hydroxylase activity by about 30% (Ulus et al., 1977a). Repeated oral administration of choline for 4 days increases the enzyme activity further, by up to 50–60%; the increase in tyrosine hydroxylase activity is not observed after intubation with saline, water, or ammonium chloride, and fails to occur in rats pretreated with cycloheximide (Ulus et al., 1977a). Similar increases in tyrosine hydroxylase activity after oral choline administration are observed in sympathetic ganglia, including the superior cervical ganglion, the stellate and celiac ganglia (Ulus et al., 1977c, 1979; Ulus and Wurtman, 1979), and the ganglia of the thoracic sympathetic chain (Ulus et al., 1977c). These increases in tyrosine hydroxylase activity are also not seen in adrenals after adrenal denervation (Ulus et al., 1977a) or after decentralization of superior cervical ganglion (Ulus et al., 1979), indicating that the action of choline is transsynaptic and that it requires intact preganglionic cholinergic nerves to affect ACh synthesis and release. Further evidence that choline administration enhances ACh release was obtained by studies in which choline was administered along with reserpine, or with other drugs that increase impulse flow in preganglionic cholinergic nerves. Injection of reserpine (2.5 mg/kg; i.p.) phenoxybenzamine (20 mg/kg; i.p.), or insulin (2 units/rat; i.p.) daily for 4 days, or of 6‐hydroxydopamine (200 mg/kg twice, with an interval of 48 h, through tail vein), all caused marked increases in adrenal tyrosine hydroxylase activities. When these treatments were combined with oral choline (2.8 g/kg; by stomach tube), the resulting increases in tyrosine hydroxylase activity were considerably greater than the sum of the changes produced by choline alone and drug alone, that is, significant potentiation occurred (Ulus et al., 1977a, b, 1978). Potentiation of a treatment‐induced rise in tyrosine hydroxylase by choline was also observed in the adrenals of rats kept in the cold (Ulus et al., 1977a, 1978) and in superior cervical ganglia of reserpine‐treated rats (Ulus et al., 1977a). Taken together, these data indicate that the availability of free choline is a major factor controlling cholinergic neurotransmission in the sympathoadrenal system. Studies using the classical perfused‐superior cervical ganglion system failed to demonstrate parallel increases in ACh release in response to elevating the choline concentration of the perfusion media (Birks and MacIntosh, 1961; Matthews, 1966; MacIntosh, 1979; O’Regan and Collier, 1981). However, when superior cervical ganglia were perfused with choline‐containing (10–14 mM) plasma or with Locke solution, they released greater amounts of ACh, by twofold, during a 1‐h stimulation (20 Hz) period than ganglia superfused without exogenous choline (Birks and MacIntosh, 1961; Matthews, 1966; MacIntosh, 1979). Furthermore, the ACh stores in perfused‐ganglia stimulated without exogenous choline were found to be partly depleted, although these ganglia had managed to synthesize some ACh by reusing free choline generated from the hydrolysis of released ACh, or from hydrolyzing membrane phospholipids like PC). PC levels and the number of synaptic vesicles in the cat’s superior cervical ganglion were found to decline significantly after stimulation of the preganglionic nerve trunk if the uptake of exogenous choline was blocked by HC‐3 (Parducz et al., 1976), or if the ganglia were perfused with a choline‐free Locke solution (Parducz et al., 1986). In striking contrast, stimulated ganglia supplied with exogenous choline maintained their ACh stores (Birks and MacIntosh, 1961; Matthews, 1966; MacIntosh, 1979), as well as membrane PC levels and the numbers of storage vesicles (Parducz et al., 1976, 1986), although they released much more ACh than they initially contained.

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Acknowledgments The authors thank Dr. Jan Krzysztof Blustajn and Ms. Carol Watkins for the critical review of this chapter. Studies described in this chapter were supported in part by grants from the National Institutes of Mental Health (MH‐28783); the NIH‐NCRR (5‐MO1RR01066–29); the Center for Brain Sciences and Metabolism Charitable Trust; and the Turkish Academy of Sciences (Ismail H. Ulus).

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Alcohol and Neural Lipids

E. Cazzaniga . A. Bulbarelli . M. Masserini

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 504

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Effects of Administration of Ethanol on Brain Morphology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 504

3 3.1 3.2 3.3 3.3.1 3.3.2 3.3.3 3.3.4 3.4 3.4.1 3.4.2 3.5 3.6 3.7 3.8

Effects of Ethanol on Neural Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 505 Membrane Fluidity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 505 Lipid Composition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 505 Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 505 Cholesterol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 505 Phospholipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 506 Plasmalogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 506 Glycosphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 506 Lipid Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 507 Studies in vitro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 507 Studies in vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 507 Lipid Peroxidation and Antioxidants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 507 Fetal Alcohol Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 508 A Unique Phospholipid: Phosphatidylethanol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 509 Ethanol and Dementia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 509

4

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 509

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_19, # Springer ScienceþBusiness Media, LLC 2009

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Abstract: Ethanol is a psychotropic drug, whose acute or chronic abuse may lead to physical dependency, cognitive dysfunctions, and brain damages. Many of the ethanol effects arise as a result of the alteration of membrane functions, which follows changes either in physicochemical properties or in the lipid composition or both of neural cells. In fact, in vitro studies have shown that ethanol affects membrane fluidity either directly, through incorporation in the bilayer, or indirectly, inducing changes in the overall unsaturated:saturated fatty acid ratio of membrane lipids. Indeed, ethanol induces more considerable changes of neural cell lipid composition; hippocampus, which plays a critical role in learning and memory, is especially sensitive. In fact, chronic exposure to ethanol in vivo and in vitro affects phospholipid and ganglioside pattern and, noteworthy, induces the formation of a unique phospholipid, phosphatidylethanol. Considering the role of phospholipids and gangliosides in neuronal development and in various signal transduction pathways, these changes may have significant implication in cognitive dysfunctions observed. Studies in vitro indicate that alcohol enhances ganglioside biosynthesis by increasing the sphingosine recycling, an hypothetic mechanism of cell defence during alcoholic stress. Chronic ethanol consumption also stimulates changes in lipid organization associated with lipid peroxidation, a critical event in many neurological diseases and disorders, in particular the fetal alcohol syndrome, caused by ethanol exposure during prenatal and developmental period. Several diet components could have a preventing effect against ethanol‐induced oxidative stress. The present review focuses on all these subjects. List of Abbreviations: Ab, b‐amyloid peptide; DAG, diacylglycerol; DHA, docosahexaenoic acid; FAS, fetal alcohol syndrome; GlcCer, glucosyl‐ceramide; PC, phosphatidyl‐choline; PE, phosphatidyl‐ethanolamine; PEth, Phosphatidylethanol; PI, phosphatidyl‐inositol; PS, phosphatidyl‐serine; PUFA, polyunsaturated fatty acids; TG, triglyceride

1

Introduction

Ethanol is a psychotropic drug that has been used since ancient times in a social context because of its euphorizing and sedative effects at low doses. However, at high doses ethanol is a central nervous system depressant and toxic doses may result in coma and death (Bass and Volpe, 1989; Leskawa et al., 1995). Moreover, chronic assumption of ethanol may lead to physical dependency and cause tissue damage in several organs, including the brain. There is evidence that many of the effects exerted by ethanol on brain and associated with acute or chronic abuse, arise as a result of the alteration of membrane functions, which follows the change either in physicochemical properties or in the lipid composition or both of neural membranes (Leonard, 1986). The present review focuses on this subject.

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Effects of Administration of Ethanol on Brain Morphology

Already in the Courville study (1955), it was reported a particularly extensive atrophy of the frontal lobes of postmortem brains of chronic alcoholics. Other studies (Horvath, 1975; Carlen et al., 1978; Epstein et al., 1978) using air encephalography or computer‐assisted tomographic (CAT) scanning devices, showed distinct dilation of the lateral ventricles and cortical sulci in alcoholic patients, a feature associated with impaired intellectual functioning. Quantitative morphometric studies showed that alcoholic brain shrinkage is related to a reduction in the volume of the white matter in the cerebral hemispheres, rather than to volume changes in the cerebral cortex. However, the cortex is also abnormal, due to a loss of neurons from the frontal region. In this and in other regions of the cortex there is shrinkage of the neuronal soma. This is reflected in a retraction of the neuronal dendritic arbour, which plays a crucial role in cell–cell communication (Harper and Kril, 1990).

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Effects of Ethanol on Neural Lipids

3.1 Membrane Fluidity Fluidity is a membrane feature typically dependent on lipids. Complex membrane lipids may contribute to this feature, forming more or less ordered environments by interaction either among their polar headgroups or their lipid moieties, in dependence of their chemical composition (e.g., fatty acid length and saturation). The information that addition of ethanol induces a short‐term increase on membrane fluidity was first reported by Chin and Goldstein (1977a, 1977b), using a paramagnetic resonance technique and suspensions of brain membranes (i.e., synaptosomal, myelin, microsomal), and since then repeatedly confirmed (Chu‐Ky et al., 2005; Sergent et al., 2005; Dolganiuc et al., 2006). As described by Holte and Gawrisch (1997), who investigated by NMR technique the localization of an ethanol molecule within a membrane in an artificial system (multilamellar liposomes), the fluidizing effect of ethanol is exerted via a disordering effect on the membrane. In fact, diffusion of alcohol occurs easily and freely within all the environments the lipid bilayer has to offer, encompassing both hydrophobic and hydrophilic moieties of membrane lipids. Ethanol exerts also long‐term effects on membrane fluidity, by affecting in living cells the fatty acid composition of membrane lipids. However, data obtained in vitro or in vivo are apparently controversial. Experiments on Escherichia Coli, grown in the presence of ethanol, showed a reduced level of saturated fatty acids in phospholipids, primarily due to a decrease of the amount available for their synthesis (Buttke and Ingram, 1978). According to the experiments carried out in E. Coli, Morrisson et al. (1984) reported that the relative degree of saturation of membrane phospholipids decreases in cultured neurons exposed to ethanol. Later, Duffy et al. (1991) showed that fatty acid polyunsaturation increases in the brain of newborn rats exposed to a chronic ethanol treatment in utero. However, contrarily to the above results, Littleton (1979) and Littleton et al. (1980) reported a decrease in the unsaturation of synaptosomal membrane lipids in mice treated with ethanol. Aloia et al. (1985) also reported a reduction in the level of polyunsaturated fatty acids (PUFA), particularly evident in phosphatidyl‐inositol (PI) and phosphatidyl‐serine (PS). The linoleic:arachidonic acid (18:2/20:4) ratio and the saturation:unsaturation ratio were also increased in the same phospholipids extracted from membranes of alcoholic animals. The authors speculated that these changes, decreasing the overall unsaturation degree of fatty acids and thus the fluidity, have the scope of contrasting the fluidizing effect of ethanol, and underlies the rapid onset of tolerance to ethanol in mice.

3.2 Lipid Composition Cell cultures. It has been demonstrated that the effects of alcohol on cell lipid composition may vary in dependence of its concentration. Membrane lipids of rat cerebellar granule cells in culture are differently affected by increasing ethanol concentrations: while 40 mM ethanol has no apparent effect, 55 mM concentration induces a significant increase of total cholesterol, 80 mM modifies the phospholipid fatty acid composition (Omodeo‐Sale` et al., 1995), and 100 mM induces triglyceride (TG) accumulation and elevated ceramide levels. Ethanol‐induced accumulation of TG in cultured neurons suggests that alcohol may enhance lipogenesis and/or reduce fatty acid degradation in neurons. Instead, the increase of ceramide level may be related to the known ethanol‐induced apoptotic pathway (Pascual et al., 2003). In agreement with this hypothesis, the employment of myriocin, a drug that reduces ceramide levels, attenuates ethanol‐induced cell death (Saito et al., 2005) in human neuroblastoma SK‐N‐SH cells.

3.3 Brain 3.3.1 Cholesterol Early body of evidence of the chronic ethanol effects on brain lipids was the increase of cholesterol to phospholipid ratio in mouse brain (Chin et al., 1978). The change was attributable to an increase of cholesterol content and may explain the alteration of membrane fluidity observed.

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In developing brain, either the maturation of synapse or the proliferation and extent of basilar dendrites are negatively affected by ethanol (Davies and Smith, 1981; Inomata et al., 1987; Guizzetti and Costa, 1996; Kotter and Klein, 1999; Ikonomidou et al., 2000; Yanni and Lindsley, 2000); these data, together with the observation that the decrease of cholesterol levels results in an impairment of several neuronal and glial cell functions (Mauch et al., 2001; Fan et al., 2002), led to the hypothesis that alcohol may alter cholesterol homeostasis in the developing brain, and that such effect may be linked to its neurodevelopmental toxicity. Guizzetti et al. (2007) found that ethanol increases cholesterol efflux and the levels of active ABCA1 (a cholesterol transporter) in fetal human or rat astrocytes. The reported effects occurred in a range of ethanol concentrations (25–100 mM) that are physiologically relevant, because they can be found in the blood of moderate to heavy drinkers.

3.3.2 Phospholipids It has been suggested that hippocampus‐related cognitive processes are especially sensitive to ethanol (Sastry, 1985; Ahmad et al., 2002). The hippocampal system plays a critical role in learning and memory; PUFA, particularly docosahexaenoic acid (22:6n‐3, DHA), are highly concentrated in hippocampus (Sastry, 1985; Ahmad et al., 2002), specifically in PS and PE (Aid et al., 2003). As a proof of the sensitivity of hippocampus to ethanol, prenatal chronic ethanol exposure lowered the levels of total PC and PS by 15–20%, primarily owing to the reduction in 1‐stearoyl‐2‐docosahexaenoyl‐PS species, while at the same time, the levels of PE increased, although the total phospholipid content did not significantly change (Wen and Kim, 2004). Since both PC and PE are synthesized from diacylglycerol (DAG), the above changes may be explained by an ethanol‐dependent increased availability of DAG substrate in the PE pathway. Concerning the changes of PS levels, it is well established that membrane PS plays an important role in various signaling pathways supporting cellular function (McPherson et al., 1999; Bittova et al., 2001), and its accumulation promotes neuronal survival under adverse conditions (Kim et al., 2000; Akbar and Kim, 2002). It is therefore conceivable that the particular reduction of PS, together with alterations in PC and PE induced by ethanol, may have significant functional consequences in hippocampus (Cui and Houweling, 2002; Wen and Kim, 2004). The alteration of the phospholipid profile in the hippocampus resulting from exposure to ethanol during prenatal and developmental stages may have significant implication with respect to the cognitive dysfunction observed in fetal alcohol syndrome (FAS) (discussed further on).

3.3.3 Plasmalogens Plasmalogens are a unique class of phospholipids characterized by an ether linkage at the sn‐1 glycerol carbon. PE plasmalogen is highly abundant in nervous tissue, but its physiological role is unknown. Studies in vitro indicate that the ether bond of plasmalogens is more susceptible to oxidative cleavage than the acyl– glycerol bond of glycerophospholipids, and suggest that PE plasmalogen may have the ability to act as an antioxidant (Kuczynski and Reo, 2006). Such possibility was supported by Sun et al. (1987) who found PE plasmalogen increased (20–40%) in cortical synaptosomes from ethanol‐treated rats.

3.3.4 Glycosphingolipids Gangliosides, particularly abundant in CNS plasma membranes, are involved in synaptic transmission, neuritogenesis, receptor binding, and recognition events. The steady‐state concentration of gangliosides within cell membranes is maintained by de novo biosynthesis, degradation, and sphingosine recycling (Tettamanti and Riboni, 1994). Moreover, due to their enrichment in lipid raft‐specialized microdomains of the plasma membrane (Harder and Simons, 1997) they are involved in membrane Signalling and in many normal neurochemical, neurophysiological, and behavioral phenomena.

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3.4 Lipid Metabolism 3.4.1 Studies in vitro Studies on the effect of ethanol on ganglioside metabolism in cultured rat cerebellar granule cells indicate that the biosynthesis of the ceramide moiety, studied by incorporation of tritiated serine in the long chain base (LCB), dramatically decreases in the presence of alcohol. On the contrary, the biosynthesis of the oligosaccharide portion, studied by incorporation of tritiated galactose in the oligosaccharide moiety, is not affected by ethanol. Since ganglioside biosynthesis necessarily requires the availability of stoichiometric equivalents of LCB and oligosaccharide moieties, this finding suggests that ethanol enhances ganglioside biosynthesis by increasing the sphingosine recycling. This hypothesis was confirmed by experiments with exogenously administered GM1 ganglioside radiolabeled in the lipid moiety (Ravasi et al., 2002), demonstrating that gangliosides deriving from GM1 anabolism (GD1b, GT1b, GD1a) dramatically increase in ethanol‐treated cultured neurons. Ethanol‐induced increase of sphingosine recycling for ganglioside biosynthesis may constitute a mechanism of cell defence during alcoholic stress. In fact, the increased availability of the sphingoid base may have important consequences on cell events such as signal transduction and apoptosis, in which sphingosine and its derivatives, e.g., sphingosine 1‐phosphate, have been implicated (Ravasi et al., 2002).

3.4.2 Studies in vivo Chronic ethanol consumption increased the content of rat brain gangliosides, in particular trisialo derivatives, but a decrease of disialo amount. Marked aberrations in ganglioside pattern were detected in hypothalamus, thalamus, and hippocampus. While the functional significance of these biochemical changes is unknown, it can be postulated that ethanol induces and/or increases the activity of transferases involved in glycolipid, or inhibits the activity of sialidases involved in degradation (Vrbaski, 1995). Opposing to these results, Ghosh et al. (1998) and Marmillot et al. (1999) observed a change of ganglioside pattern in rats chronically treated with ethanol, associated to a 10–20% decrease in the sialyltransferase activity in whole brain, Golgi, and at synaptosomal level, and a concomitant dramatic increase in sialidase activity. A decrease of galactosyltransferase activity and neutral glycolipid content was reported either in acute (Klemm et al., 1988) and chronic experiments (Omodeo‐Sale` and Palestini, 1994).

3.5 Lipid Peroxidation and Antioxidants Brain contains particularly large amounts of PUFA and of catalytically active metal ions (Fe2þ, Cuþ, Al2þ), especially in the striatum and hippocampus. In addition to this, a high rate of oxygen consumption and a relatively low level of antioxidant enzymes render this tissue particularly vulnerable to lipid peroxidation (Ostrowska et al., 2004). Lipid peroxidation products are, in fact, involved in some of the pathophysiological effects associated with oxidative stress in cells. Chronic ethanol consumption leads to important changes in brain membrane lipid organisation, displaying its harmful effects through generation of reactive metabolites and free radical species, changing cell components structures and functions. The increase of lipid peroxidation products, such as lipid hydroperoxides and their highly reactive metabolites (malondialdehyde and 4‐hydroxynonenal) may act as ‘‘secondary toxic messengers’’ by diffusion from the site of their origin to the intracellular/extracellular targets (Esterbauer et al., 1991), where they disrupt structural and protective functions of the biomembranes. Moreover, acetaldehyde oxidation may lead to formation of acetaldehyde–protein adducts or malondialdehyde–acetaldehyde–protein adducts that increase the probability of cell injury (Tuma et al., 1987). The reduction and the degradation of phospholipid hydroperoxides are also carried out by glutathione peroxidase, and this is suggested to be a pathway of cytoprotection against the deleterious effects of phospholipid hydroperoxides. Decrease in this enzyme activity during ethanol intoxication would decrease the efficiency of this protective mechanism (Kinter and Roberts, 1996).

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Moreover, other mechanisms that finally promote enhanced oxidative damage are triggered by ethanol (Kato et al., 1990; Nordmann, 1994): induction of cytochrome P4502E1 (CYP2E1) the main enzyme of microsomal ethanol‐oxidizing system (MEOS), involved in many detoxificating processes generating free radicals (Montoliu et al., 1995); overexcitation of neurons, which triggers release of excitatory neurotransmitters, loss of Ca2þ homeostasis, altered intracellular signaling cascades, and cell death (Davis and Wu, 2001); stimulation of N‐methyl‐D‐aspartate (NMDA) receptor and neuronal nitric oxide synthase activity in cultured cortical neurons as a consequent nitric oxide (NO) production (Chandler et al., 1997); increase in arachidonic acid (AA) release and cyclooxygenase (COX‐2) activity (Basavarajappa et al., 1998; Sun et al., 2002). Recently, Milne et al. (2006) testing the hypothesis that lipid peroxidation is elevated in the brain tissue of rats fed with ethanol‐containing diet, found that levels of DHA oxidation products were significantly elevated in the cerebral cortex (97%) and brainstem (68%). Study of simultaneous administration of green tea with ethanol in rats resulted in normalization of lipid peroxidation process as well as glutathione concentration and glutathione peroxidase activity in liver and brain. Green tea antioxidants protect phospholipids from enhanced peroxidation and prevent changes in biochemical parameters and morphologic changes observed after ethanol consumption (Ostrowska et al., 2004). Studies in animal models also provide information indicating the ability of grape polyphenols to ameliorate neuronal damages due to chronic ethanol consumption; in fact, resveratrol, an important component of grape polyphenols, shows protective effects on neuron cell death induced by ethanol and other oxidative agents (Sun et al., 2002; Hancock and Miller, 2006). Finally, experimental data performed with vitamin E indicate a role for this compound to reverse the level of brain lipid peroxidation and of reduced glutathione level increased after ethanol consumption (Agar et al., 2003). Chronic pretreatment with vitamin E prevents alcohol‐induced vascular injury and pathology in the brain (Altura and Altura, 1999).

3.6 Fetal Alcohol Syndrome An important consequence of the maternal exposure to ethanol during pregnancy, is a CNS pathology named the FAS (Stibler et al., 1983). FAS is characterized by impaired development of the CNS, including neurologic abnormalities, developmental delays, and intellectual deficits, presumably due to altered embryogenesis (Leskawa et al., 1995). The wide variety of cellular/biochemical effects of ethanol on fetal tissues is itself a puzzle and strongly suggests that fetal responses to ethanol reflect a multifactor setting: ethanol affects fetal cell replication (Henderson et al., 1989), membrane transport systems (Heitman et al., 1987), membrane fluidity (Kelly‐ Murphy et al., 1984; Villacara et al., 1989), Naþ–Kþ pump expression (McCall et al., 1989), and EGF receptor expression (Calamandrei and Alleva, 1989; Paria et al., 1993). Henderson et al. (1999) has documented the oxidative stress and membrane lipid peroxidation in fetal brain (gestation day 19) following a 2‐day maternal ethanol consumption. However, the low levels of antioxidants in fetal tissues and an exaggerated response of fetal mitochondria to prooxidant stimulation in vitro, suggest that fetal cells are strongly prone to oxidative stress. Additionally, the fetal tissues are likewise inclined to the formation and subsequent accumulation of at least one toxic lipid peroxidation product, 4‐hydroxynonenal. As already above pointed out, ethanol exposure significantly decreases PS in developing rat brain cortices. In the hippocampus, prenatal chronic ethanol exposure lowered the levels of total PC and PS (particularly 18:0/22:6n‐3), and increased PE, although the total phospholipid content did not significantly change. The effect remarked on PS is due to an attenuation of microsomal biosynthetic activities, in developing rat brain cortices. Considering the role of 22:6n‐3 in proper neuronal development (Birch et al., 2000; Moriguchi et al., 2000) and the involvement of PS in various important signal transduction pathways (Newton and Keranen, 1994; Kim et al., 2000; Ligeti et al., 2004; Akbar et al., 2005) these observations may have significant implication in pathophysiologic effects of ethanol pertinent to FAS (Wen and Kim, 2004, 2007). Recently, Saito et al. (2007) demonstrated that GM1 and LIGA20 (a semisynthetic derivative of GM1 ganglioside), which have been shown to be neuroprotective against insults caused by various agents, partially attenuate ethanol‐induced apoptotic neurodegeneration in the developing mouse brain. Because

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gangliosides regulate activities of growth factors and neurotrophic factors (Hakomori et al., 1998; Ledeen et al., 1998), they may facilitate or mimic the antiapoptotic nature of these factors. Activation of Trk receptors by GM1 would stimulate survival pathways such as the PI3K/Akt pathway (Yao and Cooper, 1995) and the extracellular signal‐regulated kinase pathway (Climent et al., 2002), counteracting ethanol‐induced apoptotic neurodegeneration.

3.7 A Unique Phospholipid: Phosphatidylethanol Phosphatidylethanol is formed in cell membranes only in the presence of ethanol. The reaction is catalysed by phospholipase D, an enzyme that normally catalyses the hydrolysis of phospholipids, leading to the formation of phosphatidic acid. However, phospholipase D also utilizes short‐chain alcohols as substrates resulting in the formation of the corresponding phosphatidylalcohol. Significant amounts of phosphatidylethanol have been detected after ethanol exposure in brain and other organs from rat, in astrocytes primary cultures (Gustavsson and Hansson, 1990; Gustavsson, 1995) as well as in human blood cells. The degradation of phosphatidylethanol is relatively slow and it remains in the cells after ethanol has disappeared. It is possible that this abnormal phospholipid that accumulates in cell membranes affects membrane‐associated processes. Phosphatidylethanol is a lipid with a small anionic head group and its biophysical properties are different compared with other phospholipids. Moreover, this lipid has been shown to influence membrane characteristics, enzyme activities, and levels of signaling molecules, thus representing a possible intermediate through which ethanol may disturb cell function (Omodeo‐Sale` et al., 1991; Gustavsson, 1995). In the last years it has been proposed an interesting role of phosphatidylethanol as a sensitive and specific biomarker for ethanol abuse, in addition to gamma‐glutamyl transpeptidase, mean corpuscular volume and carbohydrate‐deficient transferrin. Indeed, none of the current biological markers used is enough sensitive and specific for the diagnosis of alcoholism. Blood concentrations of phosphatidylethanol are highly correlated to ethanol intake and its diagnostic sensitivity is higher than that for previously alcohol markers (Aradottir et al., 2006; Hartmann et al., 2007).

3.8 Ethanol and Dementia Consumers of light‐to‐moderate amounts of alcohol have a lower risk of dementia and, possibly Alzheimer disease (AD), than do abstainers or heavy drinkers (Huang et al., 2002; Ruitenberg et al., 2002; Mukamal et al., 2003). Subjects whose alcohol consumption was light to moderate had less severe white matter lesions and brain infarcts than did abstainers or heavy drinkers (Den Heijer et al., 2004). The possible involvement of lipids is suggested by findings from numerous studies, showing that lipid peroxidation may be implicated in the irreversible loss of neuronal tissue after brain or spinal cord injury as well as in degenerative neurologic disorders (Ostrowska et al., 2004). There is strong evidence that oxidative and inflammatory mechanisms play an important role in the pathogenesis of AD. Increased ROS production from b‐amyloid peptide (Ab) has been shown to result in protein oxidation and lipid peroxidation, and these effects can compromise cell membrane functions. Indeed, preliminary studies with primary neuron culture have provided data illustrating the ability of ethanol to exacerbate Ab toxicity and neuron cell death mechanisms (Sun and Sun, 2001).

4

Conclusions

Alcohol action is related to its ability to disorder cell membrane structure and thereby alter the physical properties of the membrane. The adaptation (tolerance) observed after chronic ethanol exposure was explained by the development of resistance to the disordering effects on the membrane, affecting lipid constituents, through cholesterol level variations, and phospholipids/gangliosides pattern.

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Moreover, ethanol significantly impairs the antioxidant defense system and, as a consequence, leads to a significant increase in the lipid peroxidation processes. The deleterious consequences of membrane peroxidation have stimulated investigations on the efficacy and mechanisms of action of biologically relevant antioxidants, particularly naturally occurring ones, including those used as foods and beverages, as green tea antioxidants, resveratrol (a grape polyphenol) or vitamin E.

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Section 5

Lipids in Neural Disfunction and Diseases

20

Roles of Cytosolic and Secretory Phospholipases A2 in Oxidative and Inflammatory Signaling Pathways in the CNS

G. Y. Sun . A. Y. Sun . L. A. Horrocks . A. Simonyi

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 518

2

PUFA in Brain Glycerophospholipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 519

3 3.1 3.2 3.3 3.4 3.5

Cytosolic cPLA2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 520 Properties of cPLA2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 520 cPLA2 and Oxidative Signaling Pathways in Astrocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 521 cPLA2 and Oxidative Signaling Pathways in Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 521 cPLA2 and NADPH oxidase in Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 521 cPLA2 and Cerebral Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 523

4 4.1 4.2 4.3 4.4 4.5 4.6

Secretory PLA2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 523 Properties of sPLA2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 523 sPLA2 in Inflammatory Responses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 524 sPLA2 Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 525 Neurotoxic Action of Exogenous sPLA2-IIA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 525 sPLA2 in Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 525 sPLA2 and Cerebral Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 526

5

Cross Talk between sPLA2 and other PLA2 in Mediating Inflammatory Responses . . . . . . . . . . . . . 526

6

PLA2 Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 527

7

Summary and Projections for Future Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 527

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_20, # Springer ScienceþBusiness Media, LLC 2009

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Roles of cytosolic and secretory PLA2 in oxidative and inflammatory signaling pathways in the CNS

Abstract: Oxidative stress is implicated in the pathophysiology of a number of neurodegenerative diseases. There is evolving evidence for a metabolic link between reactive oxygen species (ROS) and phospholipases A2 (PLA2), enzymes that hydrolyze fatty acyl groups from the sn-2 position of membrane glycerophospholipids. Increased production of ROS and upregulation of PLA2 are important factors underlying the progression of stroke and Alzheimer’s disease. The major goal of this review is to provide recent information on the oxidative and inflammatory pathways associated with PLA2 activation in neurons and glial cells, particularly, the group IV cytosolic PLA2 and the group II secretory PLA2. Special emphasis is placed on signaling pathways and NADPH oxidase, an enzyme producing superoxide anions. Understanding the involvement of these PLA2 in the oxidative and inflammatory environment in central nervous system is an important step for developing novel therapeutic strategy for the treatment and prevention of neurodegenerative diseases. List of Abbreviations: AMPA, 2-amino-3-hydroxy-5-methyl-4-isoxazole propionate; CDR, clinical dementia ratings; CNS, central nervous system; DHA, docosahexaenoic acid; FFA, free fatty acids; HNE, hydroxynonenal; LTP, Long-term potentiation; M-CSF, macrophage colony stimulating factor; MDA, malondiadehyde; MPTP, 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine; PCR (qPCR), quantitative realtime; PKC, protein kinase C; PLA2, phospholipases A2; PLC, phospholipase C; PUFA, polyunsaturated fatty acids; ROS, reactive oxygen species; SPAN, snake presynaptic PLA2 neurotoxin; TR-FIA, time-resolved fluoroimmunoassay

1

Introduction

Due to the high rate of oxygen consumption, the enrichment of polyunsaturated fatty acids (PUFA) in neural membranes, and the relatively low abundance of anti-oxidant enzymes, cells in the central nervous system (CNS) are highly susceptible to oxidative stress. Reactive oxygen species (ROS) may play a role as signaling molecules in regulating cell metabolism under normal conditions, but excessive production is generally harmful, leading to cell damage and apoptosis. Excessive ROS production is known to cause an increase in intracellular Ca2+ and activation of Ca2+-dependent enzymes, including protein kinases, phosphatases, proteases, phospholipases, and nucleases (Mattson, 2007). These oxidative-induced reactions have been the underlying cause for the development and progression of many neurodegenerative diseases, including Alzheimer’s, Parkinson’s, multiple sclerosis, and stroke (Zipp and Aktas 2006; Aktas et al., 2007). Elucidation of the mechanism(s) for ROS production in different cell systems, including cells in the CNS, is attracting considerable interest. Recent studies provide evidence for the involvement of superoxidegenerating NADPH oxidase, a multi-subunit enzyme originally identified in immune cells for innate host defense system and bacteria killing (Quinn et al., 2006). This enzyme is now recognized to be widespread in most nonphagocytic cells, including cells in the cardiovascular systems and in CNS (Quinn et al., 2006; Bedard and Krause, 2007; Lambeth 2007). Depending on the cell type, many G-protein-coupled receptor agonists stimulate downstream signals for assembly of the NADPH oxidase units. There is increasing evidence that ROS derived from NADPH oxidase can lead to mitochondrial dysfunction and trigger apoptosis and other cell death mechanisms (Infanger et al., 2006). Increased NADPH oxidase activity is implicated in a number of diseases in the cardiovascular, renal, pulmonary, and CNS systems (Bedard and Krause, 2007). PLA2 are responsible for the hydrolysis of the sn-2 fatty acids from glycerophospholipids. These enzymes are ubiquitous in mammalian cells. They play important roles in the synthesis of lipid mediators and maintenance of cell membrane integrity. Despite more than 20 different PLA2 in different cell systems, studies to investigate the role of PLA2 in health and disease are generally limited to three major groups, i.e., the Ca2+-dependent group IV cytosolic cPLA2, the Ca2+-independent group VI iPLA2, and the low molecular weight group II secretory sPLA2 (Murakami et al., 1997; Murakami and Kudo, 2002). With advancements in molecular and analytical tools for analysis of specific phospholipids, new information about the role of PLA2 in different pathological conditions is becoming evident (Han, 2007). Studies with cPLA2 knockout mice revealed the role of cPLA2 in regulating a number of physiological functions

Roles of cytosolic and secretory PLA2 in oxidative and inflammatory signaling pathways in the CNS

20

(Hirabayashi et al., 2004). Several recent reviews focus on the role of PLA2 in brain injury and neurodegenerative diseases (Sun et al., 2004; Farooqui and Horrocks, 2006; Farooqui et al., 2007; Sun et al., 2007). In this chapter, the particular emphasis is to examine the roles of cPLA and sPLA2 in oxidative and inflammatory pathways in neurons and glial cells and to evaluate their impact on the pathophysiology of Alzheimer’s disease (AD) and stroke.

2

PUFA in Brain Glycerophospholipids

Glycerophospholipids in brain membranes are highly enriched in PUFA, especially, arachidonic acid (AA, 20:4, n-6) and docosahexaenoic acid (DHA, 22:6, n-3). With multiple double bonds, these fatty acids are highly susceptible to oxidative stress and free radical attack. Earlier studies have provided evidence for the rapid release of these fatty acids in response to neuronal excitation and injury (Bazan et al., 1981). These results led to the recognition that fatty acids in brain membrane glycerophospholipids undergo active turnover through the deacylation–reacylation cycle mediated by PLA2 and acyltransferases, respectively (Sun and MacQuarrie 1989; Farooqui et al., 2000) (> Figure 20-1). Under conditions such as brain injury

. Figure 20-1 The deacylation–reacylation reactions for turnover of membrane phosphoglycerides: mediated by phospholipases A2 and lysophospholipid:acylCoA acyltransferase

and stroke, the deacylation–reacylation cycle is disturbed. The decreased energy supply can cause the failure of conversion of free fatty acids (FFA) to their acyl-CoAs (Phillis et al., 2006). The resulting accumulation of FFA and lyso-glycerophospholipids may be the source of neuronal damage. Using tracer techniques, PUFA such as AA are rapidly taken up from the circulation system and incorporated into brain glycerophospholipids. There is dynamic metabolism of the fatty acids in brain under normal and pathological conditions (DeGeorge et al., 1989; Rapoport 2001; Rapoport et al., 2001; Rosenberger et al., 2002). AA is an important lipid mediator in the CNS. Besides serving as substrate for eicosanoid synthesis, AA and its oxidative metabolites are regarded as retrograde messengers and play a role in synaptic function including modulating neurotransmitter release (Williams et al., 1989; Nishizaki et al., 1999; Bazan, 2003; Bazan 2006), ion channel activity (Ferroni et al., 2003) and long-term potentiation (LTP) (Sang and Chen, 2006). AA is enriched in PtdIns and polyphosphoinositides (poly-PI) that are substrates for phospholipase C (PLC). A number of G-protein-coupled receptors are coupled to PLC. Hydrolysis of PtdIns-P2 produces diacylglycerols and inositol trisphosphate, second messengers for activation of protein kinase C (PKC) and mobilization of intracellular Ca2+, respectively (Berridge, 1987).

519

520

20

Roles of cytosolic and secretory PLA2 in oxidative and inflammatory signaling pathways in the CNS

Although DHA is not a substrate for prostanoid synthesis, DHA deficiency is associated with cognitive decline and altered visual aquity. DHA is enriched in PtdSer and EtnGpl and is especially abundant in the retina and brain (Kim, 2007). In astrocytes, different signaling pathways and PLA2 mediate the release of DHA as compared with pathways for the release of AA (Strokin et al., 2003). DHA released by PLA2 is oxidized to docosanoids including neuroprotectin D1, a compound with anti-oxidative properties and a powerful neuroprotective agent (Mukherjee et al., 2004; Bazan, 2005; Lukiw et al., 2005; Bazan, 2006, 2007). Oxidation of PUFA results in formation of lipid peroxidation products, for example malondiadehyde (MDA) and 4-hydroxynonenal (HNE). Enhanced lipid peroxidation has been shown in a number of disease states (Zarkovic, 2003). Isoprostanes are unique oxidative products of AA and are biomarkers of lipid peroxidation and oxidative damage in a number of human diseases including AD (Montine et al., 2004; Montine et al., 2005; Casadesus et al., 2007; Milne et al., 2007). Neuroprostanes are oxidative products derived from DHA. Because the CNS is enriched in DHA, measurement of neuroprostanes may be a more selective index of brain oxidative injury as compared with isoprostanes (Greco and Minghetti, 2004).

3

Cytosolic cPLA2

3.1 Properties of cPLA2 The group IV cytosolic cPLA2 are Ca2+-dependent enzymes ubiquitously present in the cell cytoplasm of mammalian cells (Hirabayashi et al., 2004). cPLA2a is a 85 kDa protein and contains a C2 domain that is responsible for binding Ca2+ and a catalytic domain with multiple phosphorylation sites for MAPKs, Ca2+-calmodulin kinase II, and MAPK-interacting kinase Mnk1 (Leslie, 1997, Hirabayashi et al., 2004). Upon binding with Ca2+ and phosphorylation by protein kinases, cPLA2 is translocated to membrane glycerophospholipids to release fatty acids and lyso-glycerophospholipids. Due to its active involvement in receptor signaling pathways, many studies have focused on understanding the functional role of this enzyme in different physiological and pathological conditions. Besides cPLA2a, which is the major isoform of group IV PLA2, several other isoforms, namely, cPLA2b, g, d, e, and z have been identified in different cell systems (Ghosh et al., 2006). Many cPLA2 isoforms exhibit varying degrees of lysophospholipase and transacylase activities. Upon stimulation, cPLA2a targets endoplasmic reticulum, Golgi (Evans et al., 2004), and nuclear membranes (Grewal et al., 2005). Recent studies demonstrated cellular localization of other cPLA2 isoforms; for example, cPLA2b is constitutively bound to mitochondria and early endosomes and hydrolyzes exclusively PtdEtn instead of PtdCho (Ghosh et al., 2006). cPLA2g is also constitutively bound to mitochondrial membranes, but this enzyme is Ca2+-independent (Tucker et al., 2005). In activated mouse lung fibroblasts, cPLA2z is translocated to ruffles and other dynamic vesicular structures (Ghosh et al., 2007). In neutrophils, cPLA2a is found in the plasma membranes and in the nucleus (Levy, 2006). The dual localization for this enzyme suggests that cPLA2 may play distinct functional roles in different subcellular compartments, for example for regulating receptor and enzyme activity in the plasma membranes and for regulating synthesis of eicosanoids in the cytoplasm (Levy, 2006). Despite yet unknown mechanisms, several cytoskeletal proteins including vimentin and annexin are known to bind cPLA2 (Nakatani et al., 2000). In rabbit smooth muscle cells stimulated by norepinephrine, cPLA2 is translocated to the nuclear envelope through binding with actin (Fatima et al., 2005). cPLA2 also plays a role in the cholesterol-rich membrane domain enriched with the scaffold protein, caveolin-1 (Graziani et al., 2004). Localization of PLA2 in the caveolin-1 domain of mouse primary hippocampal neurons modulates the activity of the 2-amino-3-hydroxy-5-methyl-4-isoxazole propionate (AMPA) receptor, one of the ionotropic glutamate receptors responsible for synaptic activities and LTP in the hippocampus (Gaudreault et al., 2004). However, whether binding of cPLA2 with cytoskeletal proteins may modulate neuroexocytosis has not been examined in detail. cPLA2a can also bind lipids, for example ceramide-1-phosphate is shown to bind to a cationic patch of the C2 domain of cPLA2a (Pettus et al., 2004). Binding of cPLA2 with ceramide-1-phosphate can cause a positive allosteric modulation and renders its interaction with PtdCho at lower Ca2+ concentration

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(Subramanian et al., 2005; Stahelin et al., 2007). cPLA2 can also bind anionic glycerophospholipids, especially PtdIns-P2 (Mosior et al., 1998). Exactly how the binding of cPLA2 with anionic glycerophospholipids alters catalytic activity of the enzyme has not been investigated in detail.

3.2 cPLA2 and Oxidative Signaling Pathways in Astrocytes In astrocytes, cPLA2a is activated by Ca2+-ionophores, phorbol esters, and agonists for G-protein-coupled receptors, including the P2Y2 receptor (Xu et al., 2002, Sun et al., 2004). There is also evidence linking cPLA2 activation with oxidant compounds, including H2O2 and menadione, a compound known to produce intracellular ROS by redox cycling with oxidases (Han et al., 2003; Xu et al., 2003b). Exposure of astrocytes to H2O2 led to time-dependent changes in membrane physical properties, rearrangement of F-actin, and formation of filapodia and nanotubules and these physiological changes were dependent on the p38 MAPK pathway (Zhu et al., 2005). Menadione can stimulate superoxide anion production in astrocytes through activating NADPH oxidase (Sun, unpublished data). In turn, ROS produced by NADPH oxidase can stimulate ERK and p38 MAPK and subsequently cPLA2. In neutrophils, activation of the angiotensin II receptor pathway is also linked to NADPH oxidase and cPLA2 through signaling pathways involving ERK and p38 MAPK (Hazan-Halevy et al., 2005).

3.3 cPLA2 and Oxidative Signaling Pathways in Neurons There is evolving evidence for the important role of PLA2 in maintaining different neuronal functions, for example regulation of neuronal excitation, mediation of neurite outgrowth, and neuronal viability. Prolonged inhibition of cPLA2 and iPLA2 by inhibitors such as MAFP and BEL can cause the decrease in neuronal survival in primary cultures of neurons (Forlenza et al., 2007a). Due to the putative role of AA and its metabolites as retrograde messengers, it is reasonable to direct efforts to better understand how PLA2 is regulated in neurons under oxidative conditions. Despite yet unknown mechanisms, excessive ROS production is linked to neuronal excitotoxicity due to activation of ionotropic glutamate receptors (NMDA, kainic acid, and AMPA) (Kishida et al., 2005). Studies from our laboratory demonstrated the involvement of NADPH oxidase in producing ROS under these excitatory conditions, and in turn, this pool of ROS is important in activation of cPLA2 and AA release (> Figure 20-2). Our study also demonstrated the ability of oligomeric amyloid beta peptide (Ab), a key molecule in Alzheimer’s disease, to stimulate ROS production and AA release in neurons through a similar signaling mechanism (Shelat et al., unpublished data). Consistent with the role of cPLA2 in neuronal excitotoxicity, hippocampal cultures from cPLA2a / mice or neuronal cultures treated with cPLA2 inhibitor were significantly protected from NMDA toxicity (Brady et al., 2006). Furthermore, electrophysiological studies with pyramidal neurons also provided support for the involvement of cPLA2a in NMDA-induced excitotoxicity (Shen et al., 2007). Besides studies with neurons in culture, an in vivo approach using radiolabeled AA has successfully been used to investigate glycerophospholipid metabolism in the CNS with different disease states (Rapoport, 2005). Measurement of PLA2 mRNA and protein expression supports the role of PLA2 in affective disorders. Chronic administration (21 days) of fluoxetine, an antidepressant drug, was shown to upregulate cPLA2 mRNA and protein expression in the rat frontal cortex (Rao et al., 2006). Furthermore, chronic neuronal excitation by administration of a daily sub-convulsive dose of NMDA to rats for 21 days increased cPLA2 activity, phosphorylated protein, and mRNA levels (Rao et al., 2007). These results in vitro and in vivo provide strong support for the role of cPLA2 in neuronal excitatory mechanisms.

3.4 cPLA2 and NADPH oxidase in Alzheimer’s Disease Increased oxidative stress is regarded as an early event for the development of AD (Butterfield et al., 1999, Montine et al., 2005). At least part of the oxidative effect is attributed to cytotoxicity elicited by Ab (Varadarajan et al., 2000). Although the mechanism whereby Ab elevates oxidative stress remains elusive,

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. Figure 20-2 Involvement of ROS from NADPH oxidase and cPLA2 in neuronal excitotoxicity

this compound, when folded or aggregated in particular conformation, can enhance Ca2+ influx into neurons (Mattson, 2007). Study with hippocampal neurons provided evidence for Ab to enhance Ca2+ influx through activating an L-type voltage-sensitive Ca2+ channel (Ueda et al., 1997). Activation of Ca2+-mediated pathways and increased ROS production are important underlying causes for the cytotoxicity of Ab in neurons. Abnormal signaling pathways mediated by Ab and NMDA receptor may lead to synaptic loss and neuronal damage (Shankar et al., 2007). Recent studies are beginning to unveil a novel link between Ab, ROS production, and activation of NMDA receptor in mediating the apoptotic pathways in neurons (De Felice et al., 2007). Studies by Kriem et al. (Kriem et al., 2005) further provided evidence for the involvement of cPLA2 in Ab-mediated neuronal apoptosis. Oxidative stress induced by Ab can also lead to mitochondrial dysfunction, and at least one part of the change in mitochondrial membrane potential was attributed to activation of cPLA2 and iPLA2 (Zhu et al., 2006). Soluble Ab could increase the magnitude and duration of endothelin-1-induced vasoconstriction and this pro-inflammatory pathway involves MAPK, cPLA2, and AA release (Paris et al., 2000). In conjunction with the pro-inflammatory pathway mediated by Ab, brain slices from APPsw transgenic mice also showed increased levels of COX-2 and TNFa (Quadros et al., 2003). Due to evidence supporting the involvement of ROS and cPLA2 in AD, there is increasing interest to examine whether antioxidants and antiinflammatory agents may offer protective effects against Ab-induced damage (Bate and Williams, 2007). More studies are needed to examine the expression of cPLA2 mRNA/protein and its phosphorylation status in AD brain. A microarray study demonstrated upregulation of cPLA2 mRNA in hippocampus of AD brain (Colangelo et al., 2002). On the other hand, Forlenza’s group (2007) observed decreased PLA2 activity in the frontal and parietal areas of AD brain (Forlenza et al., 2007b). The decreased activity reflected decreased glycerophospholipid turnover and subsequently changes in membrane properties and neuronal function (Forlenza et al., 2007a). Using the ‘‘shotgun lipidomic protocol,’’ Han et al. (2001) observed an

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obvious decrease in plasmalogen content in the AD cortex with mild clinical dementia ratings (CDR 0.5) (Han et al., 2001). Similar decrease in plasmalogen was also observed in transgenic AD mouse models. The underlying mechanism leading to the dramatic decline in plasmalogen and the type of PLA2 regulating plasmalogen metabolism remains to be investigated.

3.5 cPLA2 and Cerebral Ischemia Although increased oxidative stress is an important mechanism underlying neuronal damage in cerebral ischemia, little is known about the status of cPLA2 expression in neurons and glial cells after ischemic injury. Studies with transgenic mice deficient in cPLA2 provide information supporting the important role of cPLA2 in neuronal damage induced by cerebral ischemia (Bonventre and Sapirstein, 2002). Mice deficient in cPLA2 are more resistant to insults caused by cerebral ischemia and are more protected from neuronal damage by 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP), a neurotoxin used as a model for Parkinson’s disease (Sapirstein and Bonventre, 2000; Tabuchi et al., 2003). Studies with transient global cerebral ischemia using the gerbil model demonstrated an increase in lipid peroxidation products, which preceded mitochondrial dysfunction and neuronal cell death (Wang et al., 2002). Consistent with the increase in ROS in mediating ischemic damage, antioxidants such as resveratrol and curcumin were useful in inhibiting lipid peroxidation and ameliorating ischemic damage (Wang et al., 2002; Wang et al., 2005; Wang et al., 2006). In addition, apocynin, a known inhibitor for NADPH oxidase, can also ameliorate ischemia-induced neuronal damage, suggesting that NADPH oxidase serves as an important source of ROS (Wang et al., 2006). The increase in ROS in ischemia/reperfusion is linked to the increase in cPLA2 and release of FFA (Bazan et al., 1981).

4

Secretory PLA2

4.1 Properties of sPLA2 sPLA2 belong to the family of PLA2 that are Ca2+-requiring, low-molecular weight (14–19 kDa), and have 6–8 disulfide bridges. Unlike cPLA2, which is more specific in AA release, sPLA2 have wide fatty acid selectivity. To date, more than 10 secreted sPLA2s, including sPLA2-IB, IIA, IIC, IID, IIE, IIF, III, V, X, and XII, have been identified and differentially expressed in different mammalian cell systems (Murakami et al., 1997; Murakami and Kudo, 2001). Among these isoforms, sPLA2-IIA is most studied because of its association with inflammatory diseases including arthritis, atherosclerosis, sepsis, and different types of infections (Hurt-Camejo et al., 2001; Taketo and Sonoshita, 2002). Under inflammatory conditions, high levels of sPLA2-IIA are detected in plasma. sPLA2 are enriched in immune active cells, including macrophages, neutrophils, and astrocytes (Murakami and Kudo, 2002). In human kidney cells, sPLA2-IIA, IID, and V respond to stimuli to release AA and PGE2 in a manner dependent on glypican, a heparan sulfate proteoglycan, whereas sPLA2-IB, IIC, and IIE bind weakly to heparin and fail to elicit AA release (Murakami et al., 2001). sPLA2-IIF is unique because it contains a long C-terminal extension implicating its location in the plasma membranes (Murakami et al., 2002). This sPLA2 also participates in inflammatory reactions, but is independent of glypican. Using quantitative real-time PCR (qPCR) and a sensitive time-resolved fluoroimmunoassay (TR-FIA), analyses of mRNA and protein expression of sPLA2 (group 1B, GIIA, GIID, GIIE, GIIF, GV and GX) in serum and tissues of Balb/c and C57BL/6J mice revealed the prevalence of expression of several isoforms of sPLA2 in the gastrointestinal tract (Eerola et al., 2006). Group V sPLA2 is more widely distributed in all organs in Balb/c and C57BL/6J mice. In general, very low levels of these sPLA2 isoforms are found in the brain. Using TR-FIA to assay of sPLA2 isoforms in human sera from patients with septic infection, high levels of sPLA2-IIA were found in septic patients as compared to controls (Nevalainen et al., 2005).

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Apparently, mitochondria in rat brain cortex contain a form of sPLA2-IIA that is released upon reduced membrane potential (Macchioni et al., 2004). In cultured cerebellar granule neurons, sPLA2-IIA is observed in the peripheral regions of neurons and its participation in ROS production is linked to glutamate-induced neuronal death (Mathisen et al., 2007).

4.2 sPLA2 in Inflammatory Responses In astrocytes as well as in other immune cells, pro-inflammatory cytokines such as IL-1a and TNFa can induce sPLA2-IIA mRNA through the NF-kB pathway (Li et al., 1999; Schwemmer et al., 2001; Xu et al., 2003a; Ghesquiere et al., 2005; Tietge et al., 2005; Lee et al., 2006). The induction of sPLA2-IIA by proinflammatory cytokines may be regulated by intracellular kinases including PKC isoforms and MAPK (Huwiler et al., 1997; Kuwata et al., 1998). In many instances, these same protein kinases are involved in phosphorylation of cPLA2 and NADPH oxidase subunits. Thus, it is not surprising that the transcription pathway for induction of sPLA2 may involve cross talk with cPLA2 as well as ROS from NADPH oxidase (> Figure 20‐3). More studies are needed to exploit these cross-talk mechanisms.

. Figure 20-3 Transcriptional pathway for synthesis of sPLA2: possible involvement of NADPH oxidase

sPLA2 seems to target glycerophospholipids when cells are injured and membrane glycerophospholipids are perturbed, for example when cells undergo apoptosis and when acidic glycerophospholipids are exposed on the outer leaflet of the plasma membranes (Fourcade et al., 1998). Inclusion of anionic glycerophospholipids vesicles, such as vesicles containing increasing levels of PtdSer in PtdCho, can dramatically enhance interfacial binding and catalysis of the sPLA2-IIA (Bezzine et al., 2002). Besides the release of fatty acids, lyso-glycerophospholipids produced by sPLA2 have detergent-like properties and their accumulation can alter membrane structure, transbilayer glycerophospholipids migration, increase Ca2+ influx, and enhance microvesicle release and apoptosis (Wilson et al., 1999). Increased

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lyso-glycerophospholipids may also change the biophysical properties of membranes, causing an increase in membrane lipid spacing, solvation of the lower head group region of the membrane, and a greater vertical mobility of glycerophospholipids (Bailey et al., 2007).

4.3 sPLA2 Receptors sPLA2 can bind to specific membrane receptors, namely, the M type receptor that is present in muscle cells and the N type receptor that is generally neuronal. The M-type receptor (180-kDa protein) has been cloned from different animal species (Cupillard et al., 1999). A comparison of the binding properties of recombinant sPLA2 with mouse M-type receptor indicated differences in binding activity for different sPLA2 ranging from 0.3 to > 100 nM in the following order 1B, IIA, IIE, IIF, X > IIC, V > IID, III, XIIA, XIIB (Rouault et al., 2007). Besides mediating AA release, sPLA2 may also activate inflammatory responses, including the induction of degranulation of mast cells and activation of exocytosis of macrophages. Part of these responses can be attributed to an interaction of sPLA2 with its receptor and not the enzymic activity (Triggiani et al., 2006).

4.4 Neurotoxic Action of Exogenous sPLA2-IIA Extracellular forms of sPLA2-IIA, including those from human and Taipan snake venom, are neurotoxic and can modulate activities of ionotropic glutamate receptors and Ca2+ channels and trigger neuronal apoptosis (Kolko et al., 2002; Rodriguez De Turco et al., 2002; Yagami et al., 2002; Yagami et al., 2003). In some instances, sPLA2 can cause punctate swelling of neurites and enhance exocytosis of synaptic vesicles (Rigoni et al., 2004; Ikeno et al., 2005). Ammodytoxin (Atx), a snake venom sPLA2 known to exert presynaptic neurotoxicity (Petan et al., 2005), can display a strong preference for vesicles containing anionic glycerophospholipids, for example PtdSer, over the zwitterionic PtdCho. The protein snake presynaptic PLA2 neurotoxin (SPAN) is highly cytotoxic and can paralyze neuromuscular junctions. A large part of SPAN action is due to its PLA2 activity, targeting PtdCho and generating fatty acids and lyso-PtdCho (Rigoni et al., 2004; Megighian et al., 2007). Interestingly, an equimolar mixture of lyso-PtdCho and fatty acids (oleic acid) can closely mimic the biological effects of SPAN, including inducing Ca2+ influx and mediating synaptic vesicle release (Rigoni et al., 2007). LysoPtdCho + FA can also account for the changes in cell morphology and collapse of mitochondrial membrane potential within nerve terminals. Several types of sPLA2, including those from bee venom, Naja naja, and porcine and human pancreatic sPLA2 can induce endothelial cell migration. Similarly, endothelial cell migration can be mimicked by hydrolytic products of sPLA2, including AA, lyso-PtdCho, and lyso-PtdH (Rizzo et al., 2000).

4.5 sPLA2 in Alzheimer’s Disease Glial cell-mediated inflammatory responses are known to play an important role in the pathophysiology of AD (Rogers et al., 1996; Akiyama et al., 2000; Lue et al., 2001a; Mrak and Griffin, 2005). Increased levels of IL-1b were observed in the CNS associated with cognitive decline (Holmes et al., 2003). Ab activates the IL-1b pathway and causes a persistent increase in NF-kB in glial cells (Samuelsson et al., 2005). Microglial cells isolated from human postmortem brain respond to aggregated Ab to produce pro-inflammatory IL-1b, IL-6, TNFa, and macrophage colony stimulating factor (M-CSF) (Lue et al., 2001b). A study by Moses and colleagues (2006) demonstrated an increase in sPLA2-IIA mRNA and immunoreactivity in astrocytes in AD brain as compared with age-matched, nondemented (ND) controls. In agreement with studies using immortalized rat astrocytes, DITNC cells (Li et al., 1999), studied by Moses et al. (2006) also reported the ability for IL-1b and Ab to induce sPLA2-IIA in astrocytes isolated from human postmortem brain.

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4.6 sPLA2 and Cerebral Ischemia Different types of neuronal insults including cerebral ischemia have been shown to increase sPLA2-IIA mRNA and protein expression (Yagami et al., 2002; Lin et al., 2004; Adibhatla et al., 2006; Adibhatla and Hatcher, 2007). In the rat focal cerebral ischemia model induced by occlusion of the middle cerebral artery, sPLA2-IIA immunoreactivity was colocalized with the GFAP-positive cells in the penumbral area (Lin et al., 2004). Under similar conditions, cerebral ischemia-reperfusion also caused the increase in COX-2 mRNA although expression of cPLA2 mRNA was not changed (Lin et al., 2004). In agreement with an increased inflammatory response, treatment with TNFa antibody or IL-1 receptor antagonist significantly attenuated infarction volume and sPLA2-IIA protein expression and activity after transient middle cerebral artery occlusion. These results suggest a contribution of sPLA2-IIA to inflammatory responses in ischemic injury (Adibhatla and Hatcher, 2007). Although cerebral ischemia increased activity and expression of various PLA2 isoforms, more studies are needed to clarify the cellular origin and localization of these isoforms. PLA2 inhibitors with high specificity for sPLA2 offer neuroprotective effects against cerebral ischemiainduced neuronal lesions and apoptosis (Cunningham et al., 2004; Yagami et al., 2005).

5

Cross Talk between sPLA2 and other PLA2 in Mediating Inflammatory Responses

Cross-talk mechanisms exist between sPLA2 and cPLA2 in mediating AA release and regulate production of eicosanoids. Apparently, both sPLA2 and cPLA2 can cause the release of AA, which is converted to prostanoids by COX-1 and COX-2. In COX-1(/) mice, a compensatory upregulation of COX-2, accompanied by the activation of the NF-kB pathway was observed. COX-1 deficiency can affect the expression of COX-2, Ca2+-dependent PLA2, and terminal mPGES-2, to overcome defects in brain AA cascade (Choi et al., 2006). Using gene disruption and specific inhibitors, the involvement of cPLA2 and sPLA2-IIA in mediating AA release and PGE2 production in response to cytokines was shown in gastric epithelial cells and in transfected human embryonic kidney cells (Ni et al., 2006). Induction of sPLA2-IIA in response to cytokines is responsible for the late phase of AA release and production of PGE2 in these cells. Using HEK293 cells overexpressing sPLA2-IIA, there is indication that release of AA and PGE2 occurred before secretion of the enzyme into the extracellular space (Ni et al., 2006). However, there was indication in rat fibroblasts that induction of sPLA2 by pro-inflammatory stimuli involved the group VIB iPLA2 instead of cPLA2 (Kuwata et al., 2007). cPLA2 becomes phosphorylated within minutes after the addition of LPS and the phosphorylated form of cPLA2 shows enhanced activity in vitro (Dieter et al., 2002). In our earlier study, exposure of astrocytes to cytokines resulted in increased expression of sPLA2 and COX-2, but not COX-1 and cPLA2, and a timedependent increase in PGE2 (Xu et al., 2003a). Although astrocytes could respond to ATP or phorbol ester (PMA), which stimulated cPLA2 phosphorylation and AA release, relatively small increase in PGE2 was observed. However, when astrocytes were first treated with cytokines, subsequent exposure to ATP or PMA markedly increased the levels of PGE2 (Xu et al., 2003a). This study demonstrated the important role of cPLA2, sPLA2, and COX-2 in PGE2 production. Results from microarray analysis of gene expression in the cerebral cortex and hippocampus of mice deficient in COX-1 (COX-1/) or COX-2 (COX-2/) suggest that COX-1 and COX-2 differentially modulate brain gene expression (Toscano et al., 2007). The decrease in PGE2 synthesis in cPLA2/ brain is attributed to the decreased mRNA and protein levels of COX-2 but not COX-1 (Bosetti and Weerasinghe, 2003). These results suggest a coordinate induction of both cPLA2 and COX-2 mRNA by pro-inflammatory cytokines for PGE2 synthesis. Both cPLA2 and sPLA2 may be upregulated in neuroinflammation induced by cytokines. Ventricular infusion of LPS increased reactive microglia in the cerebral ventricles, pia mater, and changes in morphology of astrocytes in the cortical mantel and areas surrounding the cerebral ventricles (Rosenberger et al., 2004). LPS infusion also increased brain cPLA2 and sPLA2 activities as well as levels of nonesterified linoleic and AA, and prostaglandins E2 and D2 (Rosenberger et al., 2004).

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20

PLA2 Inhibitors

In light of the involvement of cPLA2 and sPLA2 in oxidative and inflammatory responses, control of their activities is a prerequisite for diminishing the extent of neurodegeneration (Tanaka and Arita 1995; Reid 2005; Farooqui et al., 2006; Nanda et al., 2007; Narendra Sharath Chandra et al., 2007). Besides production of lipid mediators, some sPLA2 can directly activate inflammatory cells, making therapeutic mechanisms to antagonize these enzymes an even more important endeavor (Triggiani et al., 2006). However, the presence of multiple isoforms of sPLA2 and their involvement in modulating diverse cell functions make this approach a formidable task (Yedgar et al., 2006). Reid (2005) presented a structurally diverse array of available sPLA2-IIA inhibitors and their therapeutic potential in phase II clinical trials in humans. A list of more recently used inhibitors is described in > Table 20-1. . Table 20-1 Phospholipase A2 inhibitors

Citicoline (cytidine50 -diphosphocholine)

Target action on types of PLA2 A pyrrolidine-based inhibitor for cPLA2 Mode of action unknown – not a direct PLA2 inhibitor

Indoxam

sPLA2-IIA

Me-Indoxam

sPLA2 and M-type receptor

CHEC-9

Internal fragment CHEASAAQC of the human neuroprotective polypeptide DSEP (Diffusible Survival Evasion Peptide) N-PtdEt linked to polymeric carriers – membrane anchored lipid conjugates

Name of inhibitor Pyrrolidine 1

ExPLAIs – Extracellular PLA2 inhibitors

7

Brief description of study Inhibit cPLA2 in different cell types Provides neuroprotection by attenuating the stimulation of PLA2 Neuroprotective effects against cerebral ischemic damage Decreases the affinity of sPLA2 for the recetpor Inhibit inflammatory response associated with stab wound lesion and microglial infiltration Inhibit CNS inflammatory responses associated with stab wound lesion and microglial infiltration

References Ghomashchi et al., 2001 Adibhatla et al., 2003

Yagami et al., 2002

Boilard et al., 2006 Cunningham et al., 2004

Pinto et al., 2003

Summary and Projections for Future Research

Understanding the source of ROS that underline neurodegeneration – There is strong evidence for NADPH oxidase to produce ROS in neurons and glial cells in the CNS and the involvement of this pool of ROS in the progression of a number of neurodegenerative diseases including AD and stroke. Assembly of NADPH oxidase complex requires phosphorylation of its cytosolic subunits by Ca2+-dependent pathways and protein kinases. More studies are needed to examine factors regulating assembly of these subunits and activation of NADPH oxidase under physiological and neuropathological conditions. Recognizing the important role of oxidative pathways for activation of cPLA2 in neurons and glial cells – Recent studies provide information linking oxidative signaling pathways with protein kinases that stimulate phosphorylation of cPLA2. More studies are needed to understand how the signaling pathways for

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activation of cPLA2 are controlled and the role of cPLA2 and its products for regulating neuron and glial cell functions. a. Understanding the ‘‘double-edged sword’’ mechanism for neurotoxic action of Ab – There is evolving evidence supporting the involvement of oxidative stress mediated by NADPH oxidase and subsequent membrane damage mediated by PLA2 in the neurotoxic action of Ab. Understanding this ‘‘double-edged sword’’ effect of Ab may provide new insights linking oxidative stress and neuronal dysfunction in the development of AD. Understanding the role of sPLA2 in neurodegeneration – It is recognized that sPLA2-IIA is induced in brain in response to different forms of neuronal injury. More studies are needed to examine the factors and signaling pathways underlying the induction process. Studies to develop specific inhibitors for this class of sPLA2 may provide therapeutics to combat neurodegenerative diseases. Recognizing the anti-oxidative and anti-inflammatory effects of botanical polyphenols – There is increasing evidence for dietary polyphenols as molecular targets to combat age-related neuroinflammatory diseases. Future studies should focus on the understanding whether these compounds may exert specific effects on cPLA2, sPLA2, and NADPH oxidase.

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J. R. Van Brocklyn

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 536

2 Fatty Acids and Fatty Acid Derivatives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 536 2.1 Polyunsaturated Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 536 2.2 Eicosanoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 538 3 Sphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 541 3.1 Sphingolipid Metabolites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 541 3.2 Glycosphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 544 4

Glycerolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 548

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Sterols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 549

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Conclusions and Future Directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 550

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_21, # Springer ScienceþBusiness Media, LLC 2009

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Lipids in neural tumors

Abstract: It has long been appreciated that the formation of neural tumors is accompanied by profound changes in the composition of a wide variety of different lipids. Recent years have seen a major increase in knowledge of the molecular biological roles played by lipids in these tumors, which has led to the identification of lipids, the enzymes that metabolize them and the pathways they regulate as potential diagnostic and prognostic markers and as possible therapeutic targets for these devastating malignancies. This chapter will summarize the current knowledge in this field. List of Abbreviations: CDK, cyclin‐dependent kinase; COX, cyclooxygenase; CREB, cyclicAMP response element binding protein; EGF, epidermal growth factor; ERK, extracellular signal‐regulated kinase; GBM, glioblastoma multiforme; GLA, g‐linolenic acid; GSL, glycosphingolipids; JNK, Jun N‐terminal kinase; LOX, lipoxygenase; MAP, mitogen‐activated protein; MMP, matrix metalloproteinase; NK, natural killer cell; NSAIDs, nonsteroidal anti‐inflammatory drugs; PDGF, platelet‐derived growth factor; PI, phosphatidylinositol; PKC, protein kinase C; PNET, primitive neural ectodermal tumor; PPARg, proliferator activated receptor; PTEN, phosphatase and tensin homolog deleted on chromosome 10; PUFAs, polyunsaturated fatty acids; ROS, reactive oxygen species; S1P, sphingosine‐1‐phosphate; SphK, sphingosine kinase; VEGF, vascular endothelial growth factor

1

Introduction

A variety of tumor types occur within neural tissue. Metastatic tumors represent approximately 50% of brain neoplasms, with the remaining 50% being primary neural tumors. Of primary tumors the most common category is gliomas, composed of cells resembling various glial cells. The most malignant glioma, glioblastoma multiforme (GBM), is also the most common primary brain tumor occurring in adults. GBM is a highly malignant tumor displaying aggressive growth and invasion of surrounding brain tissue, leading to a median survival time of less than 12 months. As GBMs are relatively common and highly malignant, much research involving the role of lipids in neural malignancies has focused on these tumors. However, some information is available on other tumors including oligodendrogliomas, ependymomas, meningiomas, and the pediatric tumors neuroblastoma and medulloblastoma as well as some peripheral nervous system tumors. It has long been recognized that the lipid composition of neural tumors differs from that of normal brain tissue. Brante (1949) found that intracranial tumors contained higher levels of esterified cholesterol than normal brain. Gopal et al. (1963) found that gliomas and meningiomas contained a high percentage of unsaturated fatty acids in comparison to normal brain, and Lou et al. (1965) found that astrocytomas contained an increased amount of phosphatidylcholine and sphingomyelin and decreased phopshatidylethanolamine in comparison to normal brain tissue, with a more pronounced change in more malignant tumors. In the decades since these early studies, significant advancements have been made in our understanding of not only the changes in lipid composition associated with neural malignancies, but also the biological functions of lipids within these tumors and even the potential use of lipids or modification of biological pathways regulated by lipids as therapeutic approaches. This chapter will summarize the current knowledge in these areas. In several cases, recent reviews exist, which thoroughly discuss the current understanding of the role of certain lipids in neural tumors. In such cases, we have briefly summarized the field and refer the reader to these reviews for further details.

2

Fatty Acids and Fatty Acid Derivatives

2.1 Polyunsaturated Fatty Acids A large number of studies have examined the effects of polyunsaturated fatty acids (PUFAs), in particular the o‐3 fatty acids eicosapentaenoic acid and docosahexaenoic acid and the o‐6 fatty acids g‐linolenic acid (GLA) and arachidonic acid, on tumor cells including glioma cells (reviewed in Das, 2004). PUFAs, cause

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apoptosis of a wide variety of tumor cell lines (Das, 2004) including glioma lines, but do not affect normal astrocytes (Vartak et al., 1998), suggesting their effects are targeted to tumor cells. In agreement, PUFAs caused regression of C6 gliomas in rats without apparent toxicity to normal surrounding brain tissue (Leaver et al., 2002a, b). In addition, arachidonic acid causes apoptosis in primary tumor cells derived from human glioma tissue, but had less of an effect on cells derived from surrounding normal brain tissue (Williams et al., 1998). Although the apoptotic effects of PUFAs on tumor cells may involve a variety of mechanisms, the ability of antioxidants to inhibit the apoptotic response points to the generation of reactive oxygen species (ROS) as important mediators of this effect (Das, 2004). Recently, tumor cells from glioma tissue were shown to be more sensitive to arachidonic acid‐stimulated production of ROS than nontumor cells (Leaver et al., 2004). Oxidative effects were also implicated in the cytotoxicity of cis‐paranaric acid, a plant PUFA, to rat and human glioma cell lines (Traynelis et al., 1995). Cis‐paranaric acid also was not cytotoxic for primary rat astrocytes (Traynelis et al., 1995). One mechanism of PUFA cytotoxicity to tumor cells may be enhancement of the apoptotic effect of radiation and chemotherapy. PUFAs have been shown to sensitize cell lines derived from several types of cancer to chemotherapeutic drugs (Das, 2004). PUFAs also enhanced the radiosensitivity of rat 36B10 astrocytoma cells (Vartak et al., 1997). Furthermore, docosahexaenoic acid sensitizes A172 and U‐87 cells to cytotoxic effects of doxorubicin (Rudra and Krokan, 2001). Therefore, it has been proposed that PUFAs may be useful as chemotherapeutic agents for glioma patients, particularly in combination with standard chemotherapy drugs and/or radiation (Das, 2004). A few small studies have attempted to address the usefulness of PUFAs therapeutically in human patients with malignant glioma. The first attempt treated six patients with recurrent malignant glioma with intratumoral injection of GLA. All six patients showed a significant reduction in tumor size as determined by CT scans, without significant side effects (Naidu et al., 1992). A larger study treated 15 patients with primary malignant glioma (Das et al., 1995). GLA was implanted in the brain following surgical resection and a cerebral catheter was placed into the tumor bed to allow for subsequent GLA treatments. In all patients, CT scans revealed an increase in necrotic areas of tumors and decreased midline shift, suggesting a decreased mass effect of the tumor. Twelve of the fifteen patients survived 16–20 months following surgery (Das et al., 1995). These data are encouraging, however, large scale clinical trials are necessary to validate the usefulness of this approach. Although the data discussed above clearly implicate PUFAs as cytotoxic agents for gliomas, arachidonic acid levels have been shown to be elevated in glioma and meningioma tissue compared to normal brain (Ledwozyw and Lutnicki, 1992; Ko¨koglu et al., 1998). In addition, gliomas contain high levels of linoleic acid, a o‐6 PUFA and the precursor or arachidonic acid, in comparison to nonmalignant brain tissue (Martin et al., 1996). Therefore, gliomas must have a mechanism to avoid the cytotoxic effects of these lipids. It is likely that glioma cells are able to metabolize arachidonic acid via the cyclooxygenase and lipoxygenase pathways in order to prevent apoptotic effects of these fatty acids. Interestingly, the metabolism of arachidonic acid into the bioactive lipids prostaglandins and leukotrienes has significant implications for brain tumors as discussed later. Arachidonic acid can also be metabolized into the bioactive lipids known as endocannabinoids, specifically N‐arachidonoyl‐ethanolamide, also known as anandamide, and 2‐arachidonoyl‐glycerol (reviewed in De Petrocellis et al., 2004). These lipid mediators signal through the G protein‐coupled cannabinoid receptors CB1 and CB2 (De Petrocellis et al., 2004). Recent evidence indicates that cannabinoids decrease the growth and induce apoptosis of several types of tumor cells including glioma cells in vitro and in vivo and may be therapeutically useful (reviewed in Velasco et al., 2004). Level of endocannabinoids may be altered in brain tumors, however, discrepancies exist in the literature. One study found that endocannabinoids were decreased in human meningioma and GBM in comparison to normal brain as measured by GC/MS (Maccarrone et al., 2001). On the other hand, Petersen et al. (2005) found that GBMs have enhanced levels of anandamide while meningiomas have increased 2‐arachidonoyl glycerol. Nevertheless, agreement exists that cannabinoids are antiproliferative and apoptotic for glioma cells while sparing or even protecting nontumor brain cells. D9‐tetrahydrocannabinol, the cannabinoid found in

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the cannabis plant, induced apoptosis of C6 glioma and human glioma cells but not primary astrocytes or neurons (Sanchez et al., 1998; McAllister et al., 2005). Several other cannabinoids including the endocannabinoids anandamide, and 1‐arachidonoyl glycerol (Fowler et al., 2003), nonpsychoactive synthetic cannabinoids that work through the CB2 receptor (Recht et al., 2001; Sanchez et al., 2001), stearoylethanolamide (Maccarrone et al., 2002), and the cannabinoid cannabidiol (Massi et al., 2004) also kills glioma cells and decrease growth of glioma xenografts in nude mice. In addition, the cannabinoid KM‐233, which is selective for CB2 receptors, inhibits growth of U‐87 MG tumors in SCID mice with minimal toxicity (Duntsch et al., 2005). The mechanism of cannabinoid induced apoptosis in glioma cells appears to involve generation of the proapoptotic lipid ceramide (Velasco et al., 2004). Cannabinoids activate sphingomyelin hydrolysis in C6 glioma cells (Sanchez et al., 1998), and result in sustained ceramide accumulation (Galve‐Roperh et al., 2000). In addition, cannabinoid‐induced apoptosis was blocked by an inhibitor of ceramide synthesis fumonisin‐B1 (Hinz et al., 2004). Interestingly, cannabinoids also block proliferative and survival signaling by the extracellular signal‐regulated kinase (ERK), mitogen‐activated protein (MAP) kinase, and Akt and activate the proapototic function of Bad by decreasing its phosphorylation in C6 glioma cells (Ellert‐Miklaszewska et al., 2005). In contrast however, one study found that cannabinoids enhanced the proliferation of U‐373 MG glioma cells via transactivation of the epidermal growth factor (EGF) receptor (Hart et al., 2004), and cannabinoids protect primary astrocytes from apoptosis induced by exogenously added ceramide (Gomez Del Pulgar et al., 2002). Cannabinoids may also decrease glioma angiogenesis. Cannabinoids decreased vascular endothelial growth factor (VEGF) production in glioma cells in vitro and in mice, and this response was also blocked by fumonisin‐B1 (Blazquez et al., 2004). Cannabinoids also decreased vascular growth in gliomas growing in nude mice (Blazquez et al., 2003). Addition of exogenous ceramide also decreased VEGF production (Blazquez et al., 2004). Furthermore, cannabidiol blocks migration of U‐87 MG cells by a cannabinoid receptor‐independent mechanism (Vaccani et al., 2005).

2.2 Eicosanoids The PUFA arachidonic acid is liberated from the sn‐2 position of glycerol‐based phospholipids by phospholipase A2. Arachidonic acid is metabolized to the bioactive lipids eicosanoids by the action of the enzymes cyclooxygenase (COX) and lipoxygenase (LOX), which lead to the formation of prostaglandins and leukotrienes respectively. COX enzymes have been implicated in a variety of cancers, and considerable effort has been directed toward using COX inhibitors therapeutically (reviewed in Zha et al., 2004; Hull, 2005; Mazhar et al., 2005). Two isoforms of COX, also called prostaglandin H synthase, exist. COX‐1 is constitutively expressed in a variety of tissues, while COX‐2 is an inducible enzyme. Considerable evidence indicates that COX enzymes, particularly COX‐2, play important role in cancer. COX‐2 is overexpressed in several forms of cancer, including colon, prostate, breast, lung, bladder, skin, and pancreas (Zha et al., 2004; Mazhar et al., 2005). Nonsteroidal anti‐inflammatory drugs (NSAIDs), which act as nonspecific COX inhibitors, have been associated with reduced cancer risk (Thun et al., 2002). Specific COX‐2 inhibitors have been shown to inhibit carcinogenesis in several animal models and COX‐2 genetic knockout confirms decreased carcinogenesis (reviewed in Hull, 2005). In addition, COX‐2 inhibitors have been used as cancer chemopreventative agents, particularly to prevent colon cancer in patients with familial adenomatous polyposis (Steinbach et al., 2000; Phillips et al., 2002; Hallak et al., 2003; Higuchi et al., 2003). Nevertheless, COX‐independent effects of COX‐2 inhibitors as well as concerns of the cardiovascular effects of COX‐2 inhibitors may limit the effectiveness of such therapy (Hull, 2005). The prostaglandin products of COX enzymes, particularly PGE2 but also PGD2, PGF2, PGI2, and thromboxane TXA2, have been implicated in playing direct role in several forms of cancer, particularly colon, lung, and skin (Zha et al., 2004). These prostaglandins may regulate proliferation, cell survival, and tumor angiogenesis (Zha et al., 2004). Numerous studies have investigated the potential roles played by COX and LOX enzymes and eicosanoids in brain tumors (reviewed in Nathoo et al., 2004). COX‐2 expression was detected in several types of

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brain tumor including GBM, astrocytomas, meningiomas, medulloblastomas, craniopharyngiomas, ependymomas, neurinomas, oligodendrogliomas, and primitive neural ectodermal tumors (Matsuo et al., 2001; Patti et al., 2002). COX‐2 immunostaining correlates with MIB‐1 staining, an index of cell proliferation, in GBMs (Prayson et al., 2002). COX‐2 expression was shown to correlate with high histological grade (Joki et al., 2000) and proliferative index of gliomas and with poorer survival of glioma patients (Shono et al., 2001). However, one study found that GBMs had weak immunohistochemical staining for COX‐2 in comparison to less malignant grade 2 and 3 astrocytomas (Matsuo et al., 2001). Another study found that COX‐2 expression as determined by immunohistochemistry was highly variable in both primary and recurrent GBMs, however, tumors with higher percentage of cells expressing COX‐2, greater that 25%, showed a shorter time to recurrence than those with less than 10% (Sminia et al., 2005). Thus, COX‐2 may play a significant role in the malignant behavior of astrocytomas. COX enzymes may play roles in other brain tumors as well. Increased COX‐1 expression levels were also found to correlate with increased malignancy of oligodendrogliomas and with shorter patient survival, however COX‐1 immunostaining was predominantly observed in macrophages/microglia within these tumors (Deininger et al., 2000). COX‐2 immunostaining was evident in oligodendroglioma tumor cells and correlated with poor patient survival (Castilla et al., 2003). COX‐2 was also expressed in medulloblastomas but not in surrounding normal cerebellum, and correlated with poor survival (Deng et al., 2004). COX‐2 expression was also detected in pediatric ependymomas and growth of primary ependymoma cell lines was inhibited by a COX‐2 inhibitor (Kim et al., 2004). Eicosanoid producing enzymes have been shown to contribute to the proliferation and survival of brain tumor cells. COX and LOX inhibitors blocked proliferation of human glioma cells in culture (Blomgren and Kling‐Andersson, 1992). In addition, COX‐2 inhibitors decrease occurrence and size of gliosarcomas injected intracranially in rats (Nam et al., 2004) and significantly prolong survival (Portnow et al., 2002). A specific COX‐2 inhibitor decreased both monolayer and tumor spheroid growth of U‐87 MG and U‐251 MG human glioma cells and caused an increase in apoptosis (Joki et al., 2000). Peripheral neural tumors also have been associated with COX‐2 overexpression including neuroblastoma, ganglioneuroma, and pheochromocytoma (Johnsen et al., 2005). Nonspecific COX‐1/2 inhibitors, as well as specific COX‐2 inhibitors, caused apoptosis of neuroblastoma cells in vitro, and potently decreased growth of neuroblastoma xenografts in rats (Johnsen et al., 2005). Neuroblastoma cells also contain elevated levels of the substrate of COX enzymes, arachidonic acid (Reynolds et al., 2001). Interestingly, inhibition of COX caused apoptosis in neuroblastoma cells, and this effect was more pronounced in combination with LOX inhibition and addition of arachidonic acid (Johnsen et al., 2005), suggesting that effects of COX inhibition are at least partly attributable to induction of apoptosis by increased arachidonic acid and not entirely to decreases in downstream eicosanoids. In agreement, although COX‐2 inhibitors blocked GBM cell line proliferation, and decreased levels of the prostaglandin PGE2, the potency of various inhibitors to block proliferation did not correlate with degree of suppression of prostaglandin formation, and this effect was not reversed by adding back PGE2 (Kardosh et al., 2004). COX‐2 inhibitors may also have nonspecific effects. COX‐2 inhibitors have recently been shown to also inhibit phosphoinositide‐dependent kinase (Arico et al., 2002; Kulp et al., 2004), and thus some effects of COX‐2 inhibition might occur through inhibition of the PI 3‐kinase/Akt pathway or other non‐COX‐2 dependent effects. In agreement with the potential nonspecific effects of COX‐2 inhibitors, the inhibitor NS398, decreased growth of glioma cells regardless of whether or not the cells expressed COX‐2 (Matsuo et al., 2004). Several studies have shown direct effects of prostaglandins on brain tumor cells. Gliomas and meningiomas were found to express increased levels of several prostaglandins and thromboxanes in comparison to normal brain, with no change in the relative proportion of the different eicosanoids studied (Castelli et al., 1989). The increased expression correlated with tumor histological grade (Castelli et al., 1989). Ko¨koglu et al. (1998) found increased levels of PGE2 in gliomas and meningiomas, although meningiomas had higher levels. Plasma levels of PGE2 were found to be elevated in patients with malignant gliomas and to decrease following surgical removal (Loh et al., 2002). The authors suggested that PGE2 was involved in immunosupression in glioma patients (Loh et al., 2002). Matsuo et al. (2004) recently found that an antagonist specific for the PGE2 receptor EP1 decreased glioma cell growth in vitro and glioma xenograft

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growth in mice. Surprisingly, however, addition of PGE2 decreased growth of the same glioma cell lines in vitro (Matsuo et al., 2004), an effect which the authors attributed to possible different effects of PGE2 through other receptor subtypes. Prostaglandins of the cyclopentenone group, PGA2 and PGJ2, are inhibitory to cell proliferation and have been suggested as potential cancer therapeutic agents (Ishihara et al., 2004). PGD2, from which the cyclopentenone prostaglandins are derived, inhibits proliferation and stimulates differentiation of rat C6 glioma cells (Conde et al., 1991). PGA2 repressed proliferation and IGF‐1 expression in C6 cells (Bui et al., 1997). PGJ2 blocked proliferation and induced differentiation in several glioma cell lines, at least partially through activation of the peroxisome proliferator activated receptor, PPARg (Morosetti et al., 2004). PPARg is commonly expressed in GBM tissue (Kato et al., 2002), but not in adjacent normal brain (Morosetti et al., 2004). Interestingly, PGJ2, induced apoptosis in U‐87 and A172 glioma cells through expression of the pro‐apoptotic protein Bax, but did not affect normal mouse astrocytes (Zander et al., 2002). Thus, PGJ2, or other PPARg agonists may be useful therapeutic agents for gliomas that may spare normal brain tissue. In this regard, conjugated linoleic acid, an isomer of linoleic acid which also functions as a PPARg agonist, inhibits glioma cell growth and stimulates apoptosis (Cimini et al., 2005). Injection of a phospholipase activating protein into a rat glioma increased levels of PGE2 and LTB4 and caused tumor regression (Goddard et al., 1996). This was accompanied by an acute inflammatory reaction with an infiltration of natural killer (NK) cells. Thus, while prostaglandins may contribute to the pathological behavior of gliomas they may also stimulate inflammatory reactions that could be therapeutically useful. Prostaglandins have also been linked to the induction of glioma angiogenesis. Significant evidence links COX‐2 to induction of angiogenesis by a variety of mechanisms, including direct effects of prostaglandins on endothelial cells and stimulation of production of angiogenic factors (reviewed in Zha et al., 2004). In this regard, COX‐2 immunostaining of astrocytic gliomas was found to correlate with VEGF expression and vascularity (Hara and Okayasu, 2004). COX‐2 expression in gliomas was detected in tumor cells in areas surrounding necrosis (Deininger et al., 1999). These cells have been implicated in the production of VEGF, which in turn mediates glioma angiogenesis (Fischer et al., 2005). Furthermore, the combination of temozolamide with the COX‐2 inhibitor rofecoxib led to a longer time to recurrence in GBM patients with more vascular primary tumors but not in those with low vessel density (Tuettenberg et al., 2005). Another study found that COX‐2 and thromboxane synthase are expressed in endothelial cells of high grade gliomas, and COX‐2 and thromboxane synthesis inhibitors synergistically block endothelial cell migration (Jantke et al., 2004). Thromboxane synthesis inhibitors also induced endothelial cell apoptosis (Jantke et al., 2004). Furthermore, PGE2 (Fiebich et al., 1997) and thromboxane A2 (Obara et al., 2005) both induce IL‐6 in human glioma cells. IL‐6 induces VEGF transcription in U‐87 MG glioma cells (Loeffler et al., 2005). In addition, IL‐6 expression drives autocrine proliferation of several glioma cell lines (Goswami et al., 1998), correlates with malignancy in gliomas (Rolhion et al., 2001), and is necessary for glioma development in a mouse model (Weissenberger et al., 2004). Prostaglandins may also modulate the resistance of glioma cells to radiation. The radioresistant glioma cell line T98G had higher levels of COX‐2 expression than the radiosensitive cell line A172, and COX‐2 inhibitors increased the sensitivity of T98G cells to radiation (Karim et al., 2005). COX‐2 inhibition enhanced radiation sensitivity of U‐251 MG cells in vitro and decreased xenograft growth synergistically with radiation treatment (Petersen et al., 2000). Evidence also exists, linking eicosanoids to the development of brain tumor peritumoral edema, a significant clinical problem. Prostaglandin levels were found to correlate with the degree of edema in meningioma patients (Constantini et al., 1993). Levels of the leukotriene LTC4 (Black et al., 1986) and the thromboxane TXB2 (Zhao et al., 1998) also correlated with edema in brain tumor patients. LTC4 was also found to increase transcytosis across brain–tumor barrier (Hashizume and Black, 2002). Primary microglia isolated from rats with intracranial C6 gliomas produced high levels of PGE2, and COX‐2 inhibition decreased blood–tumor barrier permeability (Badie et al., 2003). In addition, a COX‐2 inhibitor led to clinical and radiologic improvement in a child with peritumoral edema and necrosis following radiation therapy for a metastatic brain tumor (Khan et al., 2004).

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Although most of the data discussed above relate to prostaglandins, leukotrienes have also been implicated in brain tumors. 5‐LOX is expressed in astrocytic gliomas and positivity correlates with histological grade; however, after correcting for patient age, 5‐LOX staining did not significantly correlate with survival (Nathoo et al., 2006). Nevertheless, LOX inhibitors decreased in vitro proliferation, and LTB4 stimulated increased [Ca2þ]i and cell proliferation of U‐373 MG human glioma cells (Kim et al., 1998). On the other hand, leukotrienes may have beneficial effects for brain tumor patients in combination with chemotherapy drugs. LTC4 has been shown to selectively open the blood–tumor barrier while not affecting the blood–brain barrier (Black et al., 1990) and LTE4 increases permeability of blood–tumor barrier and delivery of chemotherapeutic drugs to an experimental rat tumor (Chio et al., 1995). LTC4 pretreatment before chemotherapy with cisplatin enhances survival in rat C6 glioma model (Black et al., 1998). Moreover, LOX inhibitors have been shown to block fas‐mediated apoptosis in human glioma cell lines (Wagenknecht et al., 1997). Thromboxanes, which are derived from PGH2 by the action of thromboxane synthase, may also contribute to glioma invasion. Similarly to COX‐2 inhibitors, thromboxane synthase inhibitors cause apoptosis in human glioma cells (Kurzel et al., 2002), however, they also block glioma cell motility (Giese et al., 1999), while COX‐2 inhibitors have no effect on motility (Kurzel et al., 2002). Thus thromboxanes may be involved in glioma invasiveness. In summary, eicosanoids may participate in the pathobiological behavior of brain tumors through a variety of mechanisms. Moreover, inhibition of the enzymes leading to formation of these bioactive lipids can have profound effects and may be therapeutically useful both by decreasing levels of certain eicosanoids as well as by increasing levels of their fatty acid precursors. Further work will be necessary to fully understand the roles played by the various PUFAs and eicosanoids in brain tumors. Thus, drugs targeting synthesis or action of individual eicosanoids may eventually allow for the specific clinical manipulation of brain tumor cell behaviors as well as responses to the tumors such as angiogenesis, inflammatory reactions, and edema.

3

Sphingolipids

3.1 Sphingolipid Metabolites In recent years a large amount of work has been dedicated to elucidating the biological roles played by sphingolipid metabolites. The majority of these studies have focused on two metabolites, ceramide and sphingosine‐1‐phosphate (S1P). Most studies on ceramide have shown that this lipid has growth inhibitory and apoptosis‐inducing properties, while S1P has primarily been associated with the opposite effects of mitogenesis and cell survival. These counteracting effects of two closely related lipids, and the fact that they can be easily interconverted, have led to the theory of the sphingolipid rheostat, i.e., the proposal that the balance between the cellular levels of ceramide and S1P determines cell fate, with ceramide favoring inhibition of growth and cell death, and S1P favoring survival and proliferation (reviewed in Maceyka et al., 2002). Significant evidence exists that this rheostat plays a role in regulating the growth of a variety of tumor cell types as well as apoptosis of tumor cells, or resistance to apoptosis induced by chemotherapeutic drugs (Maceyka et al., 2002; Cuvillier and Levade, 2003). Several studies have examined the roles played by ceramide in brain tumor cells. Differences in ceramide composition that may be pathologically significant exist in brain tumors. Ceramide levels in human glioma tissue have been shown to be lower than in normal surrounding brain tissue and to be inversely proportional to histological grade and patient survival (Riboni et al., 2002), suggesting that gliomas may downregulate ceramide as a means of avoiding apoptotic cell death. In addition, the individual ceramide species, distinguished by the length and saturation of the amide‐linked fatty acid chain differ in gliomas and normal brain tissue. Thus, ceramides in GBMs contain an overabundance of monounsaturated fatty acids (C18:1) in comparison to normal brain and low grade astrocytomas as determined by 1H NMR (Lombardi et al.,

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1997). In addition, differences were detected between several glioma cell lines with regard to the fatty acids present in ceramides using liquid chromatography tandem mass spectrometry (Sullards et al., 2003). As little is known regarding the differences in biological effects of various ceramide species, the significance of these observations is not currently clear. Ceramide has been shown to lead to death of glioma cells by various mechanisms. Ceramide mediates apoptotic cell death in glioma cells induced by a variety of stimuli including, TNF‐a (Sawada et al., 2004), gamma irradiation (Hara et al., 2004), the topoisomerase inhibitor etoposide (Sawada et al., 2000), and chemotherapeutic drugs such as cisplatin (Noda et al., 2001). In addition, as discussed above, generation of ceramide appears to be crucial to mediate cannabinoid‐induced glioma cell apoptosis (Sanchez et al., 1998; Galve‐Roperh et al., 2000; Hinz et al., 2004; Velasco et al., 2004), although it is not entirely clear if this response is mediated by ceramide generated by sphingomyelin hydrolysis or by de novo synthesis. Ceramide generated by hydrolysis of sphingomyelin also mediates apoptosis of human glioma cells induced by metabolites of vitamin D (Magrassi et al., 1998). Ceramide and Fas ligand have a synergistic effect on apoptosis of human glioma cells through separate signaling pathways (Wagenknecht et al., 2001). On the other hand, another group found that apoptosis induced by Fas in glioma cells required sphingomyelinase‐ mediated ceramide formation (Sawada et al., 2002). Similarly, TNF‐a‐induced ceramide formation and apoptosis involves caspase‐8, p53 and superoxide generation leading to activation of neutral sphingomyelinase (Sawada et al., 2001). Ceramide may also enhance apoptosis of glioma cells indirectly by inhibiting activation of prosurvival signals. Thus, ceramide caused dephosphorylation and inactivation of the prosurvival kinase Akt through a ceramide‐activated protein phosphatase in U‐87 MG glioma cells (Zinda et al., 2001). While most of the above described studies have been conducted in cell lines derived from astrocytic gliomas, ceramide can also regulate apoptosis in other neural tumor cells. Apoptosis induced by staurosporine or TNF‐a in human oligodendroglioma (HOG) cells is mediated by activation of caspase‐8 in rafts which signals through neutral sphingomyelinase to increase ceramide leading to caspase‐3 activation (Testai et al., 2004). Similar apoptotic signaling through neutral sphingomyelinase was induced by staurosporine in neuroblastoma cells (Kilkus et al., 2003). Moreover, increased ceramide causes translocation of the lipid phosphatase and tumor suppressor gene PTEN to lipid rafts resulting in enhanced caspase‐3 activity and apoptosis in PC12 cells and HOG cells (Goswami et al., 2005). Ceramide also causes caspase‐independent apoptosis of D283 medulloblastoma cells that involves the calpain proteases (Poppe et al., 2002). Nevertheless, some neural tumor cell lines are resistant to the apoptotic effects of ceramide. One group found that siRNA blocking expression of the x‐linked inhibitor of apoptosis sensitized several ceramide‐resistant glioma cell lines to ceramide‐induced cell death (Hatano et al., 2004). Ceramide may also cause nonapoptotic death of glioma cells in some instances. Ceramide causes nonapoptotic cell death of A172 human glioma cells that did not involve caspases or loss of mitochondrial membrane potential (Kim et al., 2005). A similar nonapoptotic cell death was induced by ceramide in two additional human glioma cell lines, U251 and T98G (Mochizuki et al., 2002). Moreover, cell death induced by ceramide in U‐373 MG and T98G cells was found to be autophagic death and to be mediated by induction of the cell death‐inducing mitochondrial protein, Bcl‐2/adenovirus E1B 19 kDa‐interacting protein 3 (BNIP3) (Daido et al., 2004). Ceramide not only induces death of glioma cells but, in some cases, causes inhibition of cell proliferation. Addition of cell permeable ceramide or sphingomyelinase activation induced by the low affinity p75NTR neurotrophin receptor induces inhibition of glioma cell growth (Dobrowsky et al., 1994). Nitric oxide treatment of C6 glioma cells caused ceramide accumulation, which resulted from decreased trafficking of ceramide from endoplasmic reticulum to Golgi leading to inhibition of cell proliferation (Viani et al., 2003), suggesting that either accumulation of ceramide or decrease in production of sphingomyelin or glucosylceramide mediates NO‐induced inhibition of glioma cell growth. Removal of the amide‐linked acyl chain from ceramide by ceramidase yields sphingosine, which can then be phosphorylated by sphingosine kinase (SphK) to create S1P (Le Stunff et al., 2004). S1P signals by binding to a group of five G protein‐coupled receptors (GPCRs), termed S1P1–5, which couple to several G proteins including, Gi, Gq, and G12/13 (Windh et al., 1999). In this way, S1P regulates a variety of cellular responses including cell proliferation, survival, motility, and differentiation (Spiegel and Milstien, 2003;

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Hla, 2004). In addition, significant evidence exists that S1P can signal via a receptor‐independent, intracellular mechanism, although the direct intracellular target of S1P remains unidentified. Significant evidence indicates that S1P has profound effects on glioma cells. S1P treatment of rat C6 glioma cells activates phospholipase C/Ca2þ signaling, ERK, MAP kinase, protein kinase C (PKC), and phospholipase D (Sato et al., 1999b) and reverses the change to an astrocyte‐like morphology induced by b‐adrenergic agonists (Tas and Koschel, 1998). C6 cells express a small amount of the receptor S1P1, which couples exclusively to Gi (Windh et al., 1999), high levels of S1P2 (Sato et al., 1999b), which couples to all three of the above mentioned G proteins but tends to favor G12/13 (Taha et al., 2004), as well as S1P5 (Malchinkhuu et al., 2003), which couples to Gi and G12 (Malek et al., 2001). Through these receptors and signaling pathways S1P stimulates expression of several immediate early genes. S1P‐stimulated Gi and PKC signaling leads to ERK activation, EGR‐1 expression, and FGF‐2 expression in C6 cells (Sato et al., 1999a). S1P also enhances c‐fos expression (Segura et al., 2005). The effect of S1P on ERK/EGR‐1/FGF‐2 is mediated by S1P1, while S1P2 mediates activation of phospholipase C, increased [Ca2þ]i and phospholipase D (Sato et al., 2000). In addition, S1P likely mediates the effects of HDL on activation of ERK, Ca2þsignaling, and FGF‐2 expression in C6 cells (Malchinkhuu et al., 2003). Consistent with activation of the above pathways, enforced overexpression of SphK, the enzyme which forms S1P, in C6 cells drives cell proliferation in the absence of serum (Vann et al., 2002). Moreover, S1P is released extracellularly by astrocytes (Anelli et al., 2005; Bassi et al., 2006) and C6 glioma cells (Edsall et al., 2001). Thus, the potential for an autocrine loop of S1P signaling that enhances growth and/or survival of glioma cells exists. Our group has focused on the roles played by S1P and SphK in regulating the malignant behavior of human gliomas. Using a panel of human glioma cell lines we determined that S1P was mitogenic for approximately 50% of the cell lines tested (Van Brocklyn et al., 2002). In addition, S1P stimulates motility and invasiveness through Matrigel of a similar percentage of human glioma cell lines (Van Brocklyn et al., 2003). Interestingly, these responses to S1P did not always occur in the same cell lines. Whether a glioma cell line responds to S1P with proliferation or motility, or both or neither, is likely due to the profile of S1P receptor expression. Thus, although all the glioma cell lines used expressed the three S1P receptors S1P1, S1P2, and S1P3, the relative levels of expression of these receptors varied. For instance, cell lines that do not respond mitogenically to S1P expressed extremely low levels of the receptor S1P1 (Van Brocklyn et al., 2003), suggesting that this receptor is crucial for mediating S1P‐stimulated glioma cell proliferation. In agreement, overexpression of S1P1 enables S1P to stimulate proliferation of glioma cells that normally are unresponsive, while siRNA‐mediated knockdown of S1P1 expression blocks S1P‐stimulated growth (Young and Van Brocklyn, 2007). Conversely, glioma cells in which S1P does not stimulate enhanced motility express high proportions of S1P2 (Van Brocklyn et al., 2003), a receptor which has previously been shown to block S1P‐induced cell motility (Okamoto et al., 2000). In agreement, Malchinkuu et al. (2005) showed that S1P inhibited migration of some glioma cell lines through S1P2 signaling. They also suggested that S1P2 is upregulated in astrocytoma cells in comparison to normal astrocytes based upon receptor expression in glioma cell lines and GBM tissue (Malchinkhuu et al., 2005). However, their analysis of GBM tissue utilized only two cases. We recently examined expression levels of S1P1, S1P2, and S1P3 by real time PCR analysis in 48 cases of GBM in comparison to 20 cases of the relatively benign pilocytic astrocytoma. We found no significant difference in expression of S1P1, S1P2, or S1P3 between these two groups (Van Brocklyn et al., 2005). Moreover, S1P2 expression in both tumor types was lower than that of S1P1, or S1P3 regardless of whether overall expression or expression as a percentage of the three receptors was examined. Thus, in individual glioma cases that express higher levels of S1P2, S1P would likely inhibit cell motility, however, this can not be generalized to astrocytomas or GBMs as there is no consistent change of S1P receptor expression in these tumors. Lepley et al. (2005) showed that migration of glioma cells can be stimulated or inhibited by S1P depending on receptor expression. Thus, cell lines that expressed high levels of S1P2, were inhibited from migration by S1P, while those expressing low levels were stimulated to migrate. Moreover, overexpression or knockdown of S1P receptor expression in these cells confirmed that the S1P2 receptor mediated inhibition of migration, while S1P1 mediated enhanced migration in response to S1P (Lepley et al., 2005). Interestingly, although S1P2 was originally shown to block migration through inhibition of the small GTPase Rac

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(Okamoto et al., 2000), S1P2 inhibition of glioma cell migration was not accompanied by a decrease in Rac activity (Lepley et al., 2005), and we have recently confirmed this finding (Young and Van Brocklyn, 2007). Thus, although S1P2 negatively regulates Rac activity in a variety of cell types, including Chinese Hamster Ovary (CHO) cells (Okamoto et al., 2000), smooth muscle cells (Ryu et al., 2002) and melanoma cells (Arikawa et al., 2003), S1P2‐mediated inhibition of glioma cell migration works by a different mechanism. This mechanism appears to involve Rho activation leading to stress fiber formation and, in some cells with excessive levels of Rho activity, cell rounding through activation of Rho kinase (Lepley et al., 2005). Interestingly, although, S1P2 decreased migration of glioma cells, overexpression of S1P2 in U‐118 MG cells enhances S1P‐stimulated invasion through Matrigel in both transwell and spheroid invasion assays (Young and Van Brocklyn, 2007). This unexpected enhancement of glioma cell invasion by S1P2 is mediated by increased cell adhesion to Matrigel and by induction of expression of the matricellular protein CCN1/Cyr61 (Young and Van Brocklyn, 2007). As mentioned above, S1P is also mitogenic for some glioma cell lines, showing a strong mitogenic response in some cell lines (Van Brocklyn et al., 2002) and a weak response in others (Lepley et al., 2005). Moreover, SphK, the enzyme which forms S1P, has been shown to be overexpressed in a variety of tumor types (French et al., 2003; Kawamori et al., 2005; Le Scolan et al., 2005). Therefore, we examined the role of SphK in glioma growth. High expression level of the SphK isoform SphK1 in GBM tissue correlates with shorter patient survival (Van Brocklyn et al., 2005). Thus, SphK1 expression could be useful as a prognostic indicator for GBM. Moreover, siRNA‐mediated knockdown of SphK1 expression decreased glioma cell proliferation by inhibiting entry into the cell cycle (Van Brocklyn et al., 2005). A pharmacological SphK inhibitor, 2‐(p‐hydroxyanilino)‐4‐(p‐chlorophenyl) thiazole, had a similar effect (Van Brocklyn et al., 2005). Interestingly, although SphK2 expression in GBM tissue did not correlate with patient survival, knockdown of SphK2 inhibited glioma cell proliferation even more potently than SphK1 knockdown did (Van Brocklyn et al., 2005). In agreement, the competitive SphK inhibitor dimethylsphingosine potently blocked proliferation of two glioma cell lines (Lepley et al., 2005). Another biological effect of S1P that has been extensively investigated is angiogenesis. S1P has been shown to induce an angiogenic response in a variety of systems (Lee et al., 1999a, b; English et al., 2000). In addition, a recent study showed that an antibody directed to S1P effectively blocked angiogenesis in a variety of mouse tumor models, although brain tumor models were not investigated (Visentin et al., 2006). Since angiogenesis is an important part of GBM pathogenesis it is tempting to speculate that S1P may be involved in mediating glioma angiogenesis. However, one report showed that, contrary to its stimulatory effect on most endothelial cells, S1P inhibited migration and tube formation by brain‐derived endothelial cells (Pilorget et al., 2005). Nevertheless, S1P could have indirect effects on angiogenesis within brain tumors through regulation of expression of other proangiogenic or antiangiogenic factors.

3.2 Glycosphingolipids Glycosphingolipids (GSL) consist of ceramide, to which is attached at the 1‐position, a variety of carbohydrate structures. Those containing sialic acid are known as gangliosides, while those without are neutral GSL. The commonly expressed ganglio‐series gangliosides contain a core structure of Galb1‐3GalNAcb1‐ 4Gal b1‐4Glc b1‐1Ceramide or a shortened version of this chain with sialic acids attached to the Gal residues. The nomenclature consists of G (for ganglio‐series), M, D, or T (for mono‐, di‐ ,or tri‐ sialic acid) and a number 1–3, referring to the number of carbohydrates present in the core structure according to the formula, 5  n (Svennerholm, 1980). Thus, for example, GM1 consists of ceramide linked to the entire 4 carbohydrate core structure shown above with a single sialic acid, while GD3 consists of ceramide linked to only the first two carbohydrates of the core but with two sialic acids. Other ganglioside series containing core structures with different sugars and linkages, such as the neolacto‐series, designated by the prefix L instead of G, and the lacto‐series, designated by isoL, also exist. It has been known for some time that the composition of cell surface glycolipids changes during neoplastic transformation (Hakomori, 1984; Feizi, 1985). Numerous studies on glycolipids in brain tumors have focused on gangliosides. The general trend indicates a decrease in the total amount of ganglioside

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compared to normal brain and a simplification of the type of gangliosides, i.e., decreased proportions of more complex, polysialylated gangliosides and increased proportions of simpler mono‐ and disialogangliosides (Kostic and Buchheit, 1970; Yates et al., 1979; Traylor and Hogan, 1980; Fredman et al., 1986; Shinoura et al., 1992). This trend increases with increasing histological tumor grade (Berra et al., 1985). In particular, GBM contains increased proportions of the disialoganglioside GD3 (Traylor and Hogan, 1980). GD3 levels increase with astrocytoma malignancy from grade 1 through grade 4 (Berra et al., 1985), with the proportion of GD3 increasing from 15% in grade 1 astrocytomas to 60% in GBM (Gaini et al., 1988). Decreased amounts and proportions of total gangliosides represented by the more complex gangliosides of the 1b pathway, i.e., GD1b, GT1b, and GQ1b, were found with increasing histologic grade of astrocytic tumors (Sung et al., 1994). These modifications of membrane glycolipids that gradually accompany the progressive increase in the malignancy of the brain tumors suggest that glycolipids could be used as tumor markers for brain tumor diagnosis and classification. Tumors with characteristics of neuronal differentiation showed a similar trend of simpler gangliosides in more undifferentiated tumors and more complex gangliosides in the more highly differentiated tumor types. The very undifferentiated primitive neural ectodermal tumors (PNETs) contained mostly simpler ganglio‐series gangliosides, whereas tumors with clear neuronal characteristics, such as gangliocytoma and neurocytoma contained a complex pattern of gangliosides, similar to normal brain tissue (Sung et al., 1994; Yates et al., 1999b). GM3 and GD3 are also the predominant gangliosides in pediatric PNET, ependymoma, neuroblastoma, dysembryoplastic neuroepithelial tumor (Pan et al., 2000), and meningioma (Davidsson et al., 1989). Beyond the changes in proportions of gangliosides, brain tumors often express gangliosides that are not normally found in adult brain tissue. The ganglioside 30 ‐isoLM1 was found to be present in 100% of grade 3 and 4 gliomas examined but only 25% of grade 2 gliomas (Fredman et al., 1988). The ganglioside 30 60 ‐isoLD1 was also present in 67% of gliomas (Wikstrand et al., 1993). These gangliosides are not expressed in normal adult brain, but only in fetal brain and reactive gliosis. Astrocytic tumors also contained the neolacto‐series gangliosides 30 ‐LM1 and 60 ‐LM1 (Sung et al., 1994). 30 ‐LM1 was also found in medulloblastomas, while 30 ‐isoLM1 was found in medulloblastomas showing astrocytic differentiation (Gottfries et al., 1990). Decreased percentages of 1b gangliosides and the presence of 60 ‐LM1 were also found with increasing histological grade of oligodendrogliomas (Sung et al., 1996). Other glycolipids could be used to differentiate between tumors of astrocytic origin versus tumors of oligodendroglial origin. The presence of asialo‐GM1 and the absence of paragloboside, a neutral GSL containing the 4‐carbohydate chain of the neolacto‐series, correlated with the diagnosis of oligodendroglioma. The reverse pattern correlated with astrocytomas (Yates et al., 1999a). A better correlation with oligogendroglioma was obtained by combining asialo‐GM1 and absence of paragloboside with the myelin galactolipid biosynthetic enzyme UDP‐galactose: ceramide galactosytransferase (Popko et al., 2002). It is interesting to note that, although the simple neutral GSL galactosylcerebroside is specific for oligodendrocytes in normal brain, it is commonly found in astrocytomas as well as oligodendrogliomas (Singh et al., 1994), and thus, would not be a useful marker for oligodendrocytic tumors. Gangliosides may also have prognostic value in gliomas. One study divided brain tumor patients into three groups, Group A whose tumors contained less than 30% 1b gangliosides and the ganglioside 60 ‐LM1, Group B whose tumors contained less than 30% 1b gangliosides but without 60 ‐LM1, and Group C containing more than 30% 1b gangliosides with or without 60 ‐LM1 (Sung et al., 1995). These groups correlated with patient survival with Group A having the shortest survival, and Group C the longest. This was even true when only tumors of WHO grade 4 were examined (Sung et al., 1995). In addition, patients with medulloblastoma whose 1b gangliosides made up over 15% of the total gangliosides survived longer that those with lower 1b ganglioside proportions (Yates et al., 1999b). Immunohistochemical staining for GD1b was also found to correlate with histological grade and patient survival for astrocytomas and oligodendrogliomas, suggesting that immunohistochemistry for this ganglioside could be a useful prognostic procedure (Comas et al., 1999). Another group divided glioma patients into three categories based upon profiles of gangliosides as well as neutral glycolipids (Jennemann et al., 1990). GSL‐Type I contained primarily the gangliosides GM3 and GD3 as well as mostly simple mono‐ and dihexaosyl‐ceramide neutral glycolipids, GSL‐Type II contained gangliosides GM2 and GD2 in addition to the other lipids contained in

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Type I, while GSL‐Type III had a glycolipid pattern similar to normal brain containing the complex gangliosides GD1a, GD1b, and GT1b. This study agreed with previous work showing that the simpler Type I glycolipid pattern tended to identify more malignant tumors, while the Type III pattern was found primarily in low grade, benign gliomas (Jennemann et al., 1990). The same group also showed that, in the few cases of anaplastic astrocytoma containing a GSL‐Type III pattern, patient survival was increased, suggesting that glycolipid patterns may have prognostic value within individual histological grades (Becker et al., 2000). The predominant gangliosides expressed in glioma cell lines, tend to be the simpler ganglio‐series gangliosides such as GM3 and GM2 (Yates et al., 1992; Fredman, 1994; Maeda et al., 1998). However, results from studies on gangliosides in brain tumor cell lines should be interpreted with caution, as in several cases, brain tumors cell lines grown in culture have been shown to have different ganglioside contents than xenografts grown in vivo or than the original tumors. The majority of primary human central nervous system neoplasms and xenografts derived from glioma cell lines express the lacto‐series gangliosides 30 ‐isoLM1 and 30 ,60 ‐isoLD1, whereas the majority of glioma cell lines did not express either 30 ‐isoLM1 or 30 ,60 ‐isoLD1 (Wikstrand et al., 1991). These gangliosides appeared in xenografts of cell line D‐54 MG which does not express the lacto series in vitro (Wikstrand et al., 1994). In fact, growing these cell lines as xenografts causes the domination of the lacto‐series (Fredman, 1994). Another cell line, U‐118 MG, does express 30 ‐LM1 in vitro as well as in vivo, however, its ganglioside pattern does change to include a higher proportion of complex gangliosides when grown as a xenograft (Fredman et al., 1990). Thus, it appears that in vivo environmental factors influence glioma ganglioside expression. These changes may result from altered glycosyltransferase expression, as GM2 synthase levels paralleled expression levels of GM2, being 20 times higher in vitro than in xenografts (Fredman et al., 1996). However, using a mouse model, Seyfried et al. (1996) suggested that differences between ganglioside profile of in vivo brain tumors versus that of glioma cells in culture is mainly due to the infiltration of host cells that contribute to the GSL composition of the solid tumors in vivo, and that these gangliosides are primarily derived from host macrophages (Seyfried et al., 1998). Similar changes occur in medulloblastoma cells grown in vitro versus in vivo (Gottfries et al., 1991), although in this case glycosyltransferases did not parallel the changes in the glycolipid pattern. Gangliosides may play several roles in regulating the biology of glioma cells. One possibility is that gangliosides may decrease tumor cell proliferation. Although both simple and complex gangliosides can have this effect, some studies show that more complex gangliosides are more potent in this effect than simpler gangliosides. This model would fit well with the observed decrease in total gangliosides and simplification of ganglioside structure seen in gliomas. In this regard, exogenously added gangliosides inhibit proliferation of several cell types including glioma cells (Bremer and Hakomori, 1982; Icard‐ Liepkalns et al., 1982; Bremer et al., 1984) and neuroblastoma cells (Carine and Schengrund, 1984). GM3 inhibits proliferation of primary cells derived from a variety of brain tumor types including GBM, ependymomas, mixed gliomas, astrocytomas, oligodendrogliomas, and gangliogliomas (Noll et al., 2001). GM3 also induced glioma cell apoptosis (Fujimoto et al., 2005) and enhanced survival of mice with an intracranial gliosarcoma xenograft (Noll et al., 2001). Complex gangliosides may play a role in cell division as b‐series gangliosides are decreased during M phase in a glioma cell line (Choi et al., 1997). Antibodies specific for GD3 blocked proliferation of glioma cell lines, while anti‐GM2 had no effect, although the cell lines used expressed at least as much GM2 as they did GD3 (Hedberg et al., 2000). In addition, overexpression of GD3 synthase in U‐1242 MG glioma cells, in order to increase endogenous expression of GD3, caused caspase‐dependent apoptosis (Saqr et al., 2006). Gangliosides likely affect cell proliferation by modifying mitogenic signal transduction. Exogenously added gangliosides inhibit the tyrosine phosphorylation of EGF receptor (Bremer et al., 1986) and platelet‐ derived growth factor (PDGF) receptor (Bremer et al., 1984). Gangliosides inhibit PDGF‐induced increased [Ca2þ]i, and mitogenesis without affecting the binding of PDGF to its receptors, and these effects are more potently induced by more complex gangliosides (Yates et al., 1993). Ganglioside inhibition of PDGF‐ induced signaling is mediated by the inhibition of PDGF receptor dimerization (Van Brocklyn et al., 1993) and recruitment of PDGF receptor to the glycolipid‐enriched microdomains (Mitsuda et al., 2002). Interestingly, different gangliosides can have different effects on various receptor tyrosine kinases.

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For example GM1, which inhibits PGDF receptor dimerization, enhances activity of the NGF receptor Trk (Rabin and Mocchetti, 1995) by stimulating Trk dimerization (Farooqui et al., 1997). Gangliosides can, however, also enhance cell growth. Gangliosides have a bimodal effect on glioma cell growth depending upon the growth state of cells when exposed to gangliosides. Thus, while GM1 treatment blocked growth of serum‐stimulated glioma cells, it stimulated growth of quiescent glioma cells (Saqr et al., 1995). The stimulatory effect of GM1 on cell growth was mediated by activation of several mitogenic signaling pathways including ERK MAP kinases, p70 S6 kinase (Van Brocklyn et al., 1997) and phosphatidylinositol (PI) 3‐kinase (Rampersaud et al., 1998). Thus, the effects of gangliosides on cell growth and mitogenic signaling are complex. Gangliosides may also influence invasiveness of glioma cells. Invasion of glioma cells in an in vitro assay was inhibited by either addition of exogenous gangliosides or by the ganglioside‐specific antibody A2B5 (Merzak et al., 1994). GM3 is also known to suppress the invasion of glioma cells in rat brain slice cultures (Fujimoto et al., 2005). On the other hand, GD3 expression on glioma cells was associated with a migratory phenotype (Gratsa et al., 1997), and transfection of GD3 synthase into rat C6 glioma cells caused these cells to produce faster growing tumors with more motile and invasive cells (Sottocornola et al., 1999). It is unclear how this reconciles with the stimulation of glioma cell apoptosis induced by GD3 synthase expression discussed above (Saqr et al., 2006), however it is possible that different levels of GD3 expression cause different biological responses. One possible mechanism for ganglioside modulation of cell invasion is through regulation of cell adhesion. Several gangliosides enhanced adhesion of glioma cell lines to fibronectin, collagen I, vitronectin, and laminin in vitro, suggesting modulation of integrin function. GD3 was the most effective adhesion‐ promoting ganglioside, while GM3 had little effect (Merzak et al., 1995). Ganglioside promotion of adhesion may be mediated by enhanced production of laminin (Koochekpour et al., 1995). Interestingly, gangliosides can also directly interact with integrins. GT1b binds to integrin a5b1 and blocks adhesion to fibronectin (Sung et al., 1998). Another possible mechanism for regulating invasion is through the regulation of metalloproteinases, which are responsible for extracellular matrix degradation during tumor cell migration. GM3 inhibits activation of matrix metalloproteinase (MMP)‐9 by blocking its interaction with integrins (Wang et al., 2003). On the other hand, exogenously added gangliosides upregulated the secretion of MMP‐2 and MMP‐9 in six of eight glioma‐derived cell lines investigated (Maidment et al., 1996). Thus, regulation of invasion by gangliosides is also complex and may involve different effects of various gangliosides on different aspects of this complex process. It should also be noted that, as discussed above, in vivo ganglioside expression is different that seen in glioma cell lines in culture. In this regard, in an invasive rat brain tumor model 30 ‐isoLM1, which, as noted above, is more commonly found in tumor cells grown as xenografts than in vitro, was expressed in the periphery of the tumor, associated with areas of tumor cell invasion, while GD3 was expressed throughout the tumor mass (Hedberg et al., 2001). Gangliosides may also regulate biology at a distance from the cells that produce them as they are shed from tumor cells (reviewed in Lauc and Heffer‐Lauc, 2006), and are thought to potentially modify immune reactions to tumors (Ladisch et al., 1983). Ganglioside shedding occurs in medulloblastoma (Chang et al., 1997) and neuroblastoma (Li and Ladisch, 1991) cells. Shedding of GD3 also occurs into CSF of astrocytoma patients (Ladisch et al., 1997), and increased levels of GD3 (Nakamura et al., 1991) and GM3 (Lo et al., 1980) were detected in serum of GBM patients, while GD2 was found in serum from neuroblastoma patients (Schulz et al., 1984). With regard to immune modulation GM2 and GM3 inhibit NK cell activity (Grayson and Ladisch, 1992). Moreover shed gangliosides induce T cell apoptosis by causing degradation of NF‐kB (Thornton et al., 2004). Furthermore, gangliosides GM2 and GD1a induced apoptosis of T‐cell lymphocytes cocultured with GBM lines (Chahlavi et al., 2005). Another important aspect of glioma pathology that gangliosides may influence is angiogenesis. Angiogenesis commonly occurs in high grade gliomas and microvascular endothelial proliferation is a diagnostic feature of GBM. Interestingly, immunostaining of GBM tissue for GD3 revealed that GD3 is highly expressed in hypervascularized areas of these tumors (Koochekpour and Pilkington, 1996). Angiogenesis in gliomas is known to be regulated by VEGF produced by cells near areas of necrosis (Fischer et al., 2005). Interestingly, GD3 potently stimulates release of VEGF from glioma cells (Koochekpour et al., 1996).

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Moreover, enhanced expression of complex gangliosides in murine ependymoblastoma cells transfected with N‐acetylgalactosaminyl transferase, a ganglioside biosynthetic enzyme, led to increased VEGF expression, increased vascularization and decreased necrosis in tumors formed by these cells in vivo (Manfredi et al., 1999). These tumors, which expressed the gangliosides GM2, GM1, and GD1a, also grew more rapidly than control tumors which expressed only GM3 (Manfredi et al., 1999). Finally gangliosides may be useful as targets for immunotherapy using specific monoclonal antibodies. Targeting gangliosides in tumor therapy is a promising immunotherapeutic strategy since gangliosides expression on the cell surface makes them accessible to ganglioside‐directed antibodies. Immunotherapy using monoclonal antiganglioside antibodies in their native form or combined with various effector molecules has been investigated as a therapeutic approach for several neural tumors. This field has been recently reviewed (Kurpad et al., 1995; Bitton et al., 2002; Raffaghello et al., 2003) and, therefore, will not be discussed in further detail here.

4

Glycerolipids

As noted above, early studies detected an increase in the amount of phosphatidylcholine in high grade gliomas. Several more recent studies have confirmed this observation. Phosphatidylcholine is present at a higher relative proportion in gliomas than in normal brain and correlates with low degree of tumor cell differentiation (Tugnoli et al., 2001). In addition, 31P‐MR spectroscopy showed higher levels of phosphatidylcholine in glioblastomas than in low grade astrocytomas (Lehnhardt et al., 2001). The increase in phosphatidylcholine appeared to be at the expense of sphingomyelin, and the authors suggested that this may reflect a shift toward generation of second messengers activating protein kinase C rather than ceramide from sphingomyelin which would likely inhibit growth (Lehnhardt et al., 2001). Lysophosphatidic acid (LPA) is a signaling lipid that has been implicated in glioma pathobiology. LPA binds to a group of four GPCRs that are closely related to the S1P receptors discussed above, called LPA1–4 (Ishii et al., 2004). LPA is thought to play a variety of roles in the development and function of the nervous system (reviewed in Chun, 1999; Steiner et al., 2002; Chun, 2005). LPA is present at micromolar concentrations in serum, and thus it has been speculated that blood–brain barrier breakdown or hemorrhage in gliomas could result in exposure of tumors to high LPA concentrations. LPA has been shown to regulate several signaling pathways in gliomas as well as to stimulate both glioma cell proliferation and migration. LPA causes release of Ca2þ from intracellular stores in rat C6 glioma cells (Hildebrandt and Hildebrandt, 1997). LPA also activates several other signaling mediators in C6 glioma cells including protein kinase C, ERK MAP kinase, PI‐3 kinase/Akt and cyclicAMP response element binding protein (CREB) and these pathways mediate a mitogenic effect of LPA on these cells (Cechin et al., 2005). In addition, LPA stimulates DNA synthesis in 1321N1 human astrocytoma cells through Gi signaling and G12/13 signaling to activate Rho GTPase (Seasholtz et al., 2004), and induces c‐fos expression in C6 glioma cells (Segura et al., 2005). Several studies have focused on the role of LPA in enhancing human glioma cell motility. LPA potently stimulates both chemotactic and chemokinetic migration of several human glioma cell lines by a signaling mechanism that required Gi protein signaling (Manning et al., 2000). Although LPA induced ERK activation and increased [Ca2þ]i, these pathways were not required for the motility response (Manning et al., 2000). LPA‐induced migration required coating of substrates with extracellular matrix proteins such as laminin and vitronectin, suggesting that LPA might enhance invasion of glioma cells along CNS blood vessels which are coated with these molecules (Manning et al., 2000). Recently LPA‐induced glioma cell migration was shown to also depend on Rac‐induced activation of Jun N‐terminal kinase (JNK) and cdc42‐ induced activation of p38 MAP kinases (Malchinkhuu et al., 2005). Indeed, LPA‐induced migration of U‐251 MG glioma cells is completely inhibited by siRNA knock down of Rac1 (Salhia et al., 2005). Additional evidence linking LPA to glioma cell invasion has come from recent work on the enzyme autotaxin (reviewed in Dennis et al., 2005). Autotaxin was originally identified as a nucleotide phosphodiesterase, expression of which is associated with invasive behavior of tumor cells (Murata et al., 1994).

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However, purification of a lysophospholipase D activity from serum, which cleaves lysophosphatidylcholine to form LPA, showed that this enzyme is identical to autotaxin (Tokumura et al., 2002; Umezu‐Goto et al., 2002). Evidence suggests that the invasion‐related function of autotaxin is due to its lysophospholipase D activity rather than its nucleotide phosphodiesterase activity (Koh et al., 2003). Autotaxin was found to be expressed specifically in invading glioma cells in comparison to core tumor cells by laser capture microdissection as well as by a glioma invasion tissue microarray (Hoelzinger et al., 2005). Moreover, the human glioma cell line U‐87 MG expresses very high levels of autotaxin and plasma from glioma patients contained significantly elevated levels of autotaxin/lysophospholipase D activity (Umezu‐ Goto et al., 2004). Recently it was shown that autotaxin stimulates migration of human GBM cell lines and that this effect is mediated by the LPA receptor LPA1 (Hama et al., 2004). In addition, GBM cell lines commonly overexpress autotaxin (Kishi et al., 2006). Thus, LPA produced by autotaxin may play a significant role in glioma invasion. Autotaxin has also been shown to be expressed in and regulate motility of neuroblastoma cells (Kawagoe et al., 1997; Dufner‐Beattie et al., 2001). Another glycerolipid‐based signaling pathway that is important in glioma growth is the phosphatidylinositol 3‐kinase (PI3 kinase)/Akt pathway, which can regulate cell growth, survival, and migration. Chromosomal changes in malignant gliomas are known to include loss of a portion or all of chromosome 10 (Ohgaki, 2005). A tumor suppressor gene located on chromosome 10 was discovered in 1997, and named phosphatase and tensin homolog deleted on chromosome 10 (PTEN) (Li et al., 1997). PTEN functions as a lipid phosphatase, dephosphorylating phosphatidylinositol at the 30 position, thus counteracting the effects of PI3 kinase (Stambolic et al., 1998). A large number of studies have shown that loss of PTEN in malignant gliomas allows for enhanced cell growth, survival, and possibly invasion and angiogenesis through enhanced signaling of the PI3 kinase/Akt pathway (reviewed in Knobbe et al., 2002). A high percentage of GBM tumors contain deletions or mutations of PTEN. In addition, many tumors which do not contain loss of PTEN have aberrations in other portions of the PI3 kinase pathway (Knobbe et al., 2002). Thus, the PTEN/PI3 kinase/Akt pathway represents a lipid‐based signaling pathway with immense potential for targeting in GBM as well as several other cancers. As this extensive field has been the subject of several recent reviews (Altomare and Testa, 2005; Hay, 2005; Cully et al., 2006) it will not be discussed in further detail here.

5

Sterols

As noted above, one of the earliest studies to examine lipids in brain tumors noted a high level of cholesteryl esters in brain tumors (Brante, 1949). More recent studies have confirmed that cholesteryl esters are elevated in brain tumors and areas surrounding brain tumors compared to normal brain (Nygren et al., 1997). Cholesteryl esters were also found to correlate positively with glioma histological grade (Tosi et al., 2003). Thus, it appears that cholesteryl esters are commonly associated with malignant gliomas. Furthermore, cholesteryl esters as measured by NMR in human gliomas correlated with the degree of vascular proliferation (Tugnoli et al., 2001). This may be due to the need of rapidly proliferating tumor cells or endothelial cells for cholesterol and either enhanced uptake of serum LDL or cholesteryl ester synthesis within tumors. However, not all cholesteryl esters are associated with enhanced brain tumor proliferation. Injection of liposomes containing 7‐b‐hydroxycholesteryl‐3‐oleate into C6 glioma tumors growing in rat brains significantly inhibited growth of the tumors, while liposomes not containing oxysterols had no effect (Werthle et al., 1994). The cytotoxic effect of this lipid may be partially due to inhibition of ERK MAP kinase (Adamczyk et al., 1998). Thus, the effects of these lipids on gliomas may be complex and involve both direct effects on tumor cells and effects on vascular growth. Nygren et al. (1997) suggested that cholesteryl esters in brain tumors were serum‐derived based on the fatty acid composition more closely resembling that of serum cholesteryl esters than those of normal brain. However, whereas patients with a variety of peripheral tumors display hypocholesterolemia, those with either primary or metastatic brain tumors do not, possibly suggesting a lack of reliance of brain tumors on

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serum cholesterol (Grieb et al., 1999). Moreover, no significant relationship was detected between mortality for brain tumor patients and serum cholesterol level (Smith et al., 1992). In addition, one study even found that increased dietary fat and cholesterol may be associated with decreased risk of gliomas (Kaplan et al., 1997). Statins are drugs which inhibit 3‐hydroxy‐3‐methylglutaryl‐coenzyme A the rate limiting step in the mevalonate pathway leading to cholesterol synthesis. As cholesterol synthesis is thought to be necessary for cell proliferation numerous studies have examined the effects of statins on a variety of tumors including brain tumors (reviewed in Jakobisiak and Golab, 2003). Statins inhibit proliferation and induce apoptosis of C6 glioma cells via JNK (Koyuturk et al., 2004). Simvastatin inhibits growth of several human glioma cells in vitro and rat and mouse models (Kikuchi et al., 1997; Murakami et al., 2001). Moreover, simvastatin synergistically potentiated the antiproliferative effects of (2‐chloroethyl)‐N‐nitrosourea or b‐interferon on human glioma cell growth in vitro (Soma et al., 1992) and on C6 glioma growth in rats (Soma et al., 1995). Lovastatin inhibits growth of human glioma cell lines and primary cells cultured from GBMs (Bouterfa et al., 2000). Lovastatin also inhibits neuroblastoma growth in mice, although this was not accompanied by a decrease in serum cholesterol (Maltese et al., 1985) suggesting an alternate mechanism of action. The effects of statins may not be due entirely to reduction in cholesterol needed for membrane biosynthesis. The mevalonate pathway also leads to production of farnesyl pyrophosphate and geranylgeranyl pyrophosphate which are needed for prenylation of ras family small GTPases. Inhibition of GBM cell growth by lovastatin was associated with decreased Ras‐MAP kinase signaling (Bouterfa et al., 2000). In addition, interference with activity of Rho proteins by prevention of prenylation has also been implicated in statin‐induced increased expression of cyclin‐dependent kinase (CDK) inhibitors (Jakobisiak and Golab, 2003). Indeed, induction of apoptosis of medulloblastoma cells by statins may be mediated by induction of CDK inhibitors (Wang and Macaulay, 2003). A few clinical trials with Statins have been run, with moderately encouraging results. One minor response was found in a patient with recurrent malignant glioma in lovastatin Phase I trial (Thibault et al., 1996). A phase I‐II trial was conducted in which Lovastatin was given alone or in combination with radiation (Larner et al., 1998). Lovastatin alone produced one minor response and one partial response from nine patients, while in combination with radiotherapy, Lovastatin produced two minor responses and two partial responses (Larner et al., 1998). In addition, in a Phase I trial of Fluvastatin in pediatric cancer patients, two of six patients with anaplastic astrocytoma showed a partial response (Lopez‐Aguilar et al., 1999). Sterols might also play important roles in mitogenic signaling in some brain tumors. The Sonic Hedgehog (SHH) pathway has been implicated in the formation of medulloblastomas. Recent evidence indicates that inhibitors of sterol synthesis block SHH‐dependent gene expression and proliferation in medulloblastoma cells, and this can be reversed by addition of cholesterol or oxysterols (Corcoran and Scott, 2006).

6

Conclusions and Future Directions

Significant gains have been made in understanding the changes in lipid profiles that accompany oncogenic transformation within neural tissue as well as the molecular significance of the lipids present in these tumors. Clearly, however, many cases exist in which certain lipids play complex roles. It is likely that several lipids can affect a variety of cellular regulatory pathways, possibly depending upon exact expression level and/or subcellular localizations. Undoubtedly, further work in these fields will elucidate how targeting pathways regulated by lipids in these tumors, probably in combination with other therapies, can be used to eventually develop effective treatments for these malignancies. It is interesting to note that in recent years the field of neuro‐oncology has seen a major advancement in the discovery that malignant brain tumors, particularly GBM and medulloblastoma, contain tumor stem cells (reviewed in Sanai et al., 2005; Vescovi et al., 2006). These brain tumor stem cells appear to be the only cells within the tumors which are capable of reforming the tumor when transplanted due to their property of self‐renewal, and thus are likely to be the most important cells in which to understand molecular abnormalities and to target therapeutically. Furthermore, brain

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tumor stem cells have proven to be more appropriate models for study than are traditional cell lines, as they form tumors in animals closely resembling the original tumors from which they were isolated, a property which the traditional cell lines lack. Thus, it will be interesting to determine whether the alterations in lipids and roles that lipids play in these tumors apply to brain tumor stem cells, and whether lipid pathways can be effectively targeted in these cells.

Acknowledgments The author thanks Dr. Hany Saqr for helpful recommendations regarding glycosphingolipids. Work in the author’s laboratory was supported by Grant # R01 NS41517 from the National Institute of Neurological Disorders and Stroke (NINDS).

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D. A. Butterfield . H. M. Abdul

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 564

2 2.1 2.2 2.3 2.3.1 2.3.2 2.3.3

Lipids in Plasma Membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 565 Types of Lipids in Plasma Membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 565 Synthesis and Transport of Phospholipids to Plasma Membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 566 Identification of Phospholipid Transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 566 Aminophospholipid Translocase (Flippase) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 567 Multidrug Resistance Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 568 Scramblase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 568

3 3.1 3.2 3.3 3.4

Lipid Alterations in AD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 569 Altered Composition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 569 Formation of Reactive Aldehydes: HNE and Acrolein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 571 TBA Reactive Substances . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 573 Isoprostanes and Neuroprostanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 573

4 4.1

Modulation in Phospholipid Asymmetry in AD by Ab (1–42) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 575 In Vitro and In Vivo Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 575

5

Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 576

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_22, # Springer ScienceþBusiness Media, LLC 2009

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Abstract: Lipids are important biological molecules. The lipids of physiological importance for humans have four major functions: (1) structural components of biological membranes; (2) energy reserves, predominantly in the form of triacylglycerols; (3) both lipids and lipid derivatives serve as vitamins and hormones, and (4) lipophilic bile acids aid in lipid solubilization. Fatty acids fill two major roles in the body: as the components of more complex membrane lipids and as the major components of stored fat in the form of triacylglycerols. Alterations in lipid structure and/or metabolism lead to many neurodegenerative diseases, among which Alzheimer’s disease (AD) is of great concern due to the increasing life-span of the world’s population. Additionally, altered cholesterol metabolism, modulation in phospholipid content, and phospholipid asymmetry in plasma membranes may play pivotal role in the progression of AD. Amyloid b-peptide [Ab (1–42)] plays a central role in the pathogenesis of AD. Ab (1–42) is heavily deposited in the brains of Alzheimer’s disease (AD) patients, and free radical oxidative stress of neuronal lipids is extensive. Research by our group and others suggests that this observation is linked to Ab-induced oxidative stress in AD brain. This chapter summarizes current knowledge on lipid alterations in AD brain, one potential cause of the external oxidative stress in AD brain, and the consequences of Ab-induced lipid peroxidation in this neurodegenerative disorder. List of Abbreviations: AD, Alzheimer’s disease; Aβ(1-42), amyloid beta-peptide; PtdSer, Phosphatidylserine; PtdEtn, Phosphatidylethanolamine; PtdIns, phosphatidylinositol; PtdCho, phosphatidylcholine; GEM, glycosphingolipid-enriched membranes; RBC, red blood cells; i-NOS, inducible nitric oxide synthase; GPC, glycerophosphatidylcholine; GPE, glycerophosphatidylethanolamine; sPLA2, Ca2+ -dependent secretory phospholipase A2; cPLA2, Ca2+ -dependent cytosolic phospholipase A2; iPLA2, Ca2+ -independent phospholipase A2; HNE, 4-hydroxy-2-nonenal; DAG, diacylglycerol; NCT, nicastrin; βaPP, β-amyloid precursor protein; TBARS, thiobarbituric acid reactive substances; IsoPs, isoprostanes; NPs, neuroprostanes; PG, prostaglandin; LV, levuglandins; NeuroKs, neuroketals; NFT, neurofibrillary tangles; SP, senile plaques; FAEE, ferulic acid ethyl ester; MCL, mild cognitive impairment; ROS, reactive oxygen species; RNS, reactive nitrogen species

1

Introduction

Alzheimer’s disease (AD) is a progressive dementing disorder characterized clinically by a loss of memory and cognition, and later aphasia (Katzman and Saitoh, 1991). Pathologically AD is characterized by senile plaques (composed of aggregated amyloid b-peptide [Ab]) and dystrophic neurites, neurofibrillary tangles (composed of hyperphosphorylation of tau), and synapse loss (Katzman and Saitoh, 1991). The AD brain is under oxidative stress, including lipid peroxidation (Butterfield and Lauderback, 2002). Ab can induce lipid peroxidation (Butterfield et al., 1994; Mark et al., 1997; Lauderback et al., 2001). Lipids are amphipathic molecules. Phospholipids derive from glycerol esters of two fatty acids and phosphoric acid. The latter moiety is esterified to an alcohol that gives the phospholipid its name. With numerous species of lipids expressed in eukaryotic plasma membranes and cells have the task of organizing the lateral and transverse distribution of membrane phospholipids to specific sites. Cells have developed a number of mechanisms to deal with this issue, one of which is to create a dynamic equilibrium to their site of function that ensures interaction of specific lipids with appropriate partner moieties. The targeting of phospholipids to specific membrane sites is essential for maintaining cell shape, homeostasis, and signal transduction cascades. Aminophospholipids also have been implicated in a diverse array of processes ranging from catabolism to inflammation and cell proliferation to cell death (> Table 22-1). The components of biological membranes are asymmetrically distributed between the membrane surfaces, and lipids are asymmetrical as they are present on both sides of the bilayer, but in different and highly variable amounts. Asymmetry is maintained by the action of a protein that requires ATP known as flippase. The cholinecontaining phospholipids (phosphatidylcholine and sphingomyelin) are found on the external surface of the plasma membrane, while the aminophospholipids (phosphatidylserine [PtdSer] and phosphatidylethanolamine [PtdEtn) are localized primarily on the cytoplasmic lamellae (Kurz et al., 2005). ATP-dependent processes (Pomorski et al., 2001) maintain phospholipid asymmetry, which is critical to normal cell function. An alteration in phospholipid distribution, particularly the appearance of PtdSer on the extracellular surface,

Lipids in Alzheimer’s disease brain

22

. Table 22-1 Arrangement of aminophospholipids in outer leaflet of cells Cell type Lipid Non-pathological cells PC-12 cells PtdEtn Macrophages PtdSer Mast cells Pathologic cells Apoptotic cells Ab-treated cells Carcinoma Diabetes

PtdSer PtdSer PtdSer PtdSer PtdSer, PtdEtn

Remarks

References

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Fadok et al. (1992) Mohmmad Abdul and Butterfield (2005) Rao et al. (1992) Wali et al. (1988)

Martin et al. (2000)

occurs during apoptosis and contributes to the recognition and destruction of these cells by macrophages (Kirschnek et al., 2005).

2

Lipids in Plasma Membranes

2.1 Types of Lipids in Plasma Membranes Plasma membranes contain a wide diversity of lipids, all of which are amphipathic. There are three main types of membrane lipids: phosphoglycerides, sphingolipids, and cholesterol. Most membrane lipids contain a phosphate group, which makes them phospholipids, and as they are built on glycerol backbone, they are called phosphoglycerides. Membrane phosphoglycerides have an additional group linked to the phosphate, most commonly choline (forming phosphatidylcholine, PtdCho), ethanolamine (forming phosphatidylethanolamine, PtdEtn), serine (forming phosphatidylserine, PtdSer), or inositol (forming phosphatidylinostol, PtdIns). Each of these groups is small and hydrophilic and together with the negatively charged phosphate to which they are attached, forms a, highly water-soluble domain at one end of the molecule, called the head group. At physiological pH, the head groups of PtdSer and PtdIns have an overall negative charge, whereas those of PtdCho and PtdEtn are zwitterions. Phosphoglycerides most often contain one saturated (a-chain) and one unsaturated fatty acyl chain (b-chain). A less abundant class of membrane lipids, which are the derivatives of sphingosine, an amino alcohol that contains a long hydrocarbon chain, is called sphingolipids. Sphingolipids consist of sphingosine linked to a fatty acid by its amino group, called ceramide. The various sphingosine-based lipids have additional groups esterified to the terminal alcohol of the sphingosine moiety. If the substitution is phosphorylcholine, the molecule is sphingomyelin. If the substitution is a carbohydrate, the molecule is a glycolipid. If the carbohydrate is a simple sugar, the glycolipid is called cerebroside; if the carbohydrate is an oligosaccharide, the glycolipid is called a ganglioside. All the sphingolipids are amphipathic and basically similar in overall structure to the phosphoglycerides. Another lipid component of certain membranes is the sterol cholesterol, which in certain animal cells may constitute up to 50% of the lipid molecules in the plasma membrane. Cholesterol is smaller than the other lipids of the membrane and less amphipathic. Cholesterol leads to increased lipid fluidity in membranes in which unsaturated lipids occur. Cholesterol is asymmetrically distributed in the membrane (Wood et al., 1990). Lipid rafts are specialized membrane domains enriched in certain lipids, cholesterol, and proteins (Simons and Ikonen, 1997). Caveolae are flask-shaped invaginations on the cell surface that are a type of

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membrane raft; these moieties were named ‘‘caveolae intracellulare’’ (Yamada, 1955). It presently seems that there could be three types of lipid rafts: caveolae; glycosphingolipid-enriched membranes (GEM); and polyphosphoinositol-rich rafts. It may also be that there are inside rafts (PIP2 rich and caveolae) and outside rafts (GEM). Lipid rafts are presumed to be signaling centers, perhaps regions of cholesterol import (Huang et al., 1999) and involved in endocytosis (Thomsen et al., 2002).

2.2 Synthesis and Transport of Phospholipids to Plasma Membrane Assembly of the phospholipid bilayer of cellular membranes is a fundamental aspect of cell growth and proliferation. Phospholipids are concomitantly synthesized and inserted at the cytoplasmic surface of the endoplasmic reticulum. Following this asymmetric assembly, transmembrane movement to the lumenal leaflet of the endoplasmic reticulum must occur in order to ensure coordinated growth of the bilayer. For phosphatidylcholine, the predominant phospholipid of eukaryotic membranes, the latter process appears to be facilitated by a specific transport protein. There are several pathways for the synthesis of aminophospholipids in eukaryotic cells (Vance and Shiao, 1996; Voelker, 2000). Phosphatidylserine (PtdSer) is synthesized by a Ca2+-dependent PtdSer synthase-catalyzed reaction in which serine is exchanged with the head group of phosphatidylcholine (PtdCho) or phosphatidylethanolamine (PtdEtn) (Kuge et al., 1986). This occurs mainly in the endoplasmic reticulum (ER) and in specialized ER-derived mitochondrial-associated membranes [MAM] (Vance and Vance, 1988) that bridge the ER to the mitochondria. The newly synthesized lipid is transported from the MAM to the mitochondrial outer membrane by an ATPdependent mechanism (Voelker, 1989). The lipid then moves to the inner membrane where it serves as a substrate for PS decarboxylase I to generate PE. The Golgi and vacuoles also synthesize PE by decarboxylating PS with decarboxylase II. Both vesicular and cytosolic protein-mediated transfer mechanisms are involved in the transport of aminophospholipids from their sites of synthesis to the plasma membrane (Sprong et al., 2001). The delivery of lipids from extracellular sources can also alter the transverse distribution of phospholipids. HDL and LDL act as lipid transport proteins in plasma to deliver phospholipids, cholesterol, and fatty acids to the cell’s outer membrane leaflet where they are utilized by the cell. Additionally, endo- and exocytosis, which involve multiple membrane fusion events, may induce lipid intermixing between membrane leaflets. In order to maintain an appropriate asymmetric aminophospholipid distribution, cells have developed several mechanisms. Lipid asymmetry is generated primarily by selective synthesis of lipids on one side of the membrane. Because passive lipid transbilayer diffusion is slow, a number of proteins have evolved to maintain this lipid gradient. These proteins fall into three classes: (1) ‘‘flippases’’ are cytofacially-directed, ATP-dependent transporters (Seigneuret and Devaux, 1984); (2) ‘‘floppases’’ are exofaciallydirected, ATP-dependent transporters (Connor et al., 1992); and (3) ‘‘scramblases’’ are bidirectional, ATPindependent transporters (Williamson et al., 1985). The flippase is highly selective for phosphatidylserine and functions to keep this lipid sequestered crossing from the external cell surface. Floppase activity has been associated with the ABC class of transmembrane transporters. Although they are primarily nonspecific, at least two members of this class display selectivity for their substrate lipid. Scramblases are inherently nonspecific and function to randomize the distribution of newly synthesized lipids in the endoplasmic reticulum or plasma membrane lipids in activated cells. It is the combined action of these proteins and the physical properties of the membrane bilayer that generate and maintain transbilayer lipid asymmetry.

2.3 Identification of Phospholipid Transporters Once lipid asymmetry has been established, it is maintained by a combination of slow transbilayer diffusion, protein–lipid interactions, and protein-mediated transport. The thermodynamic barrier to passive lipid flip-flop prevents rapid spontaneous transbilayer diffusion of phospholipids. The half time for phospholipid flip-flop is approximately long and depends on the nature of the lipid and the membrane. In the human erythrocyte, flip-flop rates are dependent on the phospholipid acyl chain length and the

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degree of unsaturation (Middelkoop et al., 1986). Although membrane lipid asymmetry has been known for many years, the mechanisms for maintaining or regulating the transbilayer lipid distribution are still not completely understood. Perhaps the most significant contributors to the maintenance of transbilayer lipid asymmetry are proteins that catalyze the movement of lipids across the membrane (> Table 22-2). These activities are classified according to the substrate specificity, requirements for energy, and direction of transport. Most of the pivotal studies of membrane phospholipid asymmetry have been performed in human red blood cells (RBC). Although these cells have provided a wealth of information on the biochemical properties of aminophospholipid transporters, further progress has been impeded by the inability to identify the active protein components in these cells by molecular biology techniques. The ultimate transbilayer distribution of lipids is determined by the specificity of the lipid transporters located in the membrane. Each of the transport activities described in a following section displays a unique specificity or non-specificity that defines its function in the determination of lipid organization. The following summarizes the current state of knowledge regarding the specificity of these transport activities.

2.3.1 Aminophospholipid Translocase (Flippase) Flippase is widely distributed and is present in most plasma membranes. Aminophospholipid flippase activity is expressed in erythrocytes and has been detected in a wide variety of cell types and membranes, including platelets, lymphocytes, spermatozoa, and synaptosomes (Zachowski and Gaudry-Talarmain, 1990; Muller et al., 1994). It is likely that this transporter is essential for any membrane in which the maintenance of PtdSer asymmetry is required. The aminophospholipid flippase is perhaps the most selective of the lipid transporters. This protein prefers PtdSer to other lipids (Daleke and Huestis, 1985), and the specificity for PtdSer is defined by each of the functional groups of the lipid. Sphingolipids (Morrot et al., 1989) and diether analogs of PtdSer are also recognized as transport substrates, but transport rates are reduced compared with diacylglycerophosphoserine. The glycerol backbone is another important recognition element. There is some flexibility in lipid backbone recognition by the enzyme. However, the enzyme displays an absolute requirement for the stereochemistry of the glycerol backbone; the sn-2,3-glycerol analog of the naturally occurring sn-1,2-glycero-lipid is not a substrate for transport (Martin and Pagano, 1987). In contrast to the polar headgroup specificity, the flippase is capable of accepting PtdSer molecules containing fatty acids of various lengths and composition, including those modified by spin, fluorescent, and photoactivatable groups (Colleau et al., 1991; Demaurex et al., 1997). This activity is associated with an ATP-dependent rapid movement (within seconds to several minutes) of both PtdSer and PtdEtn from the outer to inner membrane leaflet in virtually all eukaryotic cells. Based on its ATP dependence and sensitivity to fluoride and vanadate, Seigneuret and Devaux (Seigneuret and Devaux, 1984) postulated the activity to be associated with an aminophospholipid-specific transport protein. Thus, the aminophospholipid transporter requires the participation of a Mg2+-ATPase, whereas the varied and sometimes conflicting results raise the possibility that several distinct proteins cooperate to form a functional aminophospholipid transport complex. . Table 22-2 Lipid transporters Class Flippases

Floppases

Scramblases

Protein Erythrocyte Mg2+-ATPase ABCR P4-ATPases ABCA1 ABCB4

Specificity PS PE Amphipaths Cholesterol PC

ABCC1 ER flippase

Short-chain phospholipids None

References Daleke and Lyles (2000) Weng et al. (1999) Halleck et al. (1998) Rust et al. (1999) van Helvoort et al. (1996) Kamp and Haest (1998) Buton et al. (1996)

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ATPase II is another flippase that has been purified and cloned from bovine chromaffin granules (Moriyama and Nelson, 1988), the close homologs of which have been identified from bovine brain and human sources. These proteins are members of a new class of ATPases, the P4-ATPases (Tang et al., 1996). Defects in genes of this family produce alterations in ribosomal assembly, and familial intrahepatic cholestasis. Like the erythrocyte Mg2+-ATPase, the ATPase activity of these enzymes is selectively activated by PtdSer (Moriyama et al., 1991). Although no direct evidence for transbilayer phospholipid transport has been reported, it is likely that the P4-ATPases are involved either directly or indirectly in amphipath transport.

2.3.2 Multidrug Resistance Proteins The second class of ATP-dependent lipid transporters is the exofacially-directed floppases. Early studies in red blood cells revealed a nonspecific outward flux pathway for NBD [1-palmitoyl-2-[6(7-nitrobenz-2-oxa-1, 3-diazol-4-yl)amino]caproyl-sn-glycero-3-phosphoserine] and spin-labeled lipids (Connor et al., 1992). It was recognized subsequently that some members of the ABC transporter superfamily are also capable of transporting lipids (Borst et al., 2000). ABC transporters are a diverse group of proteins that, in general, are responsible for the ATP-dependent export of amphipathic compounds. These include the multidrug resistance proteins, which export cytotoxic xenobiotics and were first discovered in drug-resistant tumor cells. Consistent with their role in general xenobiotic amphipath export, ABC proteins are, for the most part, nonspecific. However, some members of this class demonstrate a unique specificity for their respective substrate. The most well-characterized lipid floppase activities are those catalyzed by ABCA1, ABCB1, ABCB4, and ABCC1. They are broadly classified into P-glycoprotein (MDR), multidrug resistance-associated protein (MRP), and mitoxantrone resistance protein (MXR) by their size and the number of transmembrane loops (Gottesman and Pastan, 1993). The substrates for MDR and MXR are largely hydrophobic or amphiphilic, whereas those recognized by MRP are principally anionic and likely require cotransport of glutathione disulfide (GSSG) (and/or GSSX) to be operative. Experiments using fluorescent and spinlabeled phospholipid analogs provided evidence for the existence of an ATP- and protein-dependent nonspecific floppase that transports lipid from the inner to outer membrane leaflet (Connor et al., 1992). More recent studies, however, indicate that transport is a result of MRP1 activity that expels lipids out of the cells, because their fluorescent and spin-labeled reporter tags imbued the lipid with drug-like properties (Kamp and Haest, 1998). Although these results seem to eliminate a role for the purported nonspecific floppases in the externalization of authentic phospholipids, several lipid-specific floppases have been identified that include human MDR3 (Ruetz and Gros, 1994) and ABC1 (Williamson et al., 1992), which function as true PtdCho- and PtdSer-specific floppases, respectively. Taken together, these results suggest that the steady-state equilibrium distribution of membrane phospholipids is maintained by joint activities of an aminophospholipid-specific flippase and cholinephospholipid-specific floppase (MDR family) and that acquisition of the PS-expressing apoptotic phenotype requires activation of ABC1 coincident with Ca2+-mediated inhibition of the aminophospholipid translocase (Williamson et al., 1992). It is interesting to note that not all ABC lipid transporters are floppases. ABCR is another ABC protein with lipid transport activity although it is a flippase, rather than floppase.

2.3.3 Scramblase Studies with platelet and red cell membranes revealed the existence of a Ca2+-dependent scrambling activity that seemed to result in complete intermixing of lipids between bilayer leaflets irrespective of headgroup specificity (Hamon et al., 2000). Three scramblase activities have been reported; two are involved in dissipating lipid gradients in biological membranes and the third is activated by Ca2+ in the plasma membrane of stimulated cells. The ER scramblase is known to be relatively nonspecific and was first described as a bidirectional transporter of PC and its metabolites (Chapman and Trelease, 1991). Evidence has not been found for the activity of these transporters in the plasma membrane. Thus, they may serve only to redistribute newly synthesized lipids or lipid precursors in ER and Golgi membranes. Albeit less extensive

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than for platelets, lipid scrambling has also been demonstrated in a variety of other cells such as lymphocytes, endothelial cells, red blood cells, smooth muscle cells and tumorigenic cells. An increase in the intracellular calcium concentration, for instance, evoked by cellular activation, complement pore formation, or induction of apoptosis, is an essential requisite for the onset of the scrambling process. Increased intracellular Ca2+ can also involve activity of aminophospholipid translocase activity, indicating that the loss of lipid asymmetry does not result from mere inhibition of aminophospholipid translocase activity (Kuypers et al., 1996). Scrambling requires the continuous presence of Ca2+. Removal of intracellular calcium causes an arrest of the scrambling process and may even restore aminophospholipid translocase activity, provided that its irreversible degradation by calpain is prevented. The scrambling process is bidirectional and involves all major phospholipid classes, which move at comparable rates except for sphingomyelin for which the inward movement tends to be somewhat less in comparison with the glycerophospholipids. Using fluorescent probes, it was demonstrated that lipids with unusual polar headgroups, such as the disomer form of serine (Smeets et al., 1994), or glucosyl- and galactosyl ceramides participate in the scrambling process. Lipid scrambling is not coupled to ATP hydrolysis and even occurs in resealed erythrocyte ghosts when challenged with Ca2+ (Williamson et al., 1985). Lipid scrambling is sensitive to the sulfhydryl modifications (Williamson et al., 1985).

3

Lipid Alterations in AD

3.1 Altered Composition The pathogenic mechanisms in AD have been hypothesized to involve alterations in the concentrations of phospholipids (Hazel and Williams, 1990; Prasad et al., 1998). This hypothesis is supported by the presence of high concentrations of phospholipids in the brain that contain highly oxidizable polyunsaturated free fatty acids, such as arachidonic and docosohexanoic acids (Dhillon et al., 1994), and that Ab, implicated in the pathogenesis of AD (Selkoe, 2001), prefers to localize in the phospholipid membrane core (Mason et al., 1996; Mark et al., 1997; Lauderback et al., 2001). The loss of membrane phospholipids may be an early metabolic event in the formation of SP and NFT (Pettegrew et al., 1988) and in the loss of synapses and neurons (Svennerholm and Gottfries, 1994). Because phospholipids contain unsaturated fatty acids, which are targets of free radicals, this hypothesis is linked to the oxidative stress hypothesis of neurodegeneration in AD (Markesbery and Lovell, 1998; Butterfield, 2002). In the central nervous system (CNS), the loss of membrane phospholipids occurs by at least three different mechanisms: decreased biosynthesis, increased degradation, and increased lipid peroxidation. Experimental evidence in AD suggests that phospholipase-catalyzed degradation of phospholipids is likely to decrease the concentrations of membrane phospholipids in the brain in AD (Barany et al., 1985). Increased free radical-mediated lipid peroxidation occurs in the brain in AD (Lyras et al., 1997; Markesbery and Lovell, 1998; Lauderback et al., 2001; Montine et al., 2002; Pratico and Sung, 2004; Sultana and Butterfield, 2004) and can contribute to decreases in the concentrations of phospholipids in this disorder. Furthermore, phospholipases may enhance phospholipid degradation after free radical attack, forming lipid peroxides and lipid hydroperoxides (Hall et al., 1994; Francescangeli et al., 2000, 2002; Sun et al., 2001). Studies showing the loss of membrane phospholipids have demonstrated either decreased levels of lipid phosphorus (Nitsch et al., 1992; Svennerholm and Gottfries, 1994) or increased levels of the phospholipid catabolites, glycerophosphatidylcholine (GPC) and glycerophosphatidylethanolamine (GPE) in the AD brain (Blusztajn et al., 1990). Ceramide is not only structurally, but also functionally, a key molecule in diverse kinds of sphingolipids (Sawai et al., 2005). In the past decade, ceramide has been shown to be of crucial significance in several cell functions including apoptosis, cell growth, senescence, and cell cycle control (Flores et al., 2000; Thon et al., 2005). Among these roles, the role of ceramide in apoptosis induction has extensively been studied, and ceramide-targeting molecular medicine for apoptosis-based diseases such as malignant tumors, atherosclerosis and neurodegenerative disorders is under extensive investigation (Mattson et al., 1997; Sawai et al., 2005). The recent advances in the research on ceramide-mediated apoptosis signaling show the relation of

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ceramide level through regulation of ceramide-related enzymes with diseases such as cancer, leukemia, bacterial infections, AIDS, Alzheimer’s disease, atherosclerosis, diabetes mellitus, and atopic dermatitis (Sawai et al., 2005). Amyloid beta-peptide enhances tumor necrosis factor-alpha-induced iNOS through the neutral sphingomyelinase/ceramide pathway in oligodendrocytes (Zeng et al., 2005). Cholesterol may play a pivotal role on the production of the putative AD neurotoxin, Ab (Sjogren et al., 2005). More importantly, this relationship has consistently been identified in both in vivo and in vitro studies (Sparks et al., 2003). Cholesterol-lowering drugs have been shown to cause a beneficial effect of lowering Ab levels in animal models, and epidemiological data indicate a beneficial effect on the risk of AD with prior statin use (Sparks et al., 2003). However, the results using statins may not be simply due to cholesterol lowering, but may involve other statin-sensitive pathways (Laufs et al., 2002; Mohmmad Abdul et al., 2004). Increased risk of developing late-onset AD is related to the apolipoprotein E gene found on chromosome 19 (Poirier, 2005). This gene codes for a protein that helps to carry cholesterol in the bloodstream. The APOE gene comes in several different forms, or alleles, but three occur most commonly: APOE e2, APOE e3, and APOE e4. Having one or two copies of the e4 allele increases the risk of developing AD. The e3 allele is the most common form found in the general population and may play a neutral role in AD. The e2 allele appears to be associated with a lower risk of AD. The exact degree of risk of AD for any given person cannot be determined based on APOE status. Therefore, the APOE e4 gene is called a risk factor gene for late-onset AD (Craig et al., 2004), and, consistent with the importance of lipids in AD, there are increased levels of CSF phosphorylated tau in ApoE4 carriers with mild cognitive impairment, arguably the earliest form of AD (Buerger et al., 2005). Numerous signal transduction processes involve lipids as signaling molecules. Many of these molecules are generated by phospholipases such as phospholipase A2 that releases fatty acids such as arachidonic acid and lysophospholipids. Each of these products is implicated in signal transduction processes by itself (Farooqui and Horrocks, 2005), but also serves as a precursor for eicosanoids including the prostaglandins, leukotrienes, and lipoxins or platelet-activating factor (PAF). Phospholipase A2 is a member of the class of heat-stable, calcium-dependent enzymes catalyzing the hydrolysis of the 2-acyl bond of 3-n-phosphoglycerides. The enzyme has a molecular weight of 30,000 Da. Phospholipase A2 is activated by Ca2+ and inhibited by zinc, barium, and manganese ions (Kabre et al., 1999). There are more than 19 different isoforms of PLA2 in the mammalian system, but recent studies have focused on three major groups, namely Ca2+-dependent secretory enzymes (sPLA2), the Ca2+-dependent cytosolic enzyme (cPLA2), and a Ca2+-independent PLA2 (iPLA2) (Balsinde and Dennis, 1997). There is accumulating evidence for the involvement of specific PLA2s in AD brain pathology and neurodegeneration (Macchioni et al., 2004). In two separate studies, a decrease in PLA2 activity was found in the parietal and temporal cortex (Ross et al., 1998), as well as the prefrontal cortex, of the AD brain (Talbot et al., 2000). On the other hand, immunohistochemical studies showed an increase in cPLA2 immunoreactivity associated with the glial fibrillary acidic protein-positive astrocytes in the AD brain (Stephenson et al., 1996). In a recent gene array study, profiling of 12,633 genes in the hippocampal CA1 area of AD patients indicated an increase in cPLA2 and COX-2 expression, as well as upregulation of a number of apoptotic and proinflammatory genes (Colangelo et al., 2002). Additionally, cytosolic phospholipase A2 mediates neuronal apoptosis induced by soluble oligomers of the Ab peptide (Sun et al., 2001; Kriem et al., 2005), and the brain and platelet PLA2 activity was significantly lower in AD than in controls. Mean PLA2 activity in MCI individuals was between the values of AD patients and controls, with a subgroup showing PLA as low as the lowest AD patients, but the differences from MCI were not significant from AD and control groups (Gattaz et al., 2004). These findings are in agreement with the increased oxidative and inflammatory responses and decrease in PLA2 activity associated with AD pathology. Other important phospholipases include phospholipase C, which controls the production of inositol-1,4,5trisphosphate (IP3), which, in turn, induces cytosolic Ca2+ release and diacylglycerol (DAG). The latter moiety activates protein kinase C. Phospholipase D generates phosphatidic acid (PA), which subsequently can be metabolized either by PLA2 generating lysophosphophatidic acid (Lyso PA), a potent cellular mitogen, or by phosphatidate phosphohydrolase (PAP) yielding DAG. Sphingomyelinase, a phospholipase C type enzyme, and related enzymes of sphingolipid metabolism are implicated in apoptosis and other signaling processes (Mattson et al., 1997).

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Several lines of evidence suggest the role of Ab (1–42)-induced lipid peroxidation in AD. D609, a tricyclodecanol derivative of xanthic acid, scavenges hydroxyl radicals and reacts with electrophilic products of lipid oxidation (HNE and acrolein) in a manner similar to GSH (Lauderback et al., 2003). Synaptosomes isolated from rodents, previously injected intraperitoneally with D609, followed by treatments with the oxidants Fe2+/H2O2, 2,2-azobis-(2-amidinopropane) dihydrochloride (AAPH), showed a significant reduction in 4-hydroxy-2-nonenal (HNE) [a lipid peroxidation product] (Joshi et al., 2005). Other studies suggest that D609 significantly attenuated Ab (1–42)-induced cytotoxicity and lipid peroxidation in vitro (Sultana et al., 2004) and in vivo (Joshi et al., 2005; Perluigi et al., 2006). The antioxidant property of D609 is associated with the free thiol group of xanthate. D609 is capable of detoxifying aldehydic products of lipid peroxidation by a mechanism similar to GSH (Lauderback et al., 2003). Lipid rafts are membrane microdomains enriched in cholesterol and sphingolipids, and proteins (Simons and Ikonen, 1997). Several lines of evidence suggest the involvement of lipid rafts in b- and g-cleavage of bAPP (Ehehalt et al., 2003). It has been reported that the proteins relevant to Ab generation, including presenilin, nicastrin, and a small portion of bAPP, localize in lipid rafts (Lee et al., 1998). Additionally, b-secretase localizes in lipid rafts and cholesterol depletion abrogates this localization (Riddell et al., 2001). Researchers (Ehehalt et al., 2003) have reported that bAPP exists in two pools; one associated with lipid rafts, in which b-cleavage occurs, and the other outside of lipid rafts, where a-cleavage occurs. The gamma-secretase complex is responsible for the final cleavage event in the processing of beta-amyloid precursor protein (bAPP), resulting in Ab generation. The gamma-secretase complex is a multiprotein complex composed of presenilin, nicastrin (NCT), APH-1, and PEN-2. Recent reports have suggested that g-secretase activity is predominantly localized in lipid rafts and that cholesterol can directly regulate the g-secretase activity in isolated lipid rafts (Wahrle et al., 2002). Presenilin and NCT have been reported to be localized in lipid rafts. Thus, lipid rafts offer a structural platform for examining the effect of cholesterol on Ab generation. Compositional alterations in brain phospholipids, due in part to lipid peroxidation, have been reported for AD brain (Prasad et al., 1998). Polyunsaturated fatty acids (PUFA), including arachidonic (AA), and docosohexanoic acid (DCH), are abundant in brain and highly oxidizable. Consequently, AA and DCH are vulnerable to free radical attack, and PUFA are predicted to decrease in AD brain if lipid peroxidation were increased. In an analysis, regional levels of membrane phospholipids PtdEtn, PtdIns, and PtdCho were measured in the brain of AD and control subjects and the results suggest that the levels of PtdEtn-derived and PtdIns-derived total fatty acids were significantly decreased in the hippocampus of AD subjects (Prasad et al., 1998). Additionally, significant decreases were found in PtdEtn-derived stearic, oleic and arachidonic and DCH, and in PtdIns-derived oleic and arachidonic acids (Prasad et al., 1998). In the inferior parietal lobule of AD subjects, significant decreases were found only in PtdEtn and those decreases were contributed by stearic, oleic and arachidonic acids. In the superior and middle temporal gyri and the cerebellum of AD subjects, no significant decreases were found in PtdCho-, PtdEtn- , and PtdIns-derived fatty acids. The decrease of PtdEtn and PtdIns, which are rich in oxidizable AA and DCH, but not of PtdCho, which contains lesser amounts of these fatty acids, suggests a role for oxidative stress in the increased degradation of brain phospholipids in AD (Prasad et al., 1998). Analyses using high-performance liquid chromatography (HPLC) and gas chromatography (GC) also describe a decrease in PtdEtn-plasmalogen in AD brain, confirming PtdEtn as a phospholipid exhibiting major structural modifications (Guan et al., 1999). One mechanism for decreased free fatty acids is oxidative stress-induced stimulation of phospholipase A2, and Ab added to synaptosomes led to free fatty acid release, primarily in the PtdEtn fraction, an effect blocked by the free radical scavenger vitamin E (Koppal et al., 1998).

3.2 Formation of Reactive Aldehydes: HNE and Acrolein Free radical attack on phospholipid PUFA can ultimately lead to multiple aldehydes with different carbon chain lengths, including acrolein and HNE (Esterbauer et al., 1991). These alkenals have different reactivity than free radicals due to their half-life ranging from minutes to hours. In particular, HNE is able to diffuse to sites distant from that of its formation (Butterfield and Stadtman, 1997; Butterfield, 1997).

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HNE, an a,b-unsaturated aldehyde, is one of the major products of lipid peroxidation. HNE reacts with proteins, forming stable covalent adducts to histidine, lysine, and cysteine residues through Michael addition, thereby introducing carbonyl functionalities into proteins following oxidative damage, e.g., lipid peroxidation (Esterbauer et al., 1991; Uchida and Stadtman, 1992; Subramaniam et al., 1997; Butterfield and Lauderback, 2002). HNE can inhibit synthesis of DNA, RNA, and proteins and alter activity of glycolytic, degradative, and transport proteins (Esterbauer et al., 1991). One mechanism for increasing HNE in AD may be the oxidative stress associated with Ab (Varadarajan et al., 2000; Lauderback et al., 2001). Ab is widely reported to cause lipid peroxidation in brain cell membranes, which is inhibited by free radical antioxidants (Butterfield et al., 1994; Avdulov et al., 1997; Gridley et al., 1997; Mark et al., 1997, 1999; Bruce-Keller et al., 1998; Daniels et al., 1998; Yatin et al., 2000; Boyd-Kimball et al., 2004). Further, Ab leads to HNE and 2-propen-1-al (acrolein) formation (Mark et al., 1997), and these alkenals alter the conformation of membrane proteins (Subramaniam et al., 1997; Pocernich et al., 2001). Moreover, these reactive aldehydes are toxic to neurons (Mark et al., 1997; Subramaniam et al., 1997; Lovell et al., 2001). Indeed, an increase in free- and protein-bound HNE was found in rat hippocampal neurons or synaptosomes exposed to Ab (Mark et al., 1997; Lauderback et al., 2001; Boyd-Kimball et al., 2004; Sultana et al., 2005). HNE induced apoptosis in PC12 cells or neurons (Kruman et al., 1997) suggesting that Ab-induced HNE production may contribute to an indirect mechanism of neuronal death. The concentration of free HNE is elevated in multiple brain regions and in ventricular cerebrospinal fluid (CSF) in AD (Markesbery and Lovell, 1998). Protein-bound HNE also is elevated in AD (Sayre et al., 1997; Lauderback et al., 2001; Sultana and Butterfield, 2004), and may relate to apolipoprotein E allele (Montine et al., 1997). As already sited, ApoE e4 allele is a risk factor for AD (Craig et al., 2004), suggesting that the degree of expression the lipid transporter ApoE in brain might be associated with HNE production in AD. Stereotaxic injection of HNE in rat forebrain selectively inhibited cholineacetyltransferase (ChAT) (Bruce-Keller et al., 1998), the activity of which is greatly diminished in AD. Consistent with this observation, synaptosomes treated with Ab (1–42) result in HNE being bound to ChAT (Butterfield and Lauderback, 2002), which may possibly correlate with the diminished ChAT activity in AD. HNE also may play a role in glutamate-induced neurotoxicity in AD. Glutamate, an excitotoxin that exerts its effects via stimulation of N-methyl-D-aspartate (NMDA) receptors, thereby increasing intracellular free radicals (Lafon-Cazal et al., 1993), is removed from the synapse by glutatmate transporters, particularly, the glial glutamate transporter, GLT-1 [also called as EAAT2] (Maragakis and Rothstein, 2001). Ab inhibits glutamate uptake (Harris et al., 1995; Harris et al., 1996) possibly by a mechanism that involves Ab-induced lipid peroxidation and subsequent HNE modification to glutamate transporters (Keller et al., 1997; Lauderback et al., 2001). In AD brain GLT-1 has significantly more HNE bound to it than does this transporter in aged-matched neurologically normal controls (Lauderback et al., 2001), and full-length Ab added to rodent synaptosomes induced HNE binding to GLT-1 (Lauderback et al., 2001), suggesting the possibility that these two observations are related. The activity of glutathione S-transferases (GST), an enzyme that shows high catalytic (and thus detoxifying) activity against HNE (Bruns et al., 1999), was significantly decreased in AD brain (Lovell et al., 1998). Taken together with the increased levels of lipid peroxidation in AD brain, these findings may in part account for neurodegeneration in AD brain. Additionally, the alpha class of glutathione S-transferase (GST) can detoxify HNE and plays an important role in cellular protection against oxidative stress. GST and MRP1 are themselves oxidatively modified by HNE in AD brain (Sultana and Butterfield, 2004), which, since oxidatively modified proteins are generally dysfunctional (Butterfield and Boyd-Kimball, 2004), may account in part for the elevated HNE accumulation in this disorder (Markesbery and Lovell, 1998). Protein-bound HNE is also elevated in brain from persons with MCI, suggesting that lipid peroxidation is an early event in the progression of AD (Butterfield et al., 2006). Acrolein is the most reactive of the a,b-unsaturated aldehydes produced by lipid peroxidation because of its electrophilic properties (Esterbauer et al., 1991). Even at low concentrations acrolein can structurally change transmembrane and cytoskeletal proteins (Pocernich et al., 2001). Acrolein adducts were reported in NFT in AD but not in control brain (Calingasan et al., 1999), and an increase in protein-bound acrolein in AD compared with age-matched controls was described (Lovell et al., 2001). Further, acrolein is toxic to primary hippocampal cultures (Lovell et al., 2001), and acrolein is reportedly bound in excess to

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alpha-ketoglutarate dehydrogenase in AD brain (Calingasan et al., 1999). This may suggest that acrolein can disrupt mitochondrial function (Pocernich and Butterfield, 2003).

3.3 TBA Reactive Substances One measure of lipid peroxidation is elevation of thiobarbituric acid reactive substances (TBARS). Unfortunately, reaction of nonlipid molecules with thiobarbituric acid makes TBARS a nonspecific marker of membrane lipid peroxidation, possibly accounting for disagreements in reports about TBARS levels in different brain regions in AD. An increase in TBARS in AD frontal lobe but not in the cerebellum was reported (Subbarao et al., 1990), while others (Balazs and Leon, 1994) described a significant TBARS increase in sensory and occipital cortices of AD. Only the inferior parietal lobe seemed to be affected by lipoxidation in one study (Palmer and Burns, 1994), while others (Lovell et al., 1995) showed statistically significant elevations in TBARS in the hippocampus and the cortex. A TBARS increase in all the regions of AD brain, with higher statistical significance in the temporal cortex, was described (Marcus et al., 1998):, a result confirmed by others (Tamaoka et al., 2000). One mechanism of increasing lipid peroxidation measured by TBARS is the addition of a free radical initiating source, and Ab (1–40) was shown to increase TBARS in synaptosomal membranes (Lauderback et al., 2001).

3.4 Isoprostanes and Neuroprostanes Isoprostanes (IsoPs) are prostaglandin (PG)-like compounds that are formed nonenzymatically in vivo by free radical-induced peroxidation of arachidonic acid. IsoP formation proceeds through bicyclic endoperoxide PGH2-like intermediates. The endo-peroxide intermediates are reduced to form PGF2-like compounds (F2-IsoPs) (Morrow et al., 1990) or undergo rearrangement to form E-ring and D-ring compounds (E2/D2-IsoPs) (Morrow et al., 1994) and thromboxane-like compounds [isothromboxanes] (Morrow et al., 1996). An increase in CSF levels of the isoprostane 8,12-iso-iPF2alpha-VI, a specific marker of in vivo lipid peroxidation, was demonstrated for CSF in AD patients (Pratico et al., 2004). Poor cerebral clearance of end products of oxidative reactions via CSF circulation may contribute to and sustain ongoing stress. CSF drainage via a low-flow ventriculoperitoneal shunt may improve removal of these products, reducing oxidative stress (Pratico and Sung, 2004). A novel aspect of the formation of IsoPs is that, unlike cyclooxygenase-derived prostaglandins, IsoPs are formed in situ esterified to phospholipids and subsequently released (Morrow et al., 1992). The quantification of F2-IsoPs has emerged as one of the most accurate approaches to assessing oxidant injury in vivo (Roberts and Morrow, 1997). Furthermore, IsoPs are capable of exerting potent biological activity (Roberts and Morrow, 1997). Isoketals (IsoKs) are highly reactive products of the isoprostane pathway of free radical-induced lipid peroxidation that rapidly form covalent protein adducts and form protein cross links in vitro. Isoketal adducts from an animal model of oxidative injury revealed that initial adducts were formed by isoketals esterified in phospholipids, representing a novel oxidative injury-associated modification of proteins by phospholipids (Brame et al., 2004). Studies on potassium channels revealed that isoketal adduction profoundly altered protein function, inhibiting potassium current in a dose-dependent manner (Brame et al., 2004). This result suggests that phospholipid-esterified isoketals rapidly adduct membrane proteins and that such modification can alter protein function, suggesting a generalized cellular mechanism for alteration of membrane function as a consequence of oxidative stress (Brame et al., 2004). Neuroketals are the highly reactive g-ketoaldehydes formed by the neuroprostane pathway of free radical-induced lipid peroxidation that is analogous to cyclooxygenase-derived levuglandins (LGs). Neuroketals (NeuroKs) are formed from the oxidation of DCH, which is enriched in the brain, and measurement of neuroprostanes may provide a unique marker of oxidative neuronal injury (Roberts and Morrow, 2002). Adduction of IsoK to model proteasome substrates significantly reduced their rate of degradation by the 20S proteosome (Davies et al., 2002). At lower concentrations, an IsoK-adducted protein (ovalbumin)

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and peptide (Ab1–40) significantly inhibited chymotrypsin-like activity of the 20S proteosome (Davies et al., 2002). Additionally, incubation of IsoK with P19 neuroglial cultures dose-dependently inhibited proteasome activity and induced cell death suggesting that IsoKs/NeuroKs can inhibit proteasome activity and may have relevance to the pathogenesis of neurodegenerative diseases, if overproduced (Davies et al., 2002). As noted, free radical-induced oxidation of AA results in the formation of F2- and D2/E2- isoprostanes; in contrast, analogous oxidation of gray matter-resident DCH leads to F4- and D4/E4-isoprostanes [neuroprostanes (NPs)] (Roberts et al., 1998). The level of IP, the quantitation of which directly relates to lipid peroxidation in vivo (Morrow, 2000), is increased in AD CSF (Montine et al., 1998). Consistent with Ab-associated free radical-induced lipid peroxidation (Butterfield et al., 1994; Butterfield and Lauderback, 2002), IP is significantly elevated in rat hippocampal culture after Ab addition (Mark et al., 1999). The levels of F2-IP in AD lateral ventricular fluid also were significantly elevated, and the increase was related with the extent of degeneration but independent of the distribution of NFT or the ApoE genotype (Montine et al., 1999). AD brain IP and NP levels in vivo were quantified, showing an increase in total NP level, but not total IP (Reich et al., 2001). In aggregate, these results suggest that lipid peroxidation, resulting in neurotoxic reactive aldehydes, may be important in the neurodegeneration observed in AD brain. Further, the extensive lipid peroxidation in AD brain suggests that brain-accessible, lipid-resident antioxidants that can block free radical-induced lipid peroxidation, or raising the in vivo level of glutathione, that is able to protect neurons from HNE and acrolein (Subramaniam et al., 1997; Pocernich et al., 2001) may be promising therapeutic strategies for this disorder (Drake et al., 2002). Quantification of NPs might provide a unique marker of oxidative injury in the brain. Furthermore, these compounds, like IsoPs, could potentially exert biological activity. This possibility is supported by the finding that PGF4a, the four series F-prostaglandin corresponding to the structure expected from cyclooxygenase action on C22:6, is approximately equipotent with cyclooxygenase-derived PGF2a in contracting gerbil colonic smooth muscle strips (Markesbery, 1997). In addition, the formation of NPs esterified in lipids might be expected to have significant effects on the biophysical properties of neuronal membranes, which might impair normal neuronal function. This may be particularly relevant to AD, since it has been suggested that one of the physiological functions of DCH may be to maintain a certain state of membrane fluidity and promote interactions with membrane proteins that are optimum for neuronal function (Salem and Niebylski, 1995). Neuroprostanes are readily detected esterified to lipids in the brain. However, NP measurements in humans would be limited to samples of brain removed surgically or postmortem samples of human brain. Measurements of F4-NPs made in human brain samples obtained after death could be quite problematic because of the possibility of artifactual generation of NPs by autoxidation of DCH during the time interval between death and sample procurement. Although invasive, cerebrospinal fluid is frequently obtained for diagnostic purposes in patients with suspected neurological disorders. Thus, the availability of a marker of oxidative injury in the brain that could be measured in CSF intra vitam would be an important advance. Hence, the finding that F4-NPs could be detected in human CSF clearly has potentially important clinical applications. Markers of lipid peroxidation are increased in CSF of patients with AD (Lovell et al., 1997; Montine et al., 1997). However, these assays have shortcomings related to measurement of reactive molecules, i.e., 4-hydroxynonenal, and require large volumes of fluid. However, F4-NPs were detectable using negative ion chemical ionization mass spectrometry in 1–2 ml of CSF from normal subjects, an amount that can usually be obtained safely from patients for diagnostic purposes. Although it was a limited study, the finding that F4-NP concentrations in cerebrospinal fluid from patients with AD were significantly higher than levels in age-matched control subjects highlights the potential of this approach to provide insights into the role of free radicals in the pathogenesis of neurological disorders. Another potentially important aspect of this finding is that serial measurements of F4-NPs in CSF might provide a biochemical assessment of disease progression as well as a means to monitor efficacy of therapeutic intervention, e.g., with antioxidants, during life. Elevation of tissue and urinary isoprostanes is characteristic of human atherosclerosis. Deficiency in apolipoprotein E in the mouse (apoE / ) resulted in atherogenesis and an increase in iPF2alpha-VI, an F2-isoprostane, in urine, plasma, and vascular tissue (Pratico et al., 1998). Supplementation with vitamin E significantly reduced isoprostane generation, but had no effect on plasma cholesterol levels in apoE / mice (Pratico et al., 1998).

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Therefore, there may be distinct advantages associated with measuring either IsoPs or NPs to assess oxidant injury in the brain. NPs possess important biological actions that may be relevant to the pathophysiology of oxidant injury to the brain. As mentioned, this possibility is greatly supported by the finding that C22-PGF4a is bioactive. This compound is one of the F4-NPs that would be formed, although, analogous to IsoPs, compounds in which the side chains are oriented cis likely predominate over compounds in which the side chains are oriented trans in relation to the cyclopentane ring (Morrow et al., 1990). However, in the case of the IsoPs, inversion of the stereochemistry of the upper side chain of PGF2a and PGE2 affords different and/or more potent biological actions (Roberts and Morrow, 1997). In addition, phospholipids containing esterified NPs are unnatural and unusual molecules. Thus, enhanced formation of these unusual phospholipids in neuronal membranes in settings of oxidant injury to the brain might lead to profound alterations in the biophysical properties of the membrane, e.g., degree of fluidity, which in turn might greatly impair normal neuronal function. Future studies using synthetic NP-containing aminophospholipids in model membranes to assess the extent to which these unique phospholipids alter membrane properties should provide valuable insight into the potential relevance of the formation of these phospholipids in settings of oxidative neuronal injury.

4

Modulation in Phospholipid Asymmetry in AD by Ab (1–42)

The maintenance of membrane lipid asymmetry is a dynamic process that influences many events over the lifespan of the cell. As already described, most cells restrict the bulk of the aminophospholipids to the inner membrane leaflet by means of specific transporters. For the major glycerophospholipids (PtdSer, PtdEtn, PtdCho, and PtdIns), de novo synthesis occurs on the cytosolic side of the endoplasmic reticulum [ER] (Bell et al., 1981). With the exception of PtdCho, this places the newly synthesized lipids on the side of the membrane in which they are ultimately enriched in the plasma membrane. Because of the thermodynamic barrier to spontaneous transbilayer movements, these lipids should remain enriched on the cytoplasmic side of the membrane, provided that there is no perturbation to the membrane. However, the asymmetric addition of newly synthesized phospholipids to one leaflet of the bilayer generates an unstable membrane. Accumulation of lipid on one side of the membrane can induce membrane bending and consequent membrane shape changes (Daleke and Huestis, 1985). The asymmetric distribution of PS over the cellular membrane requires much of the cell’s energy and requires the involvement of an ATP-dependent enzymatic activity (flippase) for its maintenance (Dolis et al., 1997). Indeed, if the cell fails to engage mechanisms to maintain asymmetry, aminophospholipids appear at the external cell surface. One of the important consequences of altered membrane asymmetry is the recognition and engulfment of phosphatidylserine bearing vesicles and cells by mononuclear phagocytes (Schroit et al., 1985). Loss of phospholipid asymmetry, measured by the exposure of PtdSer on the outer leaflet of the membrane bilayer, is a typical early event that follows apoptotic insult (Fadok et al., 1992). A large body of evidence indicates that lipid peroxidation is directly responsible for the generation of the apoptotic phenotype (Kagan et al., 2000).

4.1 In Vitro and In Vivo Studies An aminophospholipid, PtdSer is sequestered in the inner leaflet of the plasma membrane in nonstimulated cells. An early signal of synaptosomal apoptosis is the loss of phospholipid asymmetry and the appearance of phosphatidylserine in the outer leaflet of the membrane (Balasubramanian and Schroit, 2003). The inactivation of the transmembrane enzyme aminophospholipid-translocase (or flippase) by Ab (1–42) has been investigated (Mohmmad Abdul and Butterfield, 2005). Flippase activity depends on a critical cysteine residue, a putative site of covalent modification by the Ab (1–42)-induced lipid peroxidation products, HNE or acrolein. Pretreatment of synaptosomes with D609 (already discussed) and ferulic acid ethyl ester (FAEE), a potent antioxidant (Sultana et al., 2005) significantly protected against Ab (1–42)-induced loss of phospholipid asymmetry in synaptosomal membranes (Mohmmad Abdul and Butterfield, 2005). Annexin

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V binding and NBD-PS [1-palmitoyl-2-[6(7-nitrobenz-2-oxa-1,3-diazol-4-yl)amino]caproyl-sn-glycero-3phosphoserine] binding assays were used to study the externalization of PS. The results suggest that D609 and FAEE exert protective effects against Ab (1–42)-induced apoptosis in synaptosomal membranes (Mohmmad Abdul and Butterfield, 2005). As already noted, Ab interaction with membrane produces HNE (Markesbery, 1997; Lauderback et al., 2001). Additionally, HNE, too, was reported to lead to loss of phospholipid asymmetry in synaptosomes (Castegna et al., 2004). In vivo studies carried out at our lab recently with purified synaptosomes from AD and mild cognitive impairment (MCI) brain samples suggest a significant increase the modulation in phospholipid asymmetry (externalization of PS) compared with the aged matched controls (Bader Lange et al., 2008), consonant with the notion that oxidative stressmediated lipid peroxidation and subsequent apoptosis are early events in the progression of AD.

5

Concluding Remarks

The AD brain is under intense oxidative stress. Ab plays a central role in the pathogenesis of AD. Soluble, oligomeric Ab is postulated to insert into neuronal membranes, where, in processes that are inhibited by the chain-breaking antioxidant vitamin E and other antioxidants, Ab-induced lipid peroxidation, protein oxidation, ROS and reactive nitrogen species (RNS) formation occur (Butterfield, 1997; Varadarajan et al., 2000; Butterfield, 2002; Butterfield and Boyd-Kimball, 2004). This oxidative damage or membrane modification, resulting from the reaction of the lipid peroxidation products HNE and/or acrolein with enzymatic, transport, or structural proteins, alters synaptic membranes and leads eventually to the death of the neuron. HNE can diffuse from the site of its production, potentially modifying neuronal organelles and changing their function (Butterfield and Stadtman, 1997). Ab inhibits the function of several neuronal and glial transmembrane transport systems, including ion-motive ATPases, glutamate transporters, the glucose transporter, guanosine triphosphate (GTP)-coupled transmembrane signaling proteins, MRP-1, and the polyamine transporter (Butterfield and Lauderback, 2002). Additionally, Ab plays a pivotal role in phospholipid asymmetry in AD (both in vitro and in vivo) and compositional alterations in brain phospholipids. Each of these has functional consequences that are deleterious to neurons, such as loss of cell potential to accumulation of excitotoxic glutamate, decreased glucose availability, decreased intracellular communication, and increased neurotoxicity. Genetic mutations and other mechanisms (e.g., apolipoprotein genotype, redox metal ions, etc.) that potentially lead to an increased Ab deposition may contribute to this Ab-induced lipid peroxidation and neurotoxicity. Continued investigation of the role of Ab in oxidative stress in lipid peroxidation in AD and animal models relevant to AD, as well as studies employing therapeutic agents to block Ab-associated oxidative stress, cholesterol lowering drugs, and association of the b-secretase/active g-secretase complex with lipid rafts and its relation to cholesterol metabolism should provide greater insight into the relationships among Ab (1–42), lipid peroxidation, oxidative stress, and neurodegeneration in AD brain.

Acknowledgment This work was supported, in part, by NIH (NIA) grants to D. A. B.

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Neural Lipids in Parkinson’s Disease

M. Barichella . G. Pezzoli . A. Mauri . C. Savardi

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 584

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Toxic Agents and Oxidative Damage in PD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 584 Role of Toxic Agents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 585 Reactive Species and Oxidative Damage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 586 Oxidation Products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 587

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Lipid Influence on a‐Synuclein Behavior in PD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 587

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The Role of Cannabinoids in PD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 589

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_23, # Springer ScienceþBusiness Media, LLC 2009

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Abstract: A huge scientific effort is still ongoing in order to address the main etiologic and pathologic mechanisms responsible for the onset and the progression of Parkinson’s Disease, one of the most common chronic progressive neurological disorders. The key role of oxidative damage in PD has been widely studied. Exposition to different toxic agents can promote oxidative damages, for instance, via inhibition of mitochondrial functions. This can lead to the formation of dangerous reactive radical compounds and of their relative oxidation products which have been frequently discovered to be present in elevated concentration in the Substantia Nigra pars compacta neurons of PD affected patients. In particular, being cellular membranes a preferential site for lipid oxidation, of increasing importance are also the studies investigating the contribution of membrane lipids composition to the aggregation of the a-Synuclein protein in filamentous amyloid-like fibrils, giving origin to insoluble intracellular inclusions with the capacity to cause neuronal damage. More recently, the role of the Cannabinoid-based compounds on the endocannabinoid receptors is under investigation due to the suggestions of a close interaction of the dopaminergic transmission with the endocannabinoid system which could ultimately behave as a compensatory mechanism in case the functionality of previous one is compromised. These compounds could thus be considered as a potential therapy to alleviate motor symptoms and L-dopa associated dyskinesias in PD affected patients. List of Abbreviations: aS, a‐Synuclein; ATP, adenosine triphosphate; CB1, type 1 cannabinoid receptors; CB2, type 2 cannabinoid receptors; CNS, central nervous system; ETC, electron transport chain; GSH, glutathione; GSSG, glutathione disulfide; H2O2, hydrogen peroxide; HNE, 4‐hydroxynonenal; LB, Lewy bodies; LN, Lewy neurites; MAO, enzyme monoamine oxidase; MPPþ, 1‐methyl‐4‐phenylpyridinium; MPTP, 1‐methyl‐4‐phenil‐1,2,3,4‐tetrahydropiridine; NO, nitric oxide; NOS, nitric oxide synthase; OH, hydroxyl free radical; PD, Parkinson’s Disease; RNS, reactive nitrogen species; ROS, reactive oxygen species; Snca, human a‐Synuclein gene; SNpc, substantia nigra pars compacta; SOD, superoxide dismutase

1

Introduction

Parkinson’s Disease (PD) is one of the most common chronic progressive neurological disorders, affecting 3% of the population over the age of 65 (Lang and Lozano, 1998) with some indication of increased prevalence in recent times. PD is clinically characterized by motor symptoms, such as bradykinesia, rigidity, tremor, and postural imbalance. Subtle cognitive dysfunctions and/or depression are often present in the disease, whereas in its advanced stages dementia is not infrequent. PD is morphologically characterized by progressive and selective loss of neurons in the substantia nigra pars compacta (SNpc) (> Figure 23-1) and other subcortical nuclei associated with intracytoplasmatic filamentous aggregates called Lewy bodies (LB) and dystrophic (Lewy) neurites, (LN) mainly in subcortical nuclei and hippocampus (Jellinger, 2002). LB and LN serve as neuropathological hallmark of the disease (Forno, 1996; Giasson et al., 2000). The core biochemical pathology in parkinsonism decreased dopaminergic neurotransmission in the basal ganglia. In particular, degeneration of the nigrostriatal dopamine system results in marked loss of striatal dopamine content. Physiologically, the decreased dopaminergic activity in the striatum leads to disinhibition of the subthalamic nucleus and the medial globus pallidus, which is the predominant efferent nucleus in the basal ganglia (Fahn and Przedborski, 2005). The cause of PD in the vast majority of patients remains unknown. Research has concentrated on exogenous toxins, endogenous toxins from cellular oxidative reactions and genetics. Nevertheless it is generally believed that the environmental factors have a primary role in the pathogenesis of PD. In the following paragraphs, a more detailed biochemical description of the disease is presented, focusing the attention on the main etiologic and pathologic mechanisms responsible for disease onset and progression.

2

Toxic Agents and Oxidative Damage in PD

Among the different external and endogenous factors contributing to the onset of PD, oxidative stress contributes to the cascade leading to dopamine cell degeneration. However, oxidative stress is intimately

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. Figure 23-1 Cut section of the midbrain (left image) where a portion of substantia nigra is visible for a normal subject (upper right image) and for a Parkinson’s disease affected patient (lower right image)

linked to other components of the degenerative process, such as mitochodrial dysfunction, exo/endotoxicity, and inflammation. It is therefore difficult to determine whether oxidative stress leads to, or is a consequence of, these events.

2.1 Role of Toxic Agents The discovery that the neurotoxin 1‐methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine (MPTP) induces neuropathological and neurochemical alterations as well as clinical signs very similar to those of PD, suggesting that a similar chemical compound may cause PD (Jenner and Marsden, 1981). The chemical structure of MPTP is similar to that of many pyridines found in the environment and, in particular, to that of chemicals commonly used in agriculture. Several case control studies have shown associations between PD and rural living, well water drinking and exposure to herbicides and pesticides (Golbe and Langston, 1993). The oxidation of MPTP to its reactive metabolite 1‐methyl‐4‐phenylpyridinium (MPPþ) and its subsequent ability to impair mitochondrial function and generate oxidative stress are key components of the toxic process initiated by this agent (Jenner and Olanow, 1996). MPTP and its metabolite MPPþ can directly react with biomolecules to form free radical species (Adams and Odunze, 1991). The toxic effects of MPPþ are thought to be caused by its ability to inhibit complex I of the mitochondrial respiratory chain resulting in decreased cellular adenosine triphosphate levels (ATP) (Scotcher et al., 1990) and altered intracellular calcium content. Alterations in the homeostasis of intracellular calcium are closely linked with altered cell function and cell death. Recent studies have shown that the calcium binding protein calbindin is selectively decreased in the substantia nigra in PD (Damier et al., 1999). The inhibition of mitochondrial function may lead to increased formation of radical species in several ways, including an overspill of superoxide radicals from the electron transport chain. Free radicals are capable of reacting almost instantaneously with membrane lipids and causing lipid peroxidation, alteration in membrane fluidity and, ultimately, cell death. Superoxide dismutase (SOD) converts superoxide, a free radical involved in MPTP neurotoxicity, to hydrogen peroxide (H2O2). In the substantia nigra in PD an increase in the activity of the mitochondrial form of this enzyme has been reported with no change in total or cytosolic enzyme activity

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(Saggu et al., 1989). These data suggest an increased formation of superoxide radicals in proximity to mitochondria, which induces an increase in SOD activity. The discovery of MPTP toxicity led to the recognition that several naturally occurring molecules, such as isoquinolines (Yoshida et al., 1990), b‐carbinoles (Matsubara et al., 1995), n‐hexane (Pezzoli et al., 1995; Pezzoli et al., 2000), and similar hydrocarbons may induce MPTP‐like toxicity. Recently, D‐b Hydroxybutyrate has proved to possess neuroprotective properties against the damage induced by the toxic agent MPPþ (Kashiwaya et al., 2000).

2.2 Reactive Species and Oxidative Damage Free radicals and other so called reactive species are constantly produced in vivo by all body tissues. Some are generated by the ‘‘leakage’’ of electrons from mitochondrial electron transport chains (ETC) (Halliwell and Gutteridge, 1999) and others come from reactions of autoxidable molecules with molecular oxygen. It has been suggested that excess free radical formation plays a role in the pathogenesis of PD (Olanow et al., 1998). In particular, oxidation reactions and toxic oxygen species may contribute to the degenerative process underlying neuronal death in PD. Excess formation of free radicals may occur as a result of toxicity (see previous paragraph). Another important mechanism linked to oxidation reactions and free radical formation in the pathogenesis of PD is related to the metabolism of dopamine. Dopamine can be oxidatively metabolized by the enzyme monoamine oxidase (MAO). Both auto‐oxidation of dopamine and oxidative deamination by MAO result in the formation of H2O2. In the presence of iron, H2O2 can be reduced to form the toxic hydroxyl free radical (OH) (Fenton reaction) able to initiate and propagate the lipid peroxidation reactions. Recently a selective increase in iron content was revealed in the SNpc in PD patients (Jenner, 2003). Under normal circumstances H2O2 is normally removed from the brain glutathione system. Glutathione peroxidase catalyzes the reaction of H2O2 with glutathione (GSH) to form glutathione disulfide (GSSG). A study has shown that the concentration of GSH is reduced in the SNpc of patients with PD compared with control subjects, while the concentration of GSSG is unaltered (Jenner and Olanow, 1996). The decreased concentration of nigral GSH in PD could be the result of neuronal loss, since a positive correlation has been found between the GSH content and the severity of neuronal depletion in the parkinsonian brain. Moreover, in PD, the surviving neurons of the nigrostriatal pathway exhibit an increased dopamine turnover. This could theoretically be associated with oxidative stress as a consequence of increased production of H2O2 during the oxidative deamination of dopamine by MAO. All these evidence supports the theory that dopamine and L‐dopa (one of the most effective drugs for the treatment of PD symptoms) administration is able to promote free radical formation and oxidative stress and, hence, oxidation damage and cell death. However, there is no actual evidence to support any toxic effect of dopamine and L‐dopa in primates or in humans (Hefti et al., 1981; Lyras et al., 2002). On the contrary, the ELLDOPA study, a randomized, double‐blind controlled clinical trial comparing three dosages of carbidopa/levodopa (12.5/50 mg tid, 25/100 mg tid and 50/200 mg tid) to matching placebo for 40 weeks, suggested that levodopa does not hasten disease progression, but rather that it may slow it down (Fahn and The Parkinson Study Group, 2005). Nitric oxide (NO) also plays several roles in the process leading to PD. The toxic action of NO is likely to be mediated by peroxynitrite formed by an interaction between nitric oxide and superoxide radicals. There is evidence that NO can react with iron and stimulate lipid peroxidation (Reif and Simmons, 1990; Radi et al., 1991). NO can also inhibit mitochondrial respiration, although it inhibits complex IV rather than complex I (Bolanos et al., 1994; Cleeter et al., 1994). Studies in postmortem parkinsonian brains show that neurons that contain nitric oxide synthase (NOS) are selectively spared (Mufson and Brandabur, 1994) and that there is increased NO formation and related oxidative damage (Jenner and Olanow, 1996). Finally, the contribution of the microglia (the inflammatory cells of the brain) to the oxidative damage mechanism of PD should be mentioned. Resting microglia are sensitive to any form of disturbance within

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the neuronal microenvironment. In the presence of any degree of pathologic change they rapidly activate giving origin to a highly localized inflammatory response (Kreutzberg, 1996) and producing a wide variety of substances, including not only proinflammatory prostaglandines and cytokines, but also reactive oxygen species (ROS) and reactive nitrogen species (RNS) able to increase oxidative stress in the ways described earlier (Hunot et al., 1996).

2.3 Oxidation Products The excessive formation of ROS and RNS in PD damages key cellular components and there is evidence for increased oxidative damage to lipids, proteins, and DNA. Lipid peroxidation results in loss of membrane polyunsaturated fatty acids and in increase of oxidized phospholipids as polar species contributing to increased membrane rigidity (Farooqui et al., 1998). Lipid peroxidation leads also to the production of 4‐hydroxynonenal (HNE), and levels of HNE are increased in dopaminergic cells in SNpc and in cerebrospinal fluid of PD patients in respect to controls (Yoritaka et al., 1996; Picklo et al., 1999). HNE is a lipophilic a, b‐alkenal that is highly reactive and that forms stable adducts with nucleophilic groups on proteins such as thiols and amines. HNE can induce apoptosis with activation of caspases‐8, ‐9, and ‐3 and DNA fragmentation (Liu et al., 2000). It inhibits NK‐kB signaling pathways, decreases GSH levels and inhibits complexes I and II of the mitochondrial ETC (Camandola et al., 2000). HNE also binds to proteasomes, inhibiting the process of protein ubiquination. Baseline protein oxidation is high in the SNpc and the levels of protein carbonyls are increased in all the brain regions of PD patients (Alam et al., 1997; Floor and Wetzel, 1998). Oxidized proteins may not be adequately ubiquinated or recognized by the proteasome and thus accumulate within cells. HNE also can interact with proteins and enhance their cross‐linking. Finally mitochondrial DNA damage occurs in PD, which presumably might result in impairment of the respiratory chain leading to inhibition of complex I (Zhang et al., 1999). In conclusion, the findings of altered iron metabolism, impaired mitochondrial function and increased lipid peroxidation support the importance of oxidative stress for neuronal degeneration in the substantia nigra of patients with PD. These alterations do not explain, however, why nigral neurons are not protected by normal detoxification mechanisms. Toxic compounds formed in the brain are normally inactivated by various protective mechanisms. These mechanisms, however, may be impaired in PD, e.g., reduced activity of catalase, peroxidase, and glutathione peroxidase and diminished concentrations of reduced GSH have been found in the substantia nigra in PD. Protection against such oxidative damage could be provided by scavengers of free radicals and antioxidants such as MAO‐B inhibitors, alpha‐tocopherol (Vitamin E), ascorbic acid (Vitamin C), GSH, (Halliwell, 2001), and iron chelators, although a clear protective role towards the progression of PD has still not been clearly demonstrated. The DATATOP study, a 4‐armed randomized clinical trial comparing placebo, tocopherol supplements (2000 IU daily)þdeprenyl placebo, active deprenyl þ tocopherol placebo, and both active substances together, did not disclose any benefit with tocopherol treatment (Parkinson Study Group. 1993). However, epidemiological data suggests that a high dietary intake of vitamin E reduces the risk of developing PD (de Rijk et al., 1997).

3

Lipid Influence on a‐Synuclein Behavior in PD

a‐Synuclein (aS) is a 140‐amino acid protein that is most highly expressed in the central nervous system (CNS) and is particularly enriched in presynaptic nerve terminals. Although the normal function of aS remains poorly understood, many possible potential roles have been proposed, including membrane stabilization (Saito et al., 2004), lipid metabolism regulation (Cole et al., 2002), enzyme phospholipase D2 regulation, (Jenco et al., 1998), and dopamine uptake in synaptic vesicle regulation (Lee et al., 2001; Wersinger and Sidhu, 2003) to mention only the main ones.

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A key finding that suggested that aS is involved in the neurodegenerative PD process was that only lipofuscin‐ or neuromelanin‐laden, aging neurons show a propensity to develop aS‐positive PD‐related cytoskeletal abnormalities (Braak et al., 2001). Another PD‐related finding was that mutation or overexpression of the human aS gene (Snca) causes early‐onset autosomal dominant PD (Polymeropoulos et al., 1997; Kruger et al., 1998). The mechanisms of action, by which the toxic effects of aS are manifested in neuronal cells, are still under investigation and debate although, in recent years, many in vitro and in vivo studies are supporting the hypothesis, according to which the interaction of aS with vesicle and membrane lipids is responsible for the damage to SNpc observed in PD. In particular, the aS protein contains seven imperfect 11‐mer repeats similar to exchangeable class A apolipoproteins (Scherzer and Feany, 2004). aS resembles a apolipoprotein and is natively unfolded, when free in solution, but undergoes a structural transition to an a‐helix conformation in the presence of lipid vesicles (Clemens et al., 2004). When bound to lipid vesicles in the a‐helical conformation, aS is a relatively stable protein, vice versa, when present in the cytosolic solution at relatively high concentrations, aS can auto aggregate in b‐sheet filamentous amyloid‐like fibrils, giving origin to insoluble intracellular inclusions with the capacity to cause neuronal damage (> Figure 23-2). This theory is supported by in vitro studies, which show that aS is a conformationally dynamic molecule

. Figure 23-2 Mechanism of a‐Sinuclein fibril formation (left image) and Atomic Force Microscope 3‐dimensional view of an a‐Sinuclein fibril (right image)

whose secondary structure is very dependent on its environment. Indeed, aS has a binding affinity for the surface of lipid vesicles and lipid bilayer membranes, which is strongly influenced by the lipid composition of the membrane itself (Cole et al., 2001; Zhu et al., 2003). Moreover, at relatively high concentrations of aS, interaction with lipids leads to formation of oligomeric protofibrils with amyloidogenic properties capable of disrupting membrane bilayers by forming annular pores in the membrane (Lashuel et al., 2002). Postmortem analysis disclosed that the high molecular weight polymers of aS protein are a major component of LB, the cytoplasmic protein aggregates hallmark for PD, as well as the protein aggregates seen in axons and dendrites of PD patients, referred to as LN (Dawson and Dawson, 2003). Lipids and vesicle membranes are also found in LB, and lipid‐bound cytosolic aS levels are increased in extracts from the brains of PD patients compared to controls (Sharon et al., 2003). The effects of two PD‐linked missense mutations in the gene coding for aS, A30P and A53T have also been studied (Bussel and Eliezer, 2004). These mutations have been linked to the familial forms of early onset PD. In particular, the lipid interaction and the oligomeric autoassembly capacity of A30P and A53T aS have been investigated. Both of these mutations have been found to enhance the rate of oligomerization of the free protein (El‐Agnaf et al., 1998; Giasson et al., 1999) and both A30P and A53T mutations also appeared to decrease the extent of aS membrane binding capacity in vitro and in vivo (Perrin et al., 2000; Jo et al., 2002), thus decreasing the intracytoplasmatic amount of the a‐helix stable protein conformation, justifying the early onset of the hereditary form of PD‐linked to Snca mutation.

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In conclusion, the effect of membrane lipids on aS aggregation is very sensitive to both relative concentrations and membrane composition, as well as to the quantity of aS in the intracytoplasmatic environment, so it is difficult to predict the effects on neurons. However, it is clear that changes in lipid composition because of cellular stress, for example, could have significant effects on aS fibrillation. Membranes, in fact, are major sites of lipid peroxidation; hence, recruitment of aS to membrane environments could be an early step in the generation of pathological aS aggregates.

4

The Role of Cannabinoids in PD

It has become increasingly apparent that PD involves many transmitter systems other than dopamine. Endocannabinoids are a class of bioactive lipids responsible for the activation of type 1 (CB1) and type 2 (CB2) cannabinoid receptors. A close functional interaction between dopaminergic transmission and the endocannabinoid system during motor function has been proposed (Piomelli, 2003). In support of this statement, abundance of the cannabinoid CB1 and CB2 receptor subtypes, as well as of endocannabinoids have been found in the basal ganglia and the cerebellum, the areas that control movement. Furthermore, the demonstration of a powerful action of plant‐derived, synthetic, and endogenous cannabinoids on motor activity, and the occurrence of marked changes in endocannabinoid transmission in the basal ganglia of humans affected by several motor disorders, have strengthened the idea that cannabinoids and their receptors play a key role in the control of movement in health and disease (Fernandez and Gonzales, 2005). It has been recently found that levels of anandamide (N‐arachidonoylethanolamine), one of the endogenous ligands of CB1, extracted from the cerebrospinal fluid of PD patients, were more than doubled with respect to age matched controls (Pisani et al., 2005). Since anandamide was shown to stimulate dopaminergic release (Gubellini et al., 2002; Giuffrida et al., 2004), it is reasonable to believe that the increase in anandamide might reflect a compensatory mechanism occurring in the striatum of PD patients, where CB1 receptors are densely expressed, aimed at normalizing chronic dopamine depletion. Based on these findings, it should be borne in mind that cannabinoid‐based compounds, which act at key steps of the endocannabinoid transmission, might be of interest because of their potential ability to alleviate motor symptoms. For instance, the majority of PD patients undergoing levodopa therapy develop disabling motor complications, dyskinesias, within 10 years of treatment. Stimulation of cannabinoid receptors is emerging as a promising therapy to alleviate L‐dopa associated dyskinesias (Ferrer et al., 2003) although at present, the literature on cannabinoid therapy effects on PD patients is still very controversial (Segovia et al., 2003; Corey, 2005) and a complete vision and understanding of the subject has yet to be achieved.

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Radi R, Beckman JS, Bush KM, Freeman BA. 1991. Peroxynitrite‐induced membrane lipid peroxidation: The cytotoxic potential of superoxide and nitric oxide. Arch Biochem Biophys 288: 481-487. Reif DW, Simmons RD. 1990. Nitric oxide mediates iron release from ferritin. Arch Biochem Biophys 283: 537-541. Saggu H, Cooksey J, Dexter DH, Wells FR, Lees A, et al. 1989. A selective increase in particulate superoxide dismutase activity in parkinsonian substantia nigra. J Neurochem 53: 692-697. Saito H, Lund‐Katz S, Phillips MC. 2004. Contributions of domain structure and lipid interaction to the functionality of exchangeable human apolipoproteins. Prog Lipid Res 43: 350-380. Scherzer CR, Feany MB. 2004. Yeast genetics targets lipids in Parkinson’s disease. Trends genet 20: 273-277. Scotcher KP, Irwin I, De Lanney LE, Langston JW, Di Monte D. 1990. Effects of 1‐methyl‐4‐phenyl‐1,2,3,6‐tetrahydropyridine and 1‐methyl‐4‐phenylpyridinium ion in ATP levels of mouse brain synaptosomes. J Neurochem 54: 1295-1301. Segovia G, Mora F, Crossman AR, Brotchie JM. 2003. Effects of CB1 cannabinoid receptor modulating compounds on the hyperkinesias induced by high‐dose levodopa in reserpine‐treated rat model of Parkinson’s disease. Mov Disord 18: 138-149. Sharon R, Bar-Joseph I, Frosch MP, Walsh DM, Hamilton JA, et al. 2003. The formation of highly soluble oligomers of a‐sinuclein is regulated by fatty acids and enhanced in Parkinson’s disease. Neuron 37: 583-595. Wersinger C, Sidhu A. 2003. Attenuation of dopamine transporter activity by alpha‐sinuclein. Neurosci Lett 340: 189-192. Yoritaka A, Hattori N, Uchida K, Tanaka M, Stadtman ER, et al. 1996. Immunohistochemical detection of 4‐ hydroxynonenal protein adducts in Parkinson’s disease. Proc Natl Acad Sci USA 93: 2696-2701. Yoshida M, Niwa T, Nagatsu T. 1990. Parkinson in monkeys produced by chronic administration of an endogenous substance of the brain, tetrahydroisoquinoline: The behavioral and biochemical changes. Neurosci Lett 119: 109-113. Zhang J, Perry G, Smith MA, Robertson D, Olson SJ, et al. 1999. Parkinson’s disease is associated with oxidative damage to cytoplasmic DNA and RNA in substantia nigra neurons. Am J Pathol 154: 1423-1429. Zhu M, Li J, Fink AL. 2003. The association of a‐sinuclein with membranes affects bilayer structure, stability, and fibril formation. J Biol Chem 278 (41): 40186-40197.

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Lipids in Multiple Sclerosis

L. Rinaldi . F. Grassivaro . P. Gallo

1 Lipids in the Myelin Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 594 2 Exogenous Glycolipid Antigens and Autoimmunity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 595 3 Glycolipids and Autoimmune Demyelination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 595 4 Sulfatides in EAE and MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 597 5 Sphingolipids and Autoimmunity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 597 6 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 598

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_24, # Springer ScienceþBusiness Media, LLC 2009

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Abstract: Increasing evidence suggests that lipids antigens may be target of autoimmune attacks in inflammatory diseases of the central nervous system. Preliminary observations in multiple sclerosis and in experimental autoimmune encephalomyelitis indicate that both T cell and antibody reactivity to structural lipids of myelin may play a role in determining autoimmune-mediated demyelination. Molecular mimicry between bacterial and myelin glycolipids has been observed, and a subset of NKT cells specific for glycolipid antigens has been identified. Increased sulfatide-reactive T-cells and antibodies were demonstrated in multiple sclerosis patients. The possibility that myelin lipids may be targets for autoimmune reactions in central nervous system diseases opens new research and therapeutic perspectives for multiple sclerosis. List of Abbreviations: (APC), antigen presenting cells; (EAE), experimental autoimmune encephalomyelitis; (IFN), interferon; (LPA), lysophosphatidic acid; (MHC-I), major histocompatibility complex class I; [MBP], myelin basic protein; [MOG], myelin oligodendrocyte protein; (NOD), non-obese diabetic; (OPC), oligodendrocyte progenitor cell; [PLP], proteolipid protein; (TCR), T cell receptor; (TNF), tumor necrosis factor

1 Lipids in the Myelin Structure The myelin sheath is formed by extensions of oligodendrocyte cell membranes that wrap around the axon to form a cylindrical coating a few tens of micrometers in diameter (Boggs and Moscarello, 1978; Morell and Norton, 1980). The sheath consists of repeat units of ‘‘double’’ bilayers separated by 3- to 4-nm-thick aqueous layers that alternate between the cytoplasmic and extracellular spaces (Boggs and Moscarello, 1978). Seventy to eighty percent of the dry weight of myelin consists of lipids, a proportion that is significantly higher than in most other cell membranes of the human body. Despite lipids constitute about one-half of brain tissue dry weight, only recently their immunological properties have been recognized as possible targets/players in the immunopathological processes taking place during autoimmune diseases of central nervous system (CNS). Among the complex lipids, sphingolipids play a crucial role as modulators of transmembrane signaling, mediate cellular interaction, modulate the behavior of cellular protein and receptors, and participate in signal transduction (Merril, 2002). Indeed, sphingolipids constitute 20–35% of plasma membrane lipids, and the ratios sphingolipids:glycerophospholipids:sterols in myelin are 28:44:28. Oligodendrocytes produce huge amounts of myelin membrane, estimated at as much as 5,000–50,000 mm2 of myelin surface area per cell per day during the period of active myelin assembly (Pfeiffer et al., 1993). Myelin formation is accomplished by lipid synthesis, especially galactosylceramide and its sulfated analogue sulfatide. The composition of brain myelin is largely conserved among mammalian species. Although there are no ‘‘myelin-specific’’ lipids, myelin is significantly enriched for certain lipids, in particular galactocerebroside (GalC). Gangliosides, a class of glycosphingolipids, constitute 0.1–0.3% of myelin lipids, being the monosialogangliosides GM1 and GM4 quantitatively enriched in human myelin. Myelin has also large amounts of cholesterol and ethanolamine-containing plasmalogens as well as lecithin, while sphingomyelin is a relatively minor component. Finally, quantitatively minor lipid components of myelin include polyphosphoinositides, at least three fatty acid esters of cerebroside and glycerol-based galactosyldiglycerides. Recently, the role of lysophospholipids as intercellular signaling molecules has been recognized. Two of the best-studied lysophospholipids, lysophosphatidic acid (LPA), and sphingosine 1-phosphate (S1P), activate (through cognate G-protein-coupled receptors) many well-known intracellular signaling pathways, leading to a variety of biologically important cell responses. Lysophospholipids and their receptors have been found in a wide range of tissues and cell types, indicating their relevance in many physiological processes, including reproduction, vascular development, cancer and nervous system function (Anliker and Chun, 2004a, b; Ishii et al., 2004). A number of inflammatory demyelinating diseases, of which multiple sclerosis (MS) is the most common, result in myelin sheath de-adhesion and swelling, and ultimate vesiculation and fragmentation. Over the last decade, increasing evidence has highlighted the possibility that many of the above-mentioned

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myelin lipids may represent the antigenic targets of the autoimmune reaction(s) underlying these diseases. In this chapter, we review the current knowledge on this topic.

2 Exogenous Glycolipid Antigens and Autoimmunity Multiple Sclerosis (MS) is a complex-trait disease in which exogenous agents (viruses, bacteria, etc.), linked to the environment, are likely to cooperate with endogenous factors causing an autoimmune reaction against CNS antigens, including myelin antigens (Williams et al., 1994; Stinissen et al., 1997; Kurtzke, 2000; Franklin and Nelson, 2001; Dhib-Jalbut, 2007). Indeed, several antigens shared between pathogens and myelin/axons structures (molecular mimicry) may activate CNS-specific lymphocytes (Libbey et al., 2007). Many attempts have been made to discover homologies between foreign antigens and self-components of myelin/axons. Although myelin sheath contains less than 30% proteins, myelin proteins (myelin basic protein [MBP], myelin oligodendrocyte protein [MOG], and proteolipid protein [PLP]) are considered the main antigenic targets in MS (de Rosbo and Ben-Nun, 1998; Reindl et al., 1999; Egg et al., 2001; Lalive et al., 2006). However, in recent years, increasing evidence has revealed that self glycolipids can also be the targets of autoreactive T cells as well (Shamshiev et al., 1999; De Libero et al., 2002; De Libero and Mori, 2003). Indeed, glycolipid autoantigens have been specifically linked to the MS (Sadatipour et al., 2000; Kantner et al., 2006), and anti-glycosphingolipid autoreactivity was demonstrated to be induced by bacterial infection (Kronenberg et al., 2005; De Libero et al., 2006). Moreover, several bacteria have been implicated in the etiology of MS, including Chlamydia pneumoniae (Moses and Sriram, 2001; Contini et al., 2004; Dong-Si et al., 2004), Mycoplasma pneumoniae (Maida, 1983), and Borrelia burgdorferi (Lana-Peixoto, 1994; Chmielewska-Badora, 2000; Brorson et al., 2001; Wolfson and Talbot, 2002; Fritzsche, 2005). It is well known that B. burgdorferi infection may precede the development of an autoimmune disease (Gross et al., 1998; Steere et al., 2001), the geographic distribution of MS parallels the distribution of tickborne diseases (Brown et al., 1996), and both MS and B. burgdorferi infection have been associated with autoimmune thyroid conditions (Schwid et al., 1997; Benvenga et al., 2004). All these observations suggest that lipid antigens, shared between pathogens and myelin structures, may activate self-reactive myelin-specific lymphocytes, thus leading to the development of an autoimmune diseases of the CNS. Nevertheless, autoimmune responses to myelin lipids have been studied much less extensively than responses to myelin proteins, mainly due to the lack of suitable technologies, the large number of potential lipid antigens, the hydrophobicity of lipids, and the technical difficulty of detecting lipid-specific B- and T-cell responses. Autoimmune responses directed against phospholipids and gangliosides have been demonstrated to contribute to the pathogenesis of systemic lupus erythematosus and Guillain–Barre´ syndrome, respectively (Fredman, 1998), while the role of lipid-specific autoimmunity in MS remains controversial (Giovannoni et al., 2000). Recently, by means of a large-scale lipid microarrays for detection of autoantibodies in biological fluids, antibodies to sulfatide, sphingomyelin and oxidized lipid have been detected in cerebrospinal fluid samples from individuals with MS and in sera of mice suffering from experimental autoimmune encephalomyelitis (EAE). Furthermore, immunization of mice with sulfatide or transfer of sulfatide-specific antibodies induces severe clinical manifestations in EAE (Kanter et al., 2006). Thus, autoimmune responses to sulfatide and other lipids are present in individuals with MS and in EAE, and may contribute to the pathogenesis of autoimmune demyelination. Finally, myelin lipid compositions was demonstrated to differ between individuals with MS and healthy individuals (Gerstl et al., 1961; Alling et al., 1971). In particular, individuals affected from MS have increased cholesterol esters compared to controls (Cumings et al., 1955), thus suggesting the possibility that individuals suffering from MS may develop immune responses to lipids that are altered by oxidative processes (Alling et al., 1971).

3 Glycolipids and Autoimmune Demyelination Two bacterial-derived glycolipid antigens were found to share structural similarities with myelin GalC: the B. burgdorferi glycolipid 2 (BbGL-2) antigen (Ben-Menacem et al., 2003) and the Sphingomonas antigen

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GalAGSL, a glycosphingolipid containing galacturonic acid (Tupin et al., 2007). It has been argued that these bacterial antigens may induce autoreactivity against self-glycosphingolipids in inflammatory CNS diseases. That GalC can be a target autoantigen in MS is suggested by the observation of elevated levels of anti-GalC antibodies in MS patients compared to healthy controls (Kasai et al., 1986). In experimental models of autoimmune demyelination, anti-GalC antibodies can cause demyelination (Fry et al., 1974; Raine et al., 1981) and induce an extensive degradation of MBP (Menon et al., 1997). Particularly interesting in defining a possible role of galactocebrosides as autoantigens in MS, is the role played by NKT (Natural Killer T) cells, a peculiar T cell subpopulation discovered just over a decade ago and playing a major role in glycolipid recognition and processing (Bendelac et al., 2007). Indeed, while conventional T lymphocytes recognize peptide antigens presented by polymorphic major histocompatibility complex class I (MHC-I) or class II (MHC-II) molecules, NKT cells are specific for glycolipid antigens presented by the non-polymorphic MHC-I-like molecule CD1d (Bendelac et al., 1997; Godfrey et al., 2000; Joyce, 2001). Although NKT cells express T cell receptor (TCR) a- and b-chains, the TCR diversity for most of them is very limited, due to their expression of a single a-chain (Va14-Ja18 in mice, Va24-Ja18 in human) coupled with a b-chain rearranged with a limited Vb gene segments (Vb8.2, Vb2 and Vb7 in mice,Vb11 in human). Va14 in mice and Va24- Vb11 in human are called invariant-NKT cells (iNKT). NKT cells are particularly worth of interest in autoimmunity for their capacity to produce a wide spectrum of cytokines, including interleukin (IL)-4, IL-10, IL-13, interferon (IFN)-g, tumor necrosis factor (TNF)-a, and osteopontin (Bendelac et al., 1997; Godfrey et al., 2000; Joyce, 2001). However, NKT cells seem to produce only a set of Th1, Th2, or Th17 cytokines in different pathophysiological conditions, depending on the local cytokine milieu. Activated NKT cells may have cytotoxic activities as well. Because NKT cells lack immunological memory and express a restricted set of antigenspecific receptors, they are located at the bridge between the innate and the adaptive arm of the immune response with a main immunoregulatory role. In this context, NKT cells resemble gDd T cells and B-1 B cells (Bendelac et al., 2001). The most potent agonist of NKT cells is alpha-galactosylceramide (a-GalCer) (Wu et al., 2006), an exogenous sponge-derived glycosphingolipid containing an a-anomeric linkage between the sugar and ceramide, though, within mammalian cellular membranes, glycosphingolipids typically contain a b-anomeric linkage. A physiological role for a-GalCer in mice and humans has not been established, but a-anomeric linkage is critical for NKT activation, since b-Gal-Cer is not able to activate NKT cells. So far, no endogenous ligands for NKT cells have been identified, except for isoglobotrihexosylceramide (iGb3), which, strangely, has the ability to stimulate NKT cells despite the b-linkage between the sugar and ceramide (Zhou et al., 2004). However, the only tissue containing iGb3 is the dorsal root ganglion of mice, and no iGb3 was detected in any human tissue (Porubsky et al., 2007; Speak et al., 2007), leaving opened the search for endogenous ligands for human iNKT cells. The above mentioned Sphingomonas antigen GalAGSL may function as a ligand of iNKT cells (Kinjo et al., 2005; Mattner et al., 2005; Sriram et al., 2005), and certain Sphingomonas strains were found to be pathogenic for the human beings (Yamazaki et al., 1996). Recently, NKT cells were demonstrated to recognize B. burgdorferi’s glycolipid-2, BbGL-2 antigen (Kinjo et al., 2006). Given the structural similarities between myelin glycolipid GalC and the NKT cell agonist a-GalCer, the possibility exists that NKT cells may serve to protect against myelin-directed autoreactivity by recognizing bacterial-derived lipid antigens that share antigenic structures similar to the major myelin components. The role of NKT cells may be to recognize those exogenous antigens that can induce autoreactivity via their structural similarity to self antigens (Hammond and Godfrey, 2002). Interestingly, in MS patients NKT cells were demonstrated to be defective in IL-4 production (Linsen et al., 2005), thus suggesting that NKT cells may play a peculiar immunoregulatory role in MS and/or that MS patients have a genetic defect pertaining to NKT cells. Just as T-cell autoreactivity can be activated during microbial infection, so can NKT cells (Kinjo et al., 2005; Mattner et al., 2005). If NKT cells are deficient and do not properly combat the invading pathogen, autoreactivity may predominate and lead to the observed autoimmune demyelination in MS. A significant amount of experimental evidence in animal models of autoimmunity indicates that NKT cells can play a critical role in the regulation of autoimmune responses. Defects in NKT-cell development

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and/or function have been observed in diabetes-prone non-obese diabetic (NOD) mice, in lupus-prone lpr and NZB/NZW mice, and in SJL mice that are highly susceptible to induction of acute monophasic EAE, an experimental model for autoimmune demyelination of the SNC. Similar defects in NKT cell numbers and function were observed in patients with systemic sclerosis, type 1 diabetes, rheumatoid arthritis, lupus, and MS. Additional studies showed that transgenic over-expression of the invariant NKT TCR ameliorates diabetes (Laloux et al., 2001), whereas CD1d-defiency exacerbates diabetes (Shi et al., 2001; Wang et al., 2001) in NOD mice. Collectively, these findings have revealed a suppressive role of NKT cells in autoimmunity and suggested a central role of lipid recognition in immunoregulation.

4 Sulfatides in EAE and MS In EAE, sulfatide-reactive T cells were found to constitute 3–4% of the CNS-infiltrating T cell population, whereas iNKT cells constituted a very minor part (0.6–0.9%). The finding that sulfatide-reactive T cells were enriched in the CNS during EAE was found in contrast to what was observed in the peripheral organs of both normal and EAE animals, where iNKT cells exceeded sulfatide-reactive T cells by a factor of 5. Interestingly, sulfatide-CD1d tetramer+ T cells could not be detected in the CNS of naive mice or animals, which were immunized for disease but did not develop clinical signs of EAE (Zajonc et al., 2005). Sulfatide, presented by CNS-resident antigen presenting cells (APC), such as microglia, can activate T cells leading to inflammatory demyelination. This pathway can be targeted for intervention in EAE. Since, as mentioned above, CD1 molecules are nonpolymorphic, insight into the role of sulfatide presentation by microglia, and their interaction with a unique CD1d-restricted T cell population in the CNS, will be extremely valuable for developing HLA-independent approaches to treat autoimmune demyelinating diseases in humans. A possible role for sulfatide as autoantigen in the immunopathological process is that it is believed to induce MS arises from several observations in patients. First, anti-glycolipid, including anti-sulfatide, antibodies have been detected not only in individuals suffering from polyneuropathy syndromes but also in MS patients (Endo et al., 1984; Uhlig and Dernick, 1989; Stevens et al., 1992; Bansal et al., 1994; Ilvas et al., 2003). Second, lipid microarrays have confirmed enhanced antibody response against sulfatide and other myelin lipids in MS patients, and their potential to enhance experimental autoimmune demyelination (Kanter et al., 2006). Third, T lymphocytes reactive to glycolipids have been isolated from active MS patients, where their frequency has been shown to be threefold higher than that observed in normal individuals (Shamshiev 1999, 2000). Recently, it has been demonstrated that the ganglioside GM1 binds well to CD1b, whereas sulfatide binds promiscuously to all of the CD1 (a–d) molecules (Shamshiev 1999, 2000). Finally, in chronic-active MS lesions, CD1 molecules are up-regulated on macrophages in areas of demyelination, but not in silent lesions in the brain (Battistini et al., 1996). Taken together, these observations suggest the possibility that myelin glycolipids may be presented locally during inflammation in the CNS.

5 Sphingolipids and Autoimmunity Another recently described aspect involving lipids in autoimmune diseases, including MS, is the role of sphingolipids as main mediators of lymphocyte egress from secondary lymphoid tissues. The pathogenesis of organ-specific autoimmune diseases is classically conceived as a complex process, which starts in the periphery, with the activation of autoreactive T cells in lymphonodes, and ends in the target organ where autoreactive T cells migrate, recognize specific self-antigens, and determines the inflammatory cascade leading to tissue damage. While the molecular components involved in lymphonode homing (chemokines and chemokine receptors) had been well characterized in the last decade, the molecular mechanism(s) of lymphocyte egress from secondary lymphoid tissues to lymph still remained unclear. However, recent evidence has identified a sphingolipid, sphingosine 1-phosphate (S1P), and its receptor, S1P1, constitutively expressed by all lymphocytes, as the main players in the lymphocyte recirculation system. Most evidence of this crucial S1P/S1P1 interaction derives from studies on 2-amino-2-[2-(4-octylphenyl)

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ethyl]propane-1,3-diol hydrochloride (FTY720, fingolimod), a new synthetic compound shown to be capable to significantly prolong skin and cardiac allograft survival and host survival in lethal graft versus host reaction in rats. FTY720, after phosphorylation, is suggested to act as an agonist of the S1P1 receptor, down-regulating S1P1 expression on lymphocytes, and inhibiting S1P/S1P1-dependent lymphocyte egress from secondary lymphoid tissues and thymus (Mandala et al., 2002; Goetzl and Rosen, 2004; Sanchez and Hla, 2004; Sawicka et al., 2005). Thus, FTY720 causes the sequestration of circulating mature lymphocytes into lymphoid tissues and modulates the recirculation of lymphocytes between blood and lymphoid tissues. As a consequence, FTY720 is presumed to decrease the trafficking and the infiltration of antigenspecific T cells into grafted organs or inflammatory sites in autoimmune diseases, thereby exerting an immunomodulatory activity. This new therapeutic approach has been tested in autoimmune diseases, as well as in MS. A recent Phase II study has demonstrated that oral FTY720 is effective in reducing disease activity in relapsing MS with a favorable adverse-effect profile. These results are awaiting confirmation in the three ongoing Phase III clinical trials evaluating FTY720 for relapsing–remitting MS. A number of studies have demonstrated the expression of S1P receptor mRNA in brain cells (for a review, see Dev et al., 2008). However, the lack of good antibodies and/or radio-ligands has not allowed the investigation of the distribution of S1P receptors at both tissue and subcellular levels. Moreover, due to the lack of selective S1P receptor antagonists and agonists, it remains unclear which specific S1P receptor subtype(s) are involved in the respective effects produced by FTY720 and S1P in the brain. Nevertheless, what is noteworthy is the observation that while S1P2 receptor appears to inhibit neurite outgrowth and plays a role in neuronal development, axonal growth, and synaptic excitability, S1P1 may promote neurite outgrowth. As FTY720 is not an agonist for S1P2 receptors (i.e., it lacks affinity for S1P2), it is tempting to speculate that this drug avoids the inhibitory actions of S1P2 receptor activation, while positively regulating neuronal function via its activity on S1P1 receptors. Thus, based on this hypothesis, FTY720 may promote repairing endogenous processes, dampening those involved in neurodegeneration. In oligodendrocyte progenitor cell (OPC) cultures derived from human fetal CNS, FTY720 has been shown to exert dose and time-dependent effects on OPC process extension, differentiation, and survival (Miron et al., 2007). These functional effects, which are important cellular events in the process of remyelination, were linked to the modulation of specific S1P receptors. Moreover, FTY720 could significantly attenuate OPC apoptosis in vitro (Miron et al., 2007). While these findings support a FTY720-induced modulation of S1P receptors on human OPCs, they raise the need to define the potential impact of chronic FTY720 therapy on remyelination in MS, and to consider how S1P signaling may be used as a potential approach to promote remyelination.

6 Conclusions MS is currently considered an inflammatory and degenerative disease of the CNS due to an autoimmune attack to peptide antigens of the myelin sheath. Nevertheless, though T and B cell reactivity to myelin proteins have extensively been demonstrated in MS patients, the extent and specificity of autoimmune responses in MS remain incompletely characterized. Since lipids comprise over 70% of the myelin sheath, lipid antigens have been considered as possible targets of autoimmune demyelination. Indeed, T-cell and antibody reactivity to structural lipids of myelin have been observed not only in EAE, but also in MS. However, despite increasing evidence suggesting that myelin lipids may be target of the autoimmune response in MS, the role of lipidspecific autoimmunity in this disease still remains controversial.

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A. Brand-Yavin . E. Yavin

1

A Brief Account of the Pathophysiology of Oxidative Stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 604

2 2.1 2.1.1 2.1.2 2.2 2.2.1 2.2.2 2.2.3

Generation of Lipid-Derived Oxidation Byproducts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 605 Enzymatic Lipid Peroxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 607 Cyclooxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 607 Lipoxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 607 Nonenzymatic Lipid Peroxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 608 Isoprostanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 608 Malondialdehyde . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 609 4-Hydroxy-2-Trans-Nonenal . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 610

3 3.1 3.1.1 3.1.2 3.1.3 3.1.4 3.2 3.2.1 3.2.2 3.2.3 3.2.4

The Bioavailability of Lipid- and Water-Soluble Antioxidants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 610 Dietary Low Molecular Weight Antioxidants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 611 Ascorbic Acid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 611 Vitamin E . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 612 Carotenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 612 Lipoic Acid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 613 Endogenous Low Molecular Weight Antioxidants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 613 The Glutathione Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 613 Coenzyme Q . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 614 Membrane-Derived Antioxidants: The Case of the Vinyl Ether Bond . . . . . . . . . . . . . . . . . . . . . . . . . . 615 Membrane-Derived Antioxidants: The Case of DHA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 616

4 4.1 4.2 4.2.1 4.2.2 4.2.3 4.3 4.3.1 4.3.2

Biological Consequences of Lipid Peroxides Action . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 617 On Scrambling of Phospholipid Asymmetry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 617 On the Perinatal Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 620 Brain Ontogeny During Adequate or Adverse Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 620 Oxygen Deprivation and Its Adverse Outcome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 621 DHA: Purported Mechanisms in Neuroprotection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 621 Oxidative Stress in the Aging Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 623 Progressive Aging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 623 Pathologically Accelerated Aging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 624

5

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 624

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_25, # Springer ScienceþBusiness Media, LLC 2009

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Brain oxidative stress from a phospholipid perspective

Abstract: Oxygen-derived reactive species such as oxyl, peroxyl, and hydroxyl radicals are initiators of a damaging cascade of events known as oxidative stress (OS) whereby cellular macromolecules are attacked and cell damage may occur. Brain tissue is particularly vulnerable to damage, because on the one hand it has a high consumption of oxygen and contains ample radical-sensitive targets while on the other hand it is less endowed with a robust and diverse enzymatic and nonenzymatic antioxidant arsenal. Polyunsaturated fatty acids which are almost exclusively and ubiquitously found in membrane phospholipids (PL) are major radical-sensitive targets. As a result of the action of hydroxyl radicals, lipid peroxidation takes place and lipid hydroperoxides are generated. While these perturb membrane structure/function and can be deleterious to cells, they may also take part in redox reactions which may beneficially amend the harmful consequences of the injury. That may well depend on the topology of the PUFAs in the bilayer as well as on replenishment mechanisms to restore membrane integrity. Stress metabolites derived from docosahexaenoic acid (DHA, 22:6n-3) may act as neuroprotectants after being released from PL. On the other hand, lipid-bound DHA and the vinyl ether bonds of ethanol amine PLs constitute a large reservoir of endogenous metabolites which could also provide neuroprotection. In the developing brain the nutritional supply of DHA is crucial since the antioxidant cellular machinery is less developed. In the aging brain or in neurodegenerative disorders, the pronounced depletion of DHA-enriched PL may indicate a persistent oxidative catabolic degradation, unmatched by a proper reacylation of the PUFAs. Unravelling the lipidomic vocabulary of molecular species which contain DHA or are generated from DHA during development, aging, and trauma should pave the way for treating OS-related diseases. List of Abbreviations: AA, arachidonic acid (20:4n-6); AD, Alzheimer’s disease; amino-PL, aminophospholipids; APT, aminophospholipid translocase; CL, cardiolipin; COX, cyclooxygenases; CPG, choline phosphoglyceride; dEPG, N,N-dimethyl-EPG; DHA, docosahexaenoic acid (22:6n-3); EPA, eicosapentaenoic acid (20:5n-3); EPG, ethanolamine phosphoglyceride; FA, fatty acids; 4-HHE, 4-hydroxyhexenal; 4-HNE, 4-hydroxy-2-nonenal; LMWA, low molecular weight antioxidants; LOX, lipoxygenases; LPO, lipid peroxidation; MDA, malondialdehyde; NO, nitric oxide; OS, oxidative stress; pEPG, plasmalogen or 1-alkenyl 2-acyl EPG; PL, phospholipid; PLA2, phospholipase A2; PLSCR1, phospholipid scramblase 1; PUFA, polyunsaturated fatty acids; ROS, reactive oxygen; SPG, serine phosphoglyceride

1

A Brief Account of the Pathophysiology of Oxidative Stress

Oxidative stress (OS) is defined as the imbalance between excess generation of reactive oxygen species (ROS) and insufficient antioxidant defense mechanisms in the living organism. Oxygen free radicals are continuously formed as intermediates of enzymatic reactions during normal cellular function, e.g., mitochondrial respiratory activity and they may also be involved in growth regulation and intercellular signaling. At the cellular level, free radicals resulting from OS can produce damage by altering the function of cellular macromolecules, i.e., lipids, proteins, and nucleic acids and can ultimately cause cell death. The excessive production of ROS and other free radicals following ischemic or traumatic insults in brain tissue has been implicated in the damage and death of neural cells and has been reasoned as a main driving force in the etiology of a wide variety of acute and chronic neurological disorders (Halliwell, 2001; Reynolds et al., 2007). The sensitivity of the brain to the attack of free radicals and OS consequences which follow can be attributed in part to the high content of polyunsaturated fatty acids (PUFA), as well as to an overall lesser effective antioxidant defense. Additional reasons for susceptibility are the brain’s high oxygen consumption (about 20% of the total body oxygen intake), and accumulation of transition metal ions (i.e., iron and copper) both having the potential to exacerbate the risk of formation of hydroxyl radicals (Mattson, 2004: Gaeta and Hider, 2005). The sources of oxygen free radicals in the brain are diverse. About 95% of the oxygen taken up by the body is utilized by the mitochondria and the endoplasmic reticulum; the remaining being released as free radicals (Sheu et al., 2006; Terman and Brunk, 2006). In addition to mitochondrial metabolism, oxygen is used in various enzymatic reactions, mainly through oxidases and oxygenases. In the brain distinct enzymes form ROS as metabolic by-products, the most notorious being monoamine oxidase (MAO-A and MAO-B),

Brain oxidative stress from a phospholipid perspective

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tyrosine hydroxylase, and L-amino acid oxidase (Coyle and Puttfarcken, 1993). Likewise, nonenzymatic auto-oxidation of ascorbate and catecholamines produces hydrogen peroxide (H2O2). In the presence of divalent metal ions (i.e., iron, copper), H2O2 can react to yield highly reactive hydroxyl radicals (OH) via the Fenton reaction. The superoxide anion (O2) is one of the major ROS produced as a by-product of mitochondrial respiration and is a most deleterious compound. Therefore it is normally neutralized by superoxide dismutase (SOD), an enzyme which catalyzes the dismutation of superoxide into oxygen and H2O2, but in the presence of trace amounts of iron, it can give rise to hydroxyl radicals by the Haber–Weiss reaction. An additional source for O2 and OH is derived from N-methyl-D-aspartate receptors stimulation (Lafon-Cazal et al., 1993). The hydroxyl radical is considered a most destructive free radical capable of oxidizing amino acid residues of proteins, inactivating enzymes, damaging DNA and RNA and attacking lipids in the membrane. While H2O2, O2 and NO are practically nonreactive with lipids (Halliwell and Gutteridge, 2007), hydroxyl radicals will rapidly oxidize membrane lipids such as phospholipids, cholesterol and glycolipids, generating lipid hydroperoxides (LOOH). The latter may lead to further formation of radicals (propagation) thus disrupting membrane organization which may eventually enhance damage and cause subsequent cell death. Alterations in cell function which often lead to cell death by apoptosis or necrosis following OS, have been widely studied in animal models and in cell cultures of nervous system origin (Lehtinen and Bonni, 2006). Neuronal cells death by apoptosis (Heidenreich, 2003) has been suggested to occur in aging, Alzheimer’s disease (AD), amyotrophic lateral sclerosis, as well as in traumatic brain injuries such as stroke and ischemia. Apoptosis is characterized by cytoplasmic condensation, cell shrinkage, nuclear DNA fragmentation and membrane blebbing. Astrocytes which comprise the vast majority of cells in the adult brain appear to exert a remarkable capacity to tolerate stress and retain viability even when mitochondrial function is altered. Swelling of astrocytes is common after traumatic stimuli and in many pathological conditions, e.g., ischemia, hypoxia, and reoxygenation (Kimelberg, 2005). This is due to the impairment of ATP-driven transport systems and ion pumps which maintain the osmotic balance and regulate cell volume. The myelin forming oligodendrocytes are particular vulnerable to free radicals damage during development leading to white matter injuries (Khwaja and Volpe, 2008). Another minor cell population in brain which is actively involved in the OS cascade is the microglia. In most neurodegenerative disorders, activation of resting microglia results in the production of ROS, reactive nitrogen species, chemokines, and proinflammatory cytokines. When unleashed, these metabolites initiate a generalized inflammatory response which exacerbates neuronal injury or death (Block et al., 2007). In this review we shall briefly revisit the basic chemistry of free radicals attack on phospholipid (PL) targets and provide an account of the battery of antioxidants that may counteract this challenge in brain tissue. As such this review is by no means comprehensive in scope. It will address more specifically some of the consequences of lipid-derived oxidation products on scrambling PL asymmetry at the cellular level and will examine concisely two discrete periods in the functioning brain; one period which concerns its assembly into a functional organ around birth, and a second period which highlights its decay and aging, from a standpoint of phosphatidyl ethanolamine molecular species.

2

Generation of Lipid-Derived Oxidation Byproducts

Lipids have been identified as prime targets for free radicals of oxygen action in all organisms. Within lipids the most oxygen-sensitive molecules are the unsaturated fatty acids (FA) that contain between one to several double bonds. The higher the number of double bonds, the higher the risk to attack by free radicals and the greater the number of lipid peroxidation (LPO) products formed (Halliwell and Gutteridge, 2007). The hydrogens on the intervening methylene (–CH2) groups are especially reactive and can be easily removed by ROS generating dienyl radicals (L) which react instantly with oxygen leading to the production of peroxydienyl radicals (LOO): LH þ HO ! L þ H2 O L þ O2 ! LOO

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This reaction occurs under aerobic conditions and is the initiation step of a free radical chain reaction. During propagation of the chain reaction, the generated peroxydienyl radicals can remove hydrogen from methylene groups of neighboring PUFAs giving rise to lipid-hydroperoxide molecules and new dienyl radicals which are able to start the whole cycle all over again: LOO þLH ! LOOH þ L Termination of the free radical chain reaction will occur only when two radicals interact with each other or when chain breaking molecules such as antioxidants (e.g., alpha-tocopherol, T-OH) scavenge and neutralize the reactive radicals: LOO þT  OH ! LOOH þ T  OO The oxidized form of the antioxidant (in this example the tocopheroxyl radical) will then be reduced by endogenous redox cycles or specific enzymes to recover the original antioxidant molecule. > Table 25-1 summarizes the most relevant ROS discussed in this chapter:

. Table 25-1 Reactive oxygen species (ROS) Non lipid-derived reactive species

Lipid-derived reactive species

Free radicals Superoxide (O2•−)

Nonradicals Singlet oxygen (O21Δg)

Hydroxyl (HO•)

Hydrogen Peroxydienyl peroxide radical (LOO•) (H2O2)

Nitric oxide (NO•)

Free radicals Dienyl radical (L•)

Non-radicals Lipidhydroperoxide, LOOH

Alkoxydienyl (LO•)

Lipid peroxyl radicals can be generated as intermediates via nonenzymatic as well as via enzymaticcontrolled LPO reactions. The main difference between the two types of reaction is that during nonenzymatic reactions, peroxyl radicals can remove hydrogen radicals from any other activated C–H bond in close proximity whereas in the enzymatic reactions, the generated radicals are transformed within the enzyme complex and are not allowed to leave the complex at any time. Thus, enzymatic activities of cyclooxygenases (COX) and lipoxygenases (LOX) will produce lipid hydroperoxides in stereospecific reactions, while the nonenzymatically produced peroxyl radical, can react with other PUFA double bonds or various other activated C–H bonds in neighboring lipids, proteins or sugar residues inflicting damage and destruction of cellular macromolecules. A direct high throughput measurement of free radicals or labile hydroperoxides in the brain is difficult if not often impossible. Current methods used to assess OS are based on the quantification of relatively stable LPO end products, such as F2-isoprostanes, malondialdehyde (MDA) and 4-hydroxy-2-nonenal

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(4-HNE). A detailed account of methods available for measuring reactive species and oxidative damage was recently published (Halliwell and Whiteman, 2004).

2.1 Enzymatic Lipid Peroxidation 2.1.1 Cyclooxygenases The COX enzymes, more accurately termed as prostaglandin endoperoxide H synthases (PGHS), are responsible for the synthesis of prostanoids, i.e., prostaglandins, prostacyclin and thromboxane, a family of biologically active lipid mediators derived from FA precursors. The first step of prostanoid biosynthesis is the release of a 20-carbon essential FA containing three, four, or five double bonds from either PL by phospholipase A2 (PLA2) or from diacylglycerol via a combined action of phospholipase C and diacylglycerol lipase. The precursor FAs are either dihomo-g-linolenic acid (20:3n-6, DGLA), arachidonic acid (20:4n-6, AA) or eicosapentaenoic acid (20:5n-3, EPA), whereas the majority of eicosanoids are biosynthesized from AA. COX catalyses the synthesis of prostaglandin H2 (PGH2), a highly reactive intermediate that is the starting point for the biosynthesis of most other prostanoids. The enzyme contains two active sites: (1) a heme group with peroxidase activity, first responsible for the abstraction of a hydrogen atom from AA and (2) the COX site, where two molecules of oxygen are added to AA to form a bicyclic endoperoxide with a further hydroperoxy group in position 15 yielding prostaglandin G2 (PGG2). This hydroperoxide is then reduced by the peroxidase active site to form PGH2. All prostanoids derived from AA are of the series-2 eicosanoids (e.g., PGE2), while DGLA and EPA are the precursors of the series-1 (e.g., PGE1) and series-3 (e.g., PGE3) prostanoids, respectively (Phillis et al., 2006) (> Figure 25-1). . Figure 25-1 Arachidonic acid (AA) is the biosynthetic precursor of prostaglandin H2 (PGH2)

More recently it was found that the inducible isoform of COX-2, pre-acetylated by aspirin produced a potent group of anti-inflammatory compounds which derived from long-chain n-3 fatty acids, i.e., EPA and docosahexaenoic acid (22:6n-3, DHA). Those originating from EPA are called Resolvins of the E series (Resolvin E1 or RvE1) and those derived from DHA were called Resolvins of the D series (Resolvin D1 or RvD1) (Serhan et al., 2008). DHA also gives rise to yet another group of compounds, the docosanoids, which appear to be multifunctional regulators and neuroprotectants in brain tissue under stress. The docosanoids which contain conjugated triene structures are generated by enzymatic oxygenation of free DHA after being released by PLA2 following stress and are believed to act as endogenous mechanisms to prevent brain damage (Mukherjee et al., 2007; Bazan, 2007).

2.1.2 Lipoxygenases LOX comprise a family of iron-containing, nonheme enzymes that catalyze the dioxygenation of PUFAs. First a hydrogen atom is abstracted from the bis-allylic position of fatty acids generating a dienyl radical. Then oxygen is added to produce a peroxydienyl radical which is subsequently transformed into an anion. Finally the proton which had been abstracted in the first step of the reaction is transferred back to generate a

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lipid hydroperoxide. All reactions are carried out within the enzyme complex and the radical species formed as intermediate products are never set free. The overall reaction can be described as: LH þ O2 ! LOOH Four main enzyme types named for their specificity to oxygenate the substrate molecule at a certain carbon position (for example 12-LOX oxygenates AA at carbon-12) occur in animal tissues, i.e., 5-LOX, 8-LOX, 12-LOX, and 15-LOX, generating 5-, 8-, 12- and 15-hydroperoxyeicosatetraenoic acids (HPETE) which are rapidly reduced to the corresponding hydroxyeicosatetraenoic acids (HETE). In addition to their own biological activities HPETE and HETE are precursors for more complex molecules, e.g., leukotrienes, lipoxins and hepoxilins, some of which are also found in the brain. Three forms of LOX have been found in the brain, i.e., 5-LOX, 12-LOX, and 15-LOX. While AA is the main substrate, derivatives of DHA, which are mostly docosanoids, are also known to be formed. 12-LOX is the most common LOX in the brain and seems to be involved in modulation of synaptic function and longterm potentiation in the central nervous system (Phillis et al., 2006). 5-LOX which is expressed in neurons in various regions of the brain has been widely investigated. The physiological role of neuronal 5-LOX is not yet fully understood, but it seems to be essential for neuronal development. In the aging brain expression of the 5-LOX gene and the activity of the 5-LOX pathway are highly increased hinting at the role of LPO in this process (Manev et al., 2000). LOX can oxidize free PUFA after its release from neural membrane PLs by the action of PLA2, thus initiating the production of various eicosanoids; but it can also react directly with PLs in membranes to produce hydroperoxides that perturb membrane structure. Moreover, if free FA concentrations exceed a certain threshold level, such as following cerebral ischemia, LOX may undergo productcontrolled inhibition. As a consequence, iron may be released from the active site of the enzyme. Thus, free iron ions become available to initiate and propagate nonenzymatic LPO reactions via a one electron pathway leading to free radical chain reactions with damaging consequences (Girotti, 1998).

2.2 Nonenzymatic Lipid Peroxidation The initial products of free radical-catalyzed LPO are mostly conjugated dienic hydroperoxides that can be measured as either conjugates (Esterbauer et al., 1989) or as their stable degradation products (i.e., isoprostanes, MDA and 4-HNE). These nonenzymatic products generated during brain insults are biologically reactive molecules as they can travel within cells and reach peripheral locations distant to the original point of LPO to inflict further damage. It should be noted that at very low concentrations, some of the LPO degradation products may modulate cell functions and act as extracellular or intracellular signalling molecules whereas at high concentrations they may turn into neurotoxic compounds (Farooqui and Horrocks, 2006).

2.2.1 Isoprostanes Isoprostanes are prostaglandin-like compounds formed in vivo from the nonenzymatic, free radicalcatalyzed peroxidation of essential FA by a mechanism analogous to the enzymatic formation of prostaglandins by COX. Isoprostanes are isomeric to prostaglandins differing only in certain stereochemical aspects of the two side chains on the prostane ring. However, in contrast to prostaglandin synthesis which starts from free FAs, isoprostanes are generated from FAs that are esterified to the sn-2 position of PLs and subsequently released by the action of PLA2 (Morrow et al., 1992). Most FAs containing three or more double bonds can serve as the substrate; however, AA containing 4 double bonds is the main precursor for most of the isoprostanes. The most abundant products are isomeric to prostaglandin F2a, hence they are called F2-isoprostanes. Since their discovery in the late eighties (Morrow et al., 1990), they have been established as biomarkers for oxidative damage in an increasing number of diseases. Presently they appear to be the best available biomarker of LPO (Montuschi et al., 2007) (> Figure 25-2).

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. Figure 25-2 F2-Isoprostane

Similar to AA, DHA also undergoes nonenzymatic oxidation giving rise to the group of F4-isoprostanes, also known as neuroprostanes (Roberts et al., 1998). A number of isoprostanes and neuroprostanes have been found to possess biological activities. Many isoprostanes are very potent vasoconstrictors of brain microvasculature and are therefore implicated as mediators of OS injury (Milne et al., 2008). In addition to these derivatives more recently, another group of biologically active compounds, i.e., substituted tetrahydrofuran derivatives called isofurans and neurofurans that are formed via a slightly different pathway of lipid peroxidation were reported (Fessel and Jackson Roberts, 2005; Song et al., 2008) (> Figure 25-3). . Figure 25-3 Neuroprostane

2.2.2 Malondialdehyde MDA is one of the most frequently used indicators for LPO products. MDA concentrations are measured by variations of the 2-thiobarbituric acid reactive substances (TBARS) assays with the most reliable results obtained as MDA-TBA adducts after chromatographic separation, GC/MS or HPLC (Dotan et al., 2004). MDA is a reactive carbonyl compound which primarily attacks proteins containing lysine, arginine, and cysteine residues, which generate advanced lipoxidation end products and protein cross-links adducts. MDA can damage membrane bilayers when reacting with amino-phospholipids (amino-PL) such as ethanolamine phosphoglyceride (EPG) and serine phosphoglyceride (SPG). These chemical modifications result in the disruption of membrane and protein structure and function and enhance cell damage. MDA reactions with nucleic acids will damage DNA and contribute to impaired cellular functions (> Figure 25-4). . Figure 25-4 Malondialdehyde (MDA)

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2.2.3 4-Hydroxy-2-Trans-Nonenal Nonenzymatic peroxidative degradation of AA yields the well studied 4-HNE as a major lipid-derived product which can be easily detected in a variety of biological samples. 4-HNE is a nine-carbon a, b-unsaturated hydroxy-alkenal which contains three reactive groups consisting of an aldehyde, a doublebond on carbon 2 and a hydroxyl group on carbon 4 (> Figure 25-5). . Figure 25-5 4-Hydroxy-2-trans-nonenal (4-HNE)

When present at low concentration in the brain it may modulate cellular signalling (Keller and Mattson, 1998; Leonarduzzi et al., 2004; Uchida, 2003) and its reactivity toward nucleophiles such as Cys, His and Lys residues in proteins has been characterized (Esterbauer et al., 1991). 4-HNE can generate covalent adducts with various primary amine and thiol-containing molecules; it binds to low density lipoproteins (LDL), free amino acids, and nucleotides, thereby disrupting important cellular functions including nucleic acid synthesis. More recently it has been identified as a potential inducer of COX-2 (Uchida, 2008). Nonenzymatic peroxidative lipid degradation of DHA results in the generation of 4-hydroxyhexenal (4-HHE), a six carbon analog of 4-HNE, which in certain tissues (i.e., blood plasma, retina, specific brain regions) is fairly abundant according to some studies (Guichardant et al., 2006). Despite its similar structure 4-HHE reveals specific biological activities and neurotoxic potential that are different from 4-HNE (Long et al., 2008).

3

The Bioavailability of Lipid- and Water-Soluble Antioxidants

There is a general consensus that brain tissue is extremely sensitive to oxidative damage caused by superoxide radicals, hydrogen peroxide and hydroxyl radicals. In order to counteract the harmful consequences resulting from their action, cells normally possess a repertoire of low molecular weight antioxidants (LMWA) (e.g., ascorbic acid, a–tocopherol, and glutathione) which along with several antioxidant enzymes (e.g., SOD, catalase, glutathione peroxidase) are responsible for terminating radical chain reactions. For most of LMWAs, a major prerequisite for action is the close proximity of the antioxidant to the radical in order to trap and neutralize it before it can react with other molecules. The scavenger reacts with the free radical by hydrogen abstraction, forming a new radical of the antioxidant. A compound is able to act as an antioxidant and as a radical scavenger only if the generated antioxidant radicals have a sufficiently long lifetime to react with a second radical by forming a new molecule. For instance, after reducing lipid hydroperoxides to their corresponding alcohols by abstraction of hydrogen from the SH-groups, two glutathione radicals can combine to a stable dimer terminating the radical chain reaction. The antioxidant capacity of the system depends on the activity of the corresponding enzymes to recover the antioxidant molecule, in this case glutathione reductase which reduces glutathione disulfide (GS-SG) back to the sulfhydryl form (GSH). The defense mechanisms to prevent OS comprise of enzymatic systems including SOD, catalase, glutathione peroxidases and thioredoxin system all of which when active, are very effective in handling ROS detoxification. Nonenzymatic endogenous antioxidants on the other hand, encompass an entire spectrum of molecules which can vary in their biophysical properties, mechanism of action, abundance in tissue, source for provision and self regeneration. Within this group are included ascorbic acid, uric acid, glutathione, vitamin E, carotenoids, coenzyme Q, melatonin, melanin, billirubin and lipoic acid.

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This group of LMWAs is extremely diverse and of varying bioavailability as further discussed below. In brain tissue the naturally occurring lipid-soluble antioxidants tocopherols, carotenoids, and lipoic acid are all of dietary origin; only coenzyme Q is endogenously synthesized.

3.1 Dietary Low Molecular Weight Antioxidants Many of the LMWA members are bona fide natural compounds, mostly of plant origin, exhibiting watersoluble or lipid soluble characteristics and are provided by the diet. Among these nonenzymatic micronutrients and compounds bearing antioxidant properties most common are vitamin C (ascorbic acid), vitamin E (a-tocopherol) and carotenoids. Upon oxidation, these micronutrients need to be regenerated, hence the need for further coupling to reducing systems such as glutathione/glutathione disulfide or NADPH/NADP+ and NADH/NAD+. When orally ingested, they may redistribute into tissues according to lypohilicity and hydrophilicity properties. In organs such as the brain, their entrance may be even suppressed as a result of the blood brain barrier (BBB).

3.1.1 Ascorbic Acid L-Ascorbic

acid (Vitamin C) does not cross the BBB unless in its oxidized form as dehydroascorbic acid which enters the brain via the glucose transporter GLUT-1. Dehydroascorbic acid is then reduced to ascorbic acid which is sequestered in the brain at fairly high concentrations. L-Ascorbic acid accumulates in the mitochondria, where most of the free radicals are produced, providing protection to the mitochondrial membrane and the nuclear material. H2O2 or other ROS can oxidize ascorbic acid first to monodehydroascorbic acid and then to dehydroascorbic acid. The oxidized forms of ascorbic acid are relatively stable and nonreactive, do not cause cellular damage and are eventually reduced by appropriate enzymes to restore ascorbic acid levels (> Figure 25-6). . Figure 25-6 Ascorbic acid is oxidized to dehydroascorbic acid

In the presence of transition metals such as iron and copper, ascorbic acid has been shown to possess deleterious pro-oxidant activities. In the presence of oxygen divalent metal ions can catalyze the production of hydrogen peroxide and superoxide via an endiol-like metal-ascorbic acid-dioxygen complex. While undergoing autoxidative destruction, ascorbic acid can promote autoxidation of other macromolecules such as unsaturated lipids. For example, during brain injury when tissue organizational disruption occurs, the iron/ascorbic acid mixture is a highly potent pro-oxidant for brain membranes. Ascorbic acid is a highly hydrophilic compound present overwhelmingly in aqueous compartments of the cytoplasm, plasma and other body fluids; thus it is not immediately responsible for protecting membrane lipids from OS. However, being part of the cellular antioxidant recycling system it can reduce the tocopheroxyl as well as the beta-carotene radicals, thus restoring the antioxidant potential of two important lipophilic antioxidants (Sies et al., 1992; Buettner, 1993).

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3.1.2 Vitamin E Vitamin E is the generic term used to describe a family of at least eight naturally occurring hydrocarbon compounds characterized by a chromanol ring with a hydrophobic phytol side chain referred to as tocopherols and tocotrienols (> Figure 25-7). . Figure 25-7 a-Tocopherol

Vitamin E is a major lipid soluble antioxidant partitioning in cell membranes and lipid containing organelles where it protects membranes and lipoproteins from oxidative stress. (Wang and Quinn, 2000). Among the various vitamin E components, only a-tocopherol is actively taken up by the brain and is directly involved in nervous tissue protection, however the complex mechanisms of transport and regulation of a-tocopherol concentration in the brain are still not entirely elucidated (Spector and Johanson, 2007). a-Tocopherol is a most effective protectant against lipid peroxyl radicals acting in the membrane domain. Its hydroxyl group can easily donate a hydrogen atom to reduce a free radical; hence, it is able to break peroxyl chain propagation reactions. The resulting a-tocopheroxyl radical formed, is stabilized by delocalization of the unpaired electron about the fully substituted chromanol ring system rendering the molecule relatively unreactive. Tocopherols can be regenerated from the tocopheroxyl radicals in a redox cycle involving other endogenous antioxidants.

3.1.3 Carotenoids The antioxidant properties of carotenoids, notably beta-carotene and lycopene as well as oxycarotenoids (e.g., zeaxanthin and lutein), are associated with their radical scavenging properties and their exceptional singlet oxygen quenching abilities. Carotenoids are hydrophobic molecules and hence they are found predominantly in lipophilic regions, e.g., cell membranes, but their value as antioxidants being able to prevent membrane lipid peroxidation remains controversial (> Figures 25-8 and > 25-9). . Figure 25-8 b-Carotene

Carotenoids may lose their antioxidant effect or even turn into pro-oxidants at high oxygen concentrations and high concentration of the carotenoid itself. However, a synergistic protection seems to exist in combination with other antioxidants. It has been shown that vitamin C can repair the beta-carotene radical and hence restore its antioxidant potential (El-Agamey et al., 2004).

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. Figure 25-9 Lycopene

Another dietary constituent that crosses the BBB is lycopene. It is present in the brain at low concentrations and lycopene supplementation has recently been suggested to be potentially useful in the management of neurodegenerative diseases (Rao et al., 2006).

3.1.4 Lipoic Acid Alpha-lipoic acid is a lipid-soluble dithiol that is derived from the diet, crosses the BBB and is readily taken up by brain cells (> Figure 25-10). . Figure 25-10 Lipoic acid

Lipoic acid is not only an essential cofactor for many enzyme complexes (the most prominent being perhaps, the pyruvate dehydrogenase complex), but also acts as a chelator for transition metal ions (e.g., iron). Its ability to scavenge various ROS (e.g., peroxydienyl radicals, hydroxyl radicals and hydrogen peroxide), nitric oxide (NO) and peroxynitrite species makes it a powerful antioxidant. Moreover, in its reduced form as dihydrolipoic acid it is able to regenerate (reduce) antioxidants, such as glutathione, vitamin C and vitamin E and if it leaves the cell it acts as an extracellular antioxidant (Packer et al., 1995). Administration of lipoic acid offers some hope for treatment of neurodegenerative disorders (Packer et al., 1997; Holmquist et al., 2007).

3.2 Endogenous Low Molecular Weight Antioxidants The brain employs a fairly limited number of water-soluble and lipid-soluble LMWA in order to protect its highly sensitive membranes from OS. In addition, a few highly specific, lipid-soluble LMWAs are locally synthesized to assist in combating excess free radicals formation.

3.2.1 The Glutathione Cycle Glutathione (GSH) is a ubiquitous water-soluble thiol-containing tripeptide which plays an important role as co-factor in various enzymatic reactions and is considered one of the most important antioxidants in the brain. It is not an essential nutrient because it is endogenously biosynthesized from cysteine, glutamate and glycine, mainly in astrocytes. As the major thiol-disulfide redox buffer, glutathione is essential for the detoxification of ROS in the brain. The normal intracellular concentration of GSH is in the millimolar whereas the plasma concentration is in the micromolar range. Glutathione is found almost exclusively in its reduced form, since the enzyme

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that recovers it from its oxidized form (GSSG), glutathione reductase (GR), is constitutively active and inducible upon OS. In healthy cells and tissues, more than 90% of the total glutathione is present in the reduced form and less than 10% exists in the disulfide form. An increased GSSG-to-GSH ratio is considered indicative of OS (> Figure 25-11). . Figure 25-11 Glutathione

The glutathione peroxidases (GPx) are selenocysteine-containing proteins and constitute the most important antioxidant enzymes in the brain to prevent LPO-induced damage. GPx catalyze (1) the reduction of free hydrogen peroxide to water and (2) the reduction of lipid hydroperoxides to their corresponding alcohols: 2GSH þ H2 O2 ! GS  SG þ 2H2 O

ð1Þ

2GSH þ L  O  OH ! GS  SG þ L  OH þ H2 O

ð2Þ

After reducing a lipid hydroperoxide to its corresponding alcohol by abstraction of hydrogen from the SH-groups, two glutathione radicals can combine to a stable dimer. The antioxidant capacity of the system depends on the activity of the corresponding enzymes to recover the antioxidant molecule, in this case glutathione reductase which reduces glutathione disulfide (GSSG) back to the sulfhydryl form (GSH): GSSG þ NADPH þ Hþ ! 2 GSH þ NADPþ For efficient glutathione-dependent reduction of peroxides, neural cells contain glutathione in high concentration and have substantial activity of glutathione peroxidase, glutathione reductase and enzymes that supply the NADPH required for the glutathione reductase reaction (Savaskan et al., 2007a; Dringen and Hirrlinger, 2003; Dringen et al., 2005). In general, intracellular glutathione peroxidases comprise two distinct proteins, the classical GPx (cGPx) and the phospholipid hydroperoxide glutathione peroxidase (PHGPx), the latter of which is present in the nucleus, mitochondria and cytosol. PHGPx, also known as GPx4, is the only member of the GPx family that can use native PL hydroperoxide as substrate, while the others can only catalyze the reduction of free FA hydroperoxide (Imai and Nakagawa, 2003). Cytosolic and mitochondrial GPx4 are the major isoforms exclusively expressed by neurons in the developing brain. In contrast, following brain trauma, GPx4 is specifically upregulated in nonneuronal cells, i.e., reactive astrocytes (Savaskan et al., 2007b).

3.2.2 Coenzyme Q Ubiquinones, which are also known as coenzyme Q (CoQ), act as both lipophilic antioxidants protecting membrane lipids from oxidation as well as pro-oxidants or electron carriers that facilitate the mitochondrial respiratory chain. Being very hydrophobic molecules, ubiquinons are located within the membrane bilayer. CoQ is the only lipid-soluble antioxidant synthesized endogenously. CoQ is present in all cells and membranes, but its concentration varies greatly not only among different organs but also among various cells and regions of the same organ (e.g., different brain structures), and among different subcellular organelles (Turunen et al., 2004).

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Mitochondrial ubiquinones are essential components of the electron transport system, but they may also act as pro-oxidants as they are involved in the production of ROS (i.e., O2) which might induce oxidative damage. The most common CoQ in human mitochondria is Q-10. In its reduced form, as ubiquinol (CoQH2), it is an important antioxidant protecting membrane PLs against peroxidation. Its antioxidant potential includes (1) prevention of peroxydienyl radical production during initiation (2) its direct action as a chain breaking agent and (3) its involvement in the recycling of oxidized, radical forms of vitamin C and E, hence restoring their antioxidant capacities (James et al., 2004; Bentinger et al., 2007). The biosynthesis of CoQ was studied in great detail in bacteria and yeast and less in animal tissues (Turunen et al., 2004). However, during aging and under pathophysiological conditions tissue concentrations of CoQ are modified and Q-10 supplements have been shown to be beneficial in animal studies as well as in clinical trials suggesting that it might protect from neuronal damage induced by ischemia or neurodegenerative disorders and aging (Dhanasekaran and Ren, 2005; Galpern and Cudkowicz, 2007; Young et al., 2007).

3.2.3 Membrane-Derived Antioxidants: The Case of the Vinyl Ether Bond As pointed out in the previous section, the possible harmful consequences of ROS and reactive nitrogen species under physiological conditions are counteracted by an arsenal of defence mechanisms spanning from gene-regulated enzymatic detoxifications, to powerful nonenzymatic LMWA which exert a scavenging and chain propagation blockade. When the oxidation process by ROS is initiated in the aqueous cellular compartment, typical water-soluble antioxidants such as glutathione and ascorbic acid or water-soluble enzymes are the main contenders for the antioxidative action. With regard to the membrane compartment, while membrane-associated GPx4 is a major player, other LMWA such as a-tocopherol, coenzyme Q, b-carotene and lipoic acid take part in the detoxification process. However, not all this arsenal is available at the same time within the same cell or tissue and in sufficient concentrations to combat excess production of free radicals. Not surprisingly, a diverse spectrum of different lipid oxidation products is generated even under physiological conditions. In brain tissue, the generation of counteracting free radicals is of particular concern because of the high content of PUFA in the membrane that may become a target for lipid oxidation. Moreover, cells such as neurons bear a greater risk for free radical targeting because of a very high membrane surface/cytosol ratio (i.e., dendritic tree). Equally disadvantageous is the fact that neurons possess rather limited antioxidant defence mechanisms. Because of their firing activity neurons are by far the main consumers of the oxygen. Therefore studies of the nervous system have searched for potential endogenous antioxidants which specifically are integral parts of the hydrophobic domain where most double-bond targets for hydroxyl radicals are present. 1, 2-diacyl EPG and 1-alkenyl 2-acyl EPG (pEPG, plasmalogen) molecular species appear to fulfil this requirement and they also comprise a most robust and metabolically active reservoir. Basically pEPG consist of an aldehyde in the sn-1 position giving rise to a single vinyl ether double-bond and a long-chain hydrocarbon containing a carboxyl group esterified to the sn-2 position of the glycerol moiety possessing anywhere between 2–6 double-bonds. It is the interplay between these two side chains that creates a powerful antioxidant system that resides in the hydrophobic domain to scavenge free radicals. Plasmalogens have been proposed as natural antioxidants following OS insults based on their ability to scavenge ROS, such as peroxyl radicals and singlet oxygen, as well as to chelate potentially harmful metal ions (Zoeller et al., 1998; Zoeller et al., 1999; Leray et al., 2002; Spiteller, 2006). Furthermore, oxidation of the vinyl ether bond was suggested to prevent oxidation of adjacent PUFAs, thus keeping PUFAs untouchable from the consequences of free radical attacks and the resulting epoxide being a relatively stable molecule (Zommara et al., 1995; Engelmann, 2004) (> Figure 25-12). However, the hypothesis that degradation products of pEPG do not propagate LPO has been questioned since pEPG epoxides may give rise to highly reactive a-hydroxyaldehyde species (Spiteller, 2006). This appeared to be the case at least, in rat brain homogenates after UV- and Fe2+/ascorbate-induced oxidation (Stadelmann-Ingrand et al., 2004). Independently, it has been shown that a-hydroxyaldehydes

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. Figure 25-12 The structure of 1,2-diacyl (top) and 1-alkenyl,2-acyl (bottom) ethanolamine phosphoglycerides. EPG, ethanolamine phosphoglyceride; pEPG, plasmalogen EPG

which are considered as powerful nucleophilic reagents which may attack other biomolecules including proteins are dramatically increased during aging. Plasmalogens are ubiquitous constituents of brain tissue (Diagne et al., 1984) and contain exclusively the ethanolamine N-base unlike peripheral tissues such as heart and kidney which predominantly contain choline (Kikuchi et al., 1999; Farooqui et al., 2003). Brain pEPG are also exclusively enriched with DHA of the n-3 series and less with AA of the n-6 series while cardiac and most peripheral tissues do contain PUFA of the n-6 series. More recently it has been reported that LPO products such as hydroxy-alkenals covalently bind to the N-base of EPG in biological membranes. While 4-HNE constituted the major precursor for EPG-alkenal Michael adducts in blood platelets, EPG-4-HHE resulting from n-3 PUFA peroxidation were the main Michael adducts in the retina of diabetic rats, a tissue which is especially rich in DHA (Bacot et al., 2007). Morever, pEPG species were found to be significantly more reactive toward the hydroxy-alkenals than the 1–2-diacyl EPG species and DHA-containing EPG-species were more reactive than AA-containing species. Based on these findings the authors suggested that in addition to their antioxidant potential, pEPG might be actively involved in detoxifying cells of hydroxy-alkenals reactants after OS (Guichardant et al., 2006).

3.2.4 Membrane-Derived Antioxidants: The Case of DHA Of the several unsaturated FA esterified to the sn-2 position of the glycerol backbone of pEPG, DHA is the most prominent long-chain PUFA. The prominence of DHA in brain tissue however is even more striking, since it is characteristic of all amino-PL (i.e., 1, 2-diacyl EPG and 1, 2-diacyl SPG), independent of the substituent in the sn-1 position. Furthermore, these particular PL molecular species appear to reside predominantly in the cytofacial site of the plasma membrane (Yavin and Brand, 2005). Being the most abundant and highly unsaturated PUFA in the mammalian brain, DHA would be a prime target for OS and therefore a major source for LPO products. Dietary supplements of DHA enhance

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the endogenous esterified DHA pool, which in turn, would increase the risk for OS. Yet, most of the experimental evidence linking brain DHA levels to peroxidative events does not bear out this prediction. On the contrary, increased levels of DHA in the brain might confer protection against OS as illustrated by a decrease in LPO by serving as a preferred target for free radical attack (Kubo et al., 2000; Yavin et al., 2002). The purported dual pro-oxidant and antioxidant activity of DHA has been proposed based on several observations: (1) DHA hydroperoxides and further degradation products have been identified in vivo. (2) From a biophysical point of view, entrapment of the free radical on the nascent PL can be justified based on the very high intrinsic lateral mobility and expanded motional properties of DHA in the bilayer which may accelerate free radical penetration and oxidation (Binder and Gawrisch, 2001; Stillwell and Wassall, 2003) (3). The ethanolamine rather than the choline residue expresses direct scavenging activity for superoxide anions (Gordon et al., 1991). (4) The ethanolamine PHG is mainly present in the cytofacial site of the cell membrane; thus the chances for interaction of intracellular free radicals with EPG and pEPG are increased. (5) An oxidized DHA molecule can be enzymatically removed from the existing PL through a deacylationreacylation cycle and further cleared by peroxisomal and mitochondrial FA b-oxidation, while the lysophospholipid is reacylated with an appropriate PUFA (Farooqui et al., 2000). The deacylation-reacylation cycle is a very effective repair mechanism which can re-introduce proper PUFAs into neural membrane PL much like the repair mechanisms existing for oxidized polynucleotides such as RNA and DNA molecules. A failure to provide the appropriate PUFA or a less efficient reacylation may be reasoned as a cause for the losses of EPG and pEPG molecular species in the aging and pathological brain. In conclusion, the cascade of events which leads to partial EPG and pEPG degradation following brain injury has been summarized in the > Scheme 25-1 outlined below. Following Ca2+ influx and accumulation of free hydroxyl radicals, the EPG molecule is either hydrolyzed by a PLA2 or oxidized to lipid hydroperoxide on the DHA side chain. The free DHA can undergo nonenzymatic oxidation or LOX/COX-mediated conversion into a series of derivatives that appear to impart neuroprotection. The lipid hydroperoxide can either be reduced to a fatty acyl alcohol (LOH) by a direct action of GPx4 on the EPG molecule (Ursini et al., 1991; Imai and Nakagawa, 2003) followed by subsequent excision to give rise to a 22-carbon long free acyl alcohol. Alternatively, it can first be excised by a PLA2 and then reduced by a GPx (Grossman and Wendel, 1983). In either alternative, the nontoxic free carboxyl alcohols can be further degraded by b-oxidation which is essentially a neuroprotective pathway. The resulting lyso-EPG or lyso-pEPG can then be rapidly esterified by an acyl-CoA transferase to restore the native EPG molecular species.

4

Biological Consequences of Lipid Peroxides Action

In most circumstances while lipid-derived hydroperoxides are subject to reductive degradation, the cytopathological or cytoprotective consequences of the signalling generated may largely depend on the local environment, i.e., site of action. The generated products may trigger various signal transduction pathways which can accelerate up-regulation of detoxifying enzymes or activate cell death kinase cascades. The following section will address the consequences of some actions of lipid peroxides where the site of action is directed at the cell membrane level and will also explore a second site of action, where the brain acts as an organ while it undergoes marked structural/ functional changes during development or aging.

4.1 On Scrambling of Phospholipid Asymmetry The asymmetric distribution of PL based on polar head groups (PHG) composition across the plasma membrane bilayer, originally reported almost four decades ago (Bretscher, 1971) is ubiquitous for most mammalian cell membranes and plays essential roles in cell physiology and function (Rothman and Lenard, 1977; Daleke, 2003; Yavin and Brand, 2005; van Meer et al., 2008). Under normal conditions choline phosphoglyceride (CPG) and sphingomyelin (SMy) are the dominant constituents in the outer leaflet of the

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. Scheme 25-1 The metabolic course of EPG (pEPG) degradation products during OS. For details see text. Abbreviations: COX, cyclooxygenase; Ea, ethanolamine; GSH, glutathione; GPx4, phospholipid hydroperoxide glutathione peroxidase; LOX, lipoxygenase; PLA2, phospholipase A2

plasma membrane, while amino-PL, i.e., EPG and SPG, as well as inositol phosphoglycerides (IPG), are mainly present in the plasma membrane’s inner leaflet. The origin and maintenance of the PL transbilayer distribution is determined by the site of PL biosynthesis and is facilitated by a number of specific protein transporters. A balance of the activities of three enzymes maintains the typical asymmetry of plasma membrane lipids in mammalian cells. These are (a) the aminophospholipid translocase (APT) or ‘flippase’, an ATP-dependent, inward-directed pump, specific for amino-PL; (b) an outward-directed pump known as ‘floppase’ which shows little selectivity for the PHG of the PL; and (c) a PL scramblase, which facilitates bi-directional translocation of all PL classes independent of the PHG (Devaux et al., 2006). Activation or inhibition of any one of these enzymes may lead to a scrambling in membrane asymmetry. While there is a general consensus that PUFA are preferred targets for LPO because of their multiplicity of double-bonds, the question about targeting specific PUFA species by free radicals depending on their PHG profile has not been answered, yet. However, in the brains of subjects with AD, a condition associated with increased OS, membrane asymmetry was found to be markedly scrambled, as specifically inner membrane leaflet PL such as EPG, IPG and pEPG were significantly decreased (Prasad et al., 1998; Guan et al., 1999). Often OS may cause changes in membrane asymmetry by reshuffling the bilayer distribution of SPG and EPG amino-PL both of which are enriched in DHA. Translocation of SPG to the outer membrane leaflet is observed under pathophysiological conditions during early stages of apoptotic cell

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death (Hanshaw and Smith, 2005; Zwaal et al., 2005). More recently, it was reported that not only SPG but also EPG is redistributed on the cell surface during apoptosis, resulting in a total loss of the asymmetric distribution of amino-PL in the plasma membrane (Balasubramanian and Schroit, 2003; Yavin and Brand, 2005). Oxidation of long-chain PUFAs and generation of reactive aldehydes predominantly from AA but also from DHA may cause changes in lipid asymmetry. These changes may serve as a trigger for modulation of intracellular signalling and induce functional damage through alterations of the normal lipid environment of integral proteins. One such immediate damage has been observed in APT: the transmembrane protein which catalyzes the unidirectional transport of EPG and SPG from the outer to inner monolayer with a halflife of translocation of 5–10 min. In cortical synaptosomal preparations, APT was found to be inactivated by two prominent LPO products, i.e., 4-HNE and acrolein, presumably due to covalent modification of a critical cysteine residue (Castegna et al., 2004). In neuronal as well as glial cells, the activity of APT was significantly decreased during apoptosis following a hypoxic insult, strongly suggesting that this inhibition could be the main reason for the loss of amino-PL asymmetry in apoptotic brain cells (Das et al., 2003). Recently, APT inactivation by LPO products has been suggested to be responsible for scrambled SPG asymmetry in synaptosomes from the brain of subjects with AD and mild cognitive impairment (MCI) (Bader Lange et al., 2008). A second transmembrane protein affected by OS is the phospholipid scramblase 1 (PLSCR1). There are several scramblases which appear inherently nonspecific and their role is to randomize the distribution of newly synthesized lipids in the endoplasmic reticulum or plasma membrane lipids in activated cells (Daleke, 2003). PLSCR1 is an endofacial-oriented plasma membrane protein that has been proposed to mediate migration of membrane PLs in the presence of Ca2+ or under acidic conditions (Smeets et al., 1994). Under nonpathological conditions, translocation of EPG from the inner leaflet to the outer leaflet, an essential feature to enable cytokinesis, is mainly regulated by PLSCR1 (Emoto and Umeda, 2001). PLSCR1 was suggested to play an important role in ischemic injury and apoptotic cell death in the human hippocampus presumably by remodelling plasma membrane PLs (Rami et al., 2003). A third transmembrane protein affected by OS and by alterations in lipid asymmetry is the diacylglycerol kinase (DGK), an enzyme responsible for generation of two important lipid messengers in the membrane, i.e., diacylglycerol and phosphatidic acid. Studies have shown that DGK activity is enhanced in EPG-enriched synthetic liposomes suggesting that the lipid composition of the membrane may modulate the activity and membrane translocation of this enzyme (Fanani et al., 2004). A fourth notable protein is the phosphatidylethanolamine-binding protein (PEBP) also termed Raf-1 kinase interacting protein. As the prototype of a novel family of serine protease inhibitors, it has been shown to exhibit specific EPG binding domains. PEBPs represent a family of enzymes implicated in the control of several signalling pathways playing a role in cell proliferation, differentiation and apoptosis (Yeung et al., 1999; Hengst et al., 2001; Odabaei et al., 2004). In vitro experiments have shown, that human PEBP4 can prevent apoptosis in cell culture by inhibiting mitogen-activated protein (MAP) kinase signalling pathways and blocking membrane translocation of EPG by binding to it via its specific EPG binding domain (Wang et al., 2004). The interplay between MAP kinase signalling and EPG externalization with respect to membrane DHA enrichment or deficiency has been studied in details in a clonal cell line of oligodendroglia origin, OLN93. Like most cell lines kept under standard culture conditions, OLN 93 cells are very poor in DHA and moderate OS conditions such as exposure to H2O2 or divalent iron neither induced significant LPO nor caused cell death (Brand et al., 2000). This picture changed completely after cells membrane DHA levels, particularly enriched in EPG and SPG species, were partialy restored (increase in DHA content between six- and eightfold following 3 days supplement). Upon stress DHA-rich cells showed extracellular signal-regulated protein kinase (ERK) activation and nuclear translocation as well as a rapid and transient EPG membrane translocation followed by a delayed irreversible cell damage and apoptotic death (Brand et al., 2001). Blocking the ERK signaling pathway by the use of specific inhibitors prevented EPG externalization and cell death, indicating an intimate relation between the two events (Brand and Yavin, 2005). A different approach to modulate apoptotic signalling was carried out by preventing EPG translocation through substitution of membrane EPG by its PHG analogs, N-monomethyl- or N,N-dimethyl-EPG

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(dEPG). N,N-dimethylethanolamine supplements resulted in the formation of a newly synthesized PL species, i.e., dEPG, at the expense of existing PL such as EPG and CPG. With dEPG comprising about 50% of the total membrane PL and EPG reduced by half, OLN93 cells became resistant to OS. DHA-enrichment was now evident in the newly formed dEPG rather than EPG species. Even though DHA-derived LPO products were still found following OS, EPG translocation did not take place, ERK nuclear translocation was inhibited, and cells were saved from undergoing cell death (Brand and Yavin, 2001). These studies suggest that DHA-enriched-EPG translocation is a major consequence and a necessary signal for the apoptotic cell death cascade and intimately linked to MAP kinase signaling. In sum, altering the topology of the plasma membrane by EPG reorientation could be a way of controlling or triggering specific enzymes and signalling cascades that determine cell fate. Most recently cardiolipin (CL), a mitochondria-specific PL has attracted much interest for its possible involvement in apoptotic signalling (Gonzalvez and Gottlieb, 2007; Orrenius et al., 2007). CL is a glycerolbased PL which is found in the mitochondrial inner membrane where it interacts with the electron transport chain complexes involved in oxidative phosphorylation. Moreover, CL is closely associated with cytochrome c at the outer leaflet of the mitochondrial inner membrane. Release of cytochrome c from the mitochondria into the cytosol is a pro-apoptotic signal and a major factor in caspase activation. Two CL binding sites on cytochrome c have been proposed; one which facilitates electrostatic interactions with the negative charges of the CL head group and another which is involved in hydrophobic interactions with the fatty acyl chains. Oxidation of CL reduces cytochrome c binding and increases the level of soluble cytochrome c in the intermembrane space. Notably, CL-bound cytochrome c itself exhibits peroxidase activity and catalyzes CL peroxidation. Using advanced lipidomics techniques it was recently demonstrated that during the early phase of apoptosis induced by traumatic brain injury CL is the preferred target for LPO while more abundant PLs such as CPG and EPG remain nonoxidized (Bayir et al., 2007). However, the accumulation of CL hydroperoxides activates the release of cytochrome c and other proapoptotic factors from the mitochondria into the cytosol (Kagan et al., 2005). Interestingly, in brain tissue the distribution of aliphatic chains in neuronal CL is complex and distinctively different from that in other organs (e.g., heart, liver, etc.). While in nonneural tissues CL molecular species predominantly contain only one major type of aliphatic chain, i.e., linoleic acid, it was recently reported that the neuronal CL profile contains a very complex repertoire of CL molecular species, in which AA and DHA are markedly enriched (Cheng et al., 2008).

4.2 On the Perinatal Brain 4.2.1 Brain Ontogeny During Adequate or Adverse Environments Adequate foetal growth is a result of complex interactions between maternal supplies and the capacity of the placenta to both transfer and generate essential metabolites and growth factors to the growing foetus. Inadequate maternal nutritional supply is considered an environmental epigenetic stress, which might induce both brain and body growth retardation. The principal reason for brain vulnerability during intrauterine life is disruption of an otherwise perfectly timed and spatially-confined sequence of events leading to the cyto-architectural organization of neuronal and glial cells into functional networks (Dobbing and Sands, 1979). Apart from various neurotrophic factors (i.e., BDNF, NGF and EGF) brain ontogeny requires the presence of discrete molecules, such as low molecular weight neurotransmitters and neuromodulators, including lipid derivatives, as well as high molecular weight matrix and membrane specific proteins to maintain the delicate balance between cell differentiation and cell death by apoptosis (Buss and Oppenheim, 2004). Inadequate maternal nutritional provision may in addition, compromise brain ontogeny because of reduced levels of essential nutrients such as PUFA, the two most prominent of which, DHA and AA, could alter the normal course of neurogenesis and cerebral angiogenesis. Indeed, a dramatic accretion of both DHA and AA in the human brain, peaks during the last trimester of pregnancy (Clandinin et al., 1980), but

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continuous requirements are deemed throughout gestation including early periods between the 8th to the 16th weeks which is the hallmark of neurogenesis. In that respect accretion of DHA is of paramount importance as it is needed for dendrites and synapse formation. A deprivation of adequate nutritional constituents, particularly micronutrients, along with lesser developed antioxidant enzyme systems and immature cerebral blood flow autoregulation, could also compromise the foetal antioxidant defence system rendering the brain susceptible to OS insults.

4.2.2 Oxygen Deprivation and Its Adverse Outcome In addition to maternal nutritional deprivation, adverse environmental conditions such as birth asphyxia or prenatal hypoxic/ischemic episodes of placental vascular origin (Burke et al., 2006) are believed to be harmful stressors, likely to affect brain ontogeny. A great deal of information gathered mostly in the adult brain, has indicated that following oxygen depletion, ATP synthesis is markedly reduced and the Na+/K+ pump is unable to maintain cellular ionic gradients. In the absence of a membrane potential, large amounts of calcium ions enter through voltage-dependent ion channels into the cell. An excessive increase in levels of intracellular calcium leads to cell damage through the activation of a variety of hydrolases, such as proteases, phospholipases and endonucleases. During hypoxic/ischemic injury, apart from influx of calcium ions via voltage-gated channels, additional calcium enters the cells through glutamate-regulated ion channels. Glutamate, an excitatory neurotransmitter, is released from presynaptic vesicles during ischemia following anoxic cell depolarization. The acute depletion of cellular energy arising during ischemia induces an almost complete inhibition of cerebral protein biosynthesis which is an early indicator for possible apoptotic neuronal cell death. A second wave of neuronal cell damage takes place during the reperfusion period when flow is restored. This process while peripheral to the focal hypoxic/anoxic stress is confined to a significantly larger area, also called penumbra (Siesjo, 1993). The causes for cell damage characteristic to this area have been attributed primarily to a postischemic release of oxygen free radicals, excess production of nitric oxide (NO), initiation of inflammatory reactions involving cytokines and a disrupted equilibrium of the excitatory and inhibitory neurotransmitter systems (Berger and Garnier, 2000). Most of these signalling cascades are also present in the foetal brain (Perlman, 2006) although because of a lower pressure of oxygen during intrauterine life a limited sparing of the foetus and the neonate has been indicated (Fewell, 2005). Nevertheless, evidence from experimental animal models raises the possibility that a prenatal hypoxic/ ischemic insult may constitute the trigger for several neurodevelopmental disorders characterized by enhanced seizure activity, motor impairment and learning disabilities. These prenatal insults may give rise to severe postnatal pathologies such as cerebral palsy (CP), autism, attention deficit hyperactive disorder (ADHD) and even schizophrenia (Boksa, 2004). To this list, injury of the cerebral white matter in the neonate (Khwaja and Volpe, 2008) resulting from a higher vulnerability of immature oligodendrocytes to free radicals damage (Back et al., 2002) may be added.

4.2.3 DHA: Purported Mechanisms in Neuroprotection In a rat model of intrauterine growth retardation, administration of the ethyl ester of DHA to foetuses via the amniotic route reduced LPO production in the foetal brain after a transient restriction of the placental blood flow (Glozman et al., 1998). Furthermore, a marked decrease of DHA content in brain PLs by prior depletion of maternal supplies of n-3 a-linolenic acid enhanced LPO formation ex vivo using foetal brain slices. The intrauterine ischemic insult also caused reduction of the foetal body weight, as well as brain weight, in spite of a partial sparing of the latter (Yavin, 2006). Based on the premises of ubiquity and near absolute compartmentalization in the membrane lipid bilayer, it was postulated that DHA is a most resourceful antioxidant in the foetal brain and in this capacity may even surpass vitamin E or other water-soluble micronutrients. The antioxidant protective effect of DHA originally identified by the ability to reduce production of MDA has been attributed to its enhanced free radical scavenging capacity as attested by electron spin resonance spectroscopy (Green et al., 2001a, b).

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Lipid extracts from control brains showed little hydroxyl radical scavenging activity, in contrast to those from ethyl-DHA injected foetuses. The latter exhibited a robust scavenging trapping capacity that resided mostly in the DHA-enriched PL fraction as attested by a sharp decrease in the amount of 5,50 -dimethyl-1pyroline-N-oxide-OH (DMPO) adducts formation (Green et al., 2001b). In addition to this direct scavenging activity, a single intra amniotic injection of ethyl-DHA also enhanced production of prostaglandins and NO, both compounds reflecting enhanced enzymatic activities of COX and nitric oxide synthase (NOS), respectively. As for the former enzyme, a decrease in LPO was prevented by pretreatment of the brain slices with indomethacin, a classical COX inhibitor. That raised the possibility that activation of COX following nutritional supplements of DHA may have diverted AA oxidation from nonenzymatic ROS-generating pathways to prostanoids formation. With regard to the possible enzymatic activation of NOS, administration of ethyl-DHA to the foetuses increased NO production while reducing MDA products, an effect that could be mimicked by NO donors. The relationship between n-3 PUFA supplements and NO production or NOS activation has been the subject of ample studies in which animals whether fed or deprived of n-3 PUFA were examined. Several mechanisms have been proposed to explain the purported antioxidant effect of NO. From biophysical measurements using synthetic liposomes, it appeared that NO inhibited LPO by annihilating lipid radical species, thus terminating radical chain propagation processes (Rubbo et al., 1994). Basically NO inhibited production of MDA and transformed lipid peroxides and other MDA precursors into species no longer capable of generating MDA and reactive aldehydes by oxidative breakdown, thus conferring protection (d’Ischia et al., 2000). While these mechanisms are based on the assumption of a direct effect of NO on LPO generation, it is also clear that NO may activate COX and increase PGE2 production (Franchi et al., 1994). In this context a simplified model was proposed to account for the decreased LPO observed in the foetal rat brain after dietary DHA enrichment (Green et al., 2001a) (> Scheme 25-2). . Scheme 25-2 Oxygen utilization via reactive oxygen species (ROS), prostanoids (PN) and Nitric oxide (NO) derivatives formation and the potential role of DHA or DHA* derivatives for activation of COX and NOS enzymes

According to this scheme the flow of oxygen in a living cell is basically determined by three major pathways two of which are catalyzed by NOS and COX. Both enzymes are present as inactive (i) or active (a) forms and are regulated either by non-PL bound DHA or by one of its oxidized (DHA*) derivatives (Serhan, 2005). As a result of COX and NOS enzymes activation, a great part of free AA and arginine (Arg) are committed for increased production of PGE2 and NO, respectively. An increase in COX activity could shift most AA substrate away from a nonenzymatic reaction with ROS species and therefore cause a decrease in LPO and reactive aldehydes production. Activation of NO pathway may also decrease LPO production both directly through free radical termination, or indirectly, by COX activation. PGE2 on the other hand, has been shown to increase inducible NOS (iNOS) activity, thus contributing further to a decrease in LPO products. Since both PN and NO metabolic routes are enzyme-mediated by COX and LOX, AA targeting by free radicals is diminished. An alternative, purported way, by which sustained dietary supplements of DHA may act, is through an increase in the antioxidant capacity of the foetal brain following over expression of selected genes and their posttranslational products (Kitajka et al., 2004). EPA and DHA for example have been shown to activate NF-E2-related factor 2, (Nrf2), a master transcription factor known to regulate expression of more than 200 genes, including those involved in Phase II detoxification and antioxidant gene expression presumably by formation of J3-isoprostanes from oxidation of n-3 PUFA (Gao et al., 2007; Li et al., 2007).

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Finally, it can be argued that the beneficial effect of DHA may arise from an increased free radical trapping capacity due to biophysical changes in the bilayer properties of the PL esterified DHA molecule. Indeed compelling evidence exists for direct targeting of DHA by free radicals, as a result of which an overall decrease of EPG species was noticed (Yoshida et al., 1980). pEPG species may also be subject to direct targeting of free radicals not only because of the abundance of DHA but also because of the targeting of the vinyl ether bond (Sindelar et al., 1999; Hahnel et al., 1999). Notably, in the neonatal brain some 40% of the total aminoethanol PLs are present as pEPG.

4.3 Oxidative Stress in the Aging Brain There is a large consensus that aging is a progressive, nonescapable biological process partially related to the perpetual accumulation of oxidative damage in the organism. Oxidative damage is found in proteins, membrane lipids, nucleic acids, and glycoconjugate components of the supporting extracellular matrix. It also affects the structure/function relationship of subcellular organelles such as nucleus, plasma membrane and mitochondria. The latter are thought to play a crucial role in the aging process not only due to their role as main intracellular generators of ROS, but also because they are targets of ROS attack (Orth and Schapira, 2001). Impaired mitochondrial function may affect ATP-driven transport systems and ion pumps including the Na+-K+-ATPase, Ca2+-ATPase and the Na+-Ca2+ exchanger and ultimately cause cell damage and dysfunction (Mariani et al., 2005).

4.3.1 Progressive Aging Unlike most peripheral organs, the brain is subject to a cumulative burden of OS. While this process is presumably a universal feature of aging, the brain seems to be at higher risk as reasoned above. During normal aging the brain is subject to morphological and functional modifications affecting the dendritic tree, the synaptic structure/function relationship, the neurotransmission apparatus and the effective crosstalk between neurons and glial cell elements. A decrease in calcium homeostasis and a decrement of the neurotransmitter apparatus efficiency are the most characteristic neuronal changes during aging (Thibault et al., 2007) that are likely to lead to alterations in motor and sensory systems, and in humans, to losses of higher brain functions. Part of the age-related deficits in brain function may also result from a decline in the efficiency of normal antioxidant defence mechanisms such as decreased enzymatic activities and lower LMWA levels (Kitani et al., 2003). A decline in cerebral blood flow can also contribute to the aging process (Spilt et al., 2005). Of the main products of LPO, reactive aldehydes species (i.e., MDA and 4-HNE) have received increasing attention as they appear to be prominent biomarkers identified in the aging brain. 4-HNE in particular, an aldehyde derived from oxidation of linoleic acid and AA is a most powerful endogenous neurotoxin (Kruman et al., 1997) which may accumulate in membranes at variable concentrations (up to 5 mM) in response to oxidative insults (Esterbauer et al., 1991) and is largely responsible for cytopathological effects observed during OS in vivo (Uchida, 2003). 4-HNE inhibits neurite outgrowth, disrupts neuronal cytoskeleton and alters cellular tubulin in nerve cells. Disruption of microtubules integrity occurs via Michael adducts. Interestingly these molecular events were not accompanied by cytotoxicity (Neely et al., 1999) possibly indicating that longer time periods are required to activate cell-death cascades. Other reactive aldehydes including neuroketals, 4-hydroxy-trans-2-cis-6-nonadienal and 4-HHE have been identified as DHA-derived products (Van Kuijk et al., 1990) yet neither their impact as endogenous neurotoxins nor the signalling cascades they activate have been well established. In general, 4-HHE appears in quantities that are substantially lower than those found for 4-HNE (Yamada et al., 2004) although in DHA-rich tissues such as retina, 4-HHE constitutes the main LPO product (Guichardant et al., 2006). Finally, progressive aging is not a generalized phenomenon and even long-lived cells such as neurons may follow different courses depending on environmental conditions. Thus, emerging data from recent

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studies indicate that regional stability and plasticity can be contained in many brain regions of the aged brain and neuronal losses may be compensated by expansion of dendritic arborization and increased synaptogenesis in the remaining neurons (Bertoni-Freddari et al., 1996). The basis for the survival of selective neuronal populations is not yet clear. A key to this selectivity may be found from studies on the perinatal and young adult brain, as almost identical intercellular signals and intracellular transduction pathways that regulate neurite outgrowth, synaptogenesis and cell survival during development may provide a lead to understanding the molecular basis for their decline during aging (Keller and Mattson, 1998).

4.3.2 Pathologically Accelerated Aging Many clues regarding the role of LPO in aging have been discovered through compositional studies of brain specimens from various neurodegenerative disorders in humans and from animal models subjected to OS insults (Butterfield et al., 2001; Reynolds et al., 2007). Elevated levels of LPO products such as reactive aldehydes and isoprostanes were found in AD (Sayre et al., 1997) Down syndrome (Odetti et al., 1998) Parkinson’s disease (Barichella et al., 2008, vol 8 this series) and other dementias. Animal models mimicking AD have been instrumental in identifying lipid components that may participate in exacerbation of neurotoxicity or in unravelling the mechanisms which may bring about neuroprotection (Pappolla et al., 1998; Calon and Cole, 2007). Apart from reactive aldehydes and isoprostane species which are mostly derived from 1, 2-diacyl phospholipids, an increasing attention was given to pEPG species and their associated PUFAs as potential targets for OS both in aging and in neurodegenerative disorders (Farooqui et al., 2003). Significant reductions in pEPG and DHA content were noted in AD (So¨derberg et al., 1991, Guan et al., 1999), Zellweger disease (Martinez, 1989) and bipolar disorders (McNamara and Carlson, 2006; Mazza et al., 2007). The decrease in pEPG levels has been attributed to an increase of hydrolysis by PLs-specific PLA2 acting selectively on pEPG (Farooqui and Horrocks, 2001). Lack of pEPG replenishment in these disorders may be indicative of a poorly enzyme-catalyzed reacylation. Losses of PUFAs have been reported in the ageing neuronal membranes of the mouse brain presumably leading to memory deficits, learning disabilities, cognitive alterations and impaired visual acuity (Yehuda et al., 2002). On the other hand, supplements of n-3 PUFA in the diet appeared beneficial in a wide variety of pathophysiological conditions from cardiovascular to inflammatory diseases and caused a slow down of neurodegenerative diseases. Thus, numerous epidemiological studies suggest that increased fatty fish consumption and high DHA intake are associated with reduced risk of AD (Morris et al., 2003). Nevertheless, definitive answers for the molecular basis for n-3 PUFA neuroprotective effects remain unclear. Recent studies however have identified a whole spectrum of DHA-derived molecules including resolvins, docosatrienes, and neuroprotectins (Lukiw et al., 2005; Serhan, 2005) all of which appear to demonstrate some degree of neuroprotection. Added to these, the potential of DHA to accept hydroxyl radicals and thus abolish radical propagation in the lipid bilayer has not been decisively evaluated (Yavin et al., 2002). It would appear that any prediction about the dual pro-oxidant and antioxidant properties of DHA will remain hampered until quantitative data are obtained and the relative contribution of each pathway is resolved (see > Scheme 25-1).

5

Conclusions

The unleashed attack of free radicals on cellular macromolecules is believed to be a common denominator characteristic of a large number of pathophysiological conditions in the organism subjected to stress. In the central nervous system, disruption of the molecular cyto-architecture following excess free radicals formation is believed to be a major episode which predates aging, chronic neurodegenerative and neurodevelopmental disorders. The membrane lipids constitute a principal target in this sequel of events as a result of which lipid hydroperoxides are formed as intermediates of peroxidative reactions. A major target for these reactions are the PUFAs that contain multiple double-bonds such as AA (20:4 n-6) and DHA (22:6 n-3).

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It is generally accepted that AA-oxidized products are by far more damaging than those of DHA-oxidized products, yet the presence of the latter appears to confer protection. That raises two basically unresolved fundamental questions as to (1) how DHA, in such a highly selective yet indiscriminative fashion, imparts protection in so many human afflictions including cancer, heart disease, inflammation ischemic stroke and bipolar disorders and (2) what could underlie the molecular mechanism(s) for this action. While the details of the molecular mechanisms still remain unclear, the principal conclusion is that the ubiquity of DHA and its asymmetry in the hydrophobic compartment may provide the clue for its beneficial action. In other words an attempt to underpin the possible role of a specific group of PL molecular species containing ethanolamine as the polar head group and esterified DHA as bona fide antioxidants is a valid hypothesis. The purported role of such DHA is to target hydroxyl radicals and terminate free radicals propagation with supportive enzymatic mechanisms by generating nondamaging long-chain fatty acyl alcohols. Furthermore, the scrambling of the plasma membrane lipid asymmetry resulting from DHA hydroperoxides formation may alter signalling transduction cascades and affect cell survival or cell death. In addition, free DHA released by the action of PLA2 constitutes an important resource for the biosynthesis of a group of specific lipid messengers, resolvins and NPD1 which hold a great potential to provide neuroprotection. The antioxidant properties of DHA and its derivatives will be better understood with the advent of lipidomics technologies and use of high resolution high throughput techniques to decipher the complete vocabulary of PL molecular species as they appear both in health and disease.

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G. V. Raymond

1 1.1 1.2 1.2.1 1.2.2 1.3 1.3.1 1.4 1.4.1 1.4.2 1.4.3 1.4.4 1.4.5 1.4.6 1.4.7 1.4.8 1.4.9

Clinical Aspects of Peroxisomal Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 632 Introduction: Classification of Peroxisomal Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 632 Conditions Resulting from Defective Peroxisome Biogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 632 Zellweger Spectrum Disorders (ZSD) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 632 Neonatal Adrenoleukodystrophy (NALD) and Infantile Refsum Disease (IRD) . . . . . . . . . . . . . . . . 633 Therapy of Zellweger Spectrum Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 633 Rhizomelic Chondrodysplasia Punctata (RCDP), Type 1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 633 Defects of Single Peroxisomal b-Oxidation Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 634 Adrenoleukodystrophy (X-ALD) and Adrenomyeloneuropathy (AMN) . . . . . . . . . . . . . . . . . . . . . . . . 634 Childhood Cerebral Form of Adrenoleukodystrophy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 634 Adrenomyeloneuropathy (AMN) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 635 Adrenoleukodystrophy with Addison’s Disease Only . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 635 Asymptomatic Patients with the Biochemical Defect of Adrenoleukodystrophy . . . . . . . . . . . . . . . 635 Symptomatic Heterozygotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 635 Therapies of Adrenoleukodystrophy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 635 Acyl-CoA Oxidase Deficiency (ACOX1) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 636 D-Bifunctional Enzyme Deficiency (DBP) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 636

2 2.1 2.2 2.3 2.4 2.5 2.6 2.7 2.8

Lipid Metabolism in Peroxisomal Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 637 Elevated VLCFA in Peroxisomal Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 637 Vlcfa b-Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 637 Docosahexaenoic Acid Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 640 Branched-Chain Fatty Acid Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 641 Bile Acid Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 643 a-Methylacyl-CoA Racemase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 644 Plasmalogen Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 644 Cholesterol Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 645

3 3.1 3.2 3.3 3.4 3.5 3.6 3.7 3.8

Molecular Aspects of Peroxisomal Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 645 X-Linked Adrenoleukodystrophy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 647 Peroxisome Biogenesis Disorders, Zellweger Spectrum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 647 Rhizomelic Chondrodysplasia Punctata (RCDP) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 649 D-Bifunctional Protein Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 650 Acyl-CoA Oxidase (ACOX1) Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 650 Adult Refsum Disease (ARD) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 650 SCPx Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 650 a-Methylacyl-CoA Racemase (a-MACR) Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 651

4 4.1 4.2 4.3

Pathology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 651 Abnormalities in Neuronal Migration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 651 Abnormalities in Myelin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 655 Postdevelopmental Neuronal Degeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 659

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_26, # Springer ScienceþBusiness Media, LLC 2009

632

26

Peroxisomal disorders

List of Abbreviations: ACOX1, acyl-CoA oxidase deficiency; ACSVL1, very long-chain acyl-CoA synthetase; ADHAPS, alkyl-DHAP synthase; AMN, adrenomyeloneuropathy; ARD, adult refsum disease; CNS, central nervous systems; CTLs, cytotoxic lymphocytes; DBP, D-bifunctional enzyme deficiency; DHA, docosahexaenoic acid; DHAP, dihydroxyacetone phosphate; DHAPAT, dihydroxyacetonephosphate acyltransferase; DHCA, dihydroxycholestanoic acid; DPA, docosapentaenoic acid; DRG, dorsal root ganglia; FAR, fatty acyl-CoA reductase; HMG-CoA, b-Hydroxy-b-methylglutaryl-CoA; HNE, hydroxynonenal; I-CAM, intercellular adhesion molecules; IRD, infantile Refsum disease; LBP, L-bifunctional protein; a-MACR, a-methylacyl-CoA racemase; MRS, magnetic resonance spectroscopy; NALD, neonatal adrenoleukodystrophy; PAS, periodic acid-Schiff; PBD, peroxisome biogenesis disorders; PEX, peroxisome assembly; PNS, peripheral nervous systems; RCDP, rhizomelic chondrodysplasia punctata; SCPx, sterol carrier protein-X; THCA, trihydroxycholestanoic acid; TNF, tumor necrosis factor; VLCFA, very long chain fatty acids; WM, white matter; ZS, Zellweger syndrome; ZSD, Zellweger spectrum disorders

1

Clinical Aspects of Peroxisomal Disorders

1.1

Introduction: Classification of Peroxisomal Disorders

Peroxisomal disorders are divided into the two categories: (1) disorders of peroxisome assembly or peroxisome biogenesis disorders (PBD) and (2) single protein defects (it may be better to refer to these as single protein defects, since ABCD1 encodes a protein that appears to be a transporter and not an enzyme) (Raymond, 2001). In the first, the peroxisome fails to form and there are abnormalities of multiple peroxisomal enzymes. It is now understood that these disorders are defects in protein importation or membrane incorporation. The PBDs can be further divided by their clinical and biochemical features into the Zellweger spectrum disorders (ZSD) and rhizomelic chondrodysplasia punctata (RCDP). The second major group consists of a growing number of disorders in which there is a genetically determined abnormality of a single peroxisomal protein and peroxisomal structure is intact.

1.2

Conditions Resulting from Defective Peroxisome Biogenesis

Zellweger syndrome (ZS) and RCDP are the respective prototypes that result from defective peroxisomal biogenesis. Although it is recognized that these disorders are similar in their mechanisms of pathogenesis, the clinical and biochemical variation between these two types of assembly defects still makes it useful to discuss them separately.

1.2.1

Zellweger Spectrum Disorders (ZSD)

Zellweger (cerebrohepatorenal) syndrome was first described by Bowen et al. (Bowen et al., 1964; Opitz, 1985). This was later followed by the clinical descriptions of neonatal adrenoleukodystrophy (NALD) and infantile Refsum disease (IRD). Although classical ZS is the most severe form with a characteristic phenotype, there is clinical overlap between the three forms and more appropriately they should be referred to as a spectrum of clinical abnormalities. 1.2.1.1 Clinical Features of Zellweger Spectrum Disorders Zellweger syndrome is a multiple congenital anomaly syndrome secondary to a defect in one of the peroxisome assembly (PEX) genes. It is characterized by eye abnormalities, hearing loss, neuronal migration defects, hepatomegaly, chondrodysplasia punctata, and near complete absence of peroxisomes. The craniofacial features include a high forehead, hypoplastic supraorbital ridges, epicanthal folds, midface hypoplasia, and a large fontanel. Severe weakness and hypotonia manifest in the newborn period and is often accompanied by seizures and apnea (Heymans, 1984; Wilson

Peroxisomal disorders

26

et al., 1986a). Very little psychomotor development ensues and affected infants typically die in the first year (Wilson et al., 1986a).

1.2.2

Neonatal Adrenoleukodystrophy (NALD) and Infantile Refsum Disease (IRD)

NALD and IRD were described separately in children with developmental delays, seizures, liver disease, sensorineural hearing loss, and retinitis pigmentosa. The generalized peroxisomal dysfunction was only later documented (Poulos and Sharp, 1984; Poll-The et al., 1986, 1987; Poulos et al., 1986; Wanders et al., 1986). There is now a clearer understanding that alteration in PEX genes may result in clinical phenotypes that are less severe than ZS, but even the milder forms generally have mental retardation, retinal degeneration, and motor handicaps. Children have less severe dysmorphism than infants with ZS, renal cysts may be absent, and there is usually no stippling of the cartilage on radiographs. Patients with milder forms may present in the neonatal period with mild-to-moderate hypotonia, poor feeding, and hepatomegaly. There is usually delayed motor and cognitive development. Even with hypotonia, individuals may walk although gait is often unsteady. Retinal pigmentary degeneration often results in visual loss in the first years of life. Sensorineural hearing loss is associated with limited language development (Scotto et al., 1982; Weleber et al., 1984; Aubourg et al., 1986; Budden et al., 1986; Poll-The et al., 1987; Torvik et al., 1988; Wanders et al., 1990). Adrenal dysfunction may develop with age (Govaerts et al., 1984; Poll-The et al., 1987). Liver dysfunction is often present and detectable by persistent elevation in liver function tests. A bleeding diathesis that responds to vitamin K may also develop and several children have developed esophageal varices consistent with portal hypertension. Lifespan is variable and individuals have survived to adulthood. Several older patients have now been described. All identified patients have been visually impaired with sensorineural hearing loss. A progressive leukodystrophy has been seen in a number of patients with variable age of onset. It does result in loss of previously acquired skills and in most progresses to a vegetative state and death. All of the disorders in this group are inherited in an autosomal-recessive fashion and so it is important for genetic counseling that an accurate diagnosis is obtained. Carrier detection is not possible by biochemical determination in blood or fibroblasts, but may be performed by DNA mutational analysis.

1.3

Therapy of Zellweger Spectrum Disorders

Since many of the abnormalities are present in the affected fetus, potential for therapy is presently limited and likely to remain so. Treatment is primarily supportive. In the milder phenotypes, rehabilitative approaches including communication training and physical and occupational therapy are helpful. There have been attempts to normalize some of the biochemical abnormalities (Van Duyn et al., 1984; Bjorkhem et al., 1985; Lazarow et al., 1985; Wilson et al., 1986a, b; Greenberg et al., 1987; Setchell et al., 1992; Martinez, 1996; Maeda et al., 2002), but the clinical effectiveness of these interventions has yet to be demonstrated.

1.3.1

Rhizomelic Chondrodysplasia Punctata (RCDP), Type 1

RCDP is characterized by disproportionate shortening of the proximal portions of the extremities, short stature, microcephaly, dysmorphic facial appearance, cataracts, ichthyosis, and severe mental retardation. Radiologic highlights include shortening of the proximal limbs, metaphyseal cupping, and disturbed ossification with epiphyseal and extra-epiphyseal calcification (Spranger et al., 1971). This disorder is characterized by the triad of biochemical defects: (1) marked reduction of plasmalogen levels because of impaired plasmalogen synthesis (Heymans, 1984); (2) impaired oxidation of phytanic acid with increased plasma levels (Hoefler et al., 1988b); and (3) presence of 3-oxoacyl-CoA thiolase in the precursor form.

633

634

26

Peroxisomal disorders

RCDP is inherited in an autosomal-recessive manner. Treatment is very limited and consists of palliative orthopedic care and cataract removal. While dietary restriction of phytanic acid has been proposed, there is no evidence that it affects outcome.

1.4

Defects of Single Peroxisomal b-Oxidation Proteins

1.4.1

Adrenoleukodystrophy (X-ALD) and Adrenomyeloneuropathy (AMN)

1.4.1.1 Clinical and Pathologic Features of X-ALD and AMN One of the unique features of adrenoleukodystrophy is the variety of presentations that have been reported. All of these patients demonstrate accumulation of saturated very long chain fatty acids (VLCFA) and molecular studies show ABCD1 gene mutations. Various phenotypes have been recognized to occur within the same pedigree, so neither the genetic mutation nor biochemical abnormality predicts the clinical presentation. The clinical differences are mirrored in the pathologic findings.

1.4.2

Childhood Cerebral Form of Adrenoleukodystrophy

The childhood cerebral variant is the most common and fulminant form of X-ALD. The boys are normal until 4–8 years of age, when they manifest behavior problems and failure in school. This results from the rapid loss of auditory discrimination, spatial orientation, speech, and writing. Seizures occur in 30% of patients and in rare instances it may be the initial symptom. The MRI scan reveals parieto-occipital white matter lesions (85%) or frontal lesions (15%) at this stage, with contrast accumulation at the leading edge of the lesion (> Figure 26-1) (Duda and Huttenlocher, 1976; Eiben and Di Chiro, 1977; Kumar et al., 1987). There is rapid clinical deterioration to spastic quadriparesis, swallowing difficulty, and visual loss culminating in a vegetative state usually within 2 years of the initial symptoms. Although boys come to medical attention because of the neurologic deficits, impaired cortisol response can be identified by ACTH stimulation in 85% of this group (Moser et al., 1991). Rarely there is an adolescent presentation that is similar to the childhood form. . Figure 26-1 MRI in cerebral childhood adrenoleukodystrophy. Note posterior symmetric hyperintensity on T2-weighted image on the left. The image on the right shows the characteristic garland of contrast enhancement at the leading edge of the lesion

Peroxisomal disorders

1.4.3

26

Adrenomyeloneuropathy (AMN)

The neurologic manifestations of this adult form of X-ALD consist of an insidious onset and slow progression of spastic paraparesis, impaired vibratory sense in the lower extremity, and bladder or bowel dysfunction. Onset is typically in the third decade of life. The development of cerebral demyelination has been seen in approximately 15–20% of men with AMN (van Geel et al., 2001). This needs to be differentiated from those individuals with long tract findings on MRI (Loes et al., 2003). Cerebral disease is similar in time course to the childhood form of the disease and leads to a dementia, spasticity, blindness, and death. Approximately half of AMN patients appear to have some degree of cerebral involvement, with mild-to-moderate abnormalities noted on MRI in 46% (see > Figure 26-1) (Kumar et al., 1995). These abnormalities consist most frequently of parietal-occipital white matter and optic radiation involvement. Adrenal insufficiency or Addison’s disease precedes the onset of neurologic symptoms in 42% of patients; in some, this may occur 3–35 years earlier. Adrenal insufficiency was evident in 67% of patients in the series. Serum testosterone levels were abnormally low in 22% of patients, and early onset of sexual dysfunction occurred in one third.

1.4.4

Adrenoleukodystrophy with Addison’s Disease Only

Adrenoleukodystrophy with Addison’s disease only includes patients who have isolated Addison’s disease in the absence of neurologic signs and symptoms and is more common than previously recognized. Recognition of these patients is vital for genetic counseling and to monitor for the development of AMN or cerebral symptoms. Onset of adrenal dysfunction does not correlate with neurologic involvement.

1.4.5

Asymptomatic Patients with the Biochemical Defect of Adrenoleukodystrophy

Individuals may be diagnosed by measurement of plasma VLCFA during screening tests of relatives of symptomatic patients. These individuals may be of any age. The elevation of VLCFA levels in these males is comparable with that of severely affected members.

1.4.6

Symptomatic Heterozygotes

Women who are carriers for the adrenoleukodystrophy gene may manifest myelopathy findings resembling AMN between the third and fifth decades of life (Penman, 1960; Pilz and Schiener, 1973; Heffungs et al., 1980; Moser et al., 1980; Dooley and Wright, 1985; Noetzel et al., 1987). Spasticity with hyperreflexia and diminished vibration sense in the legs are present in two thirds of the patients, although only 25% complain of symptoms. In one series, approximately 14% had severe spinal involvement requiring assistance with ambulation and 5% had dementia (Naidu and Moser, 1990). In contrast to males, adrenal insufficiency rarely occurs in these women (el-Deiry et al., 1997). Elevation of VLCFA in plasma or skin fibroblasts allowed identification of 85% of obligate heterozygotes (Moser et al., 1983). The remaining cases can be identified by molecular analysis for the gene defect. Mutation analysis helps identify heterozygotes once the mutation in the index case is known (Boehm et al., 1999).

1.4.7

Therapies of Adrenoleukodystrophy

The two forms of therapy under study are dietary and bone marrow transplantation. Early on, it was determined that dietary restriction alone did not alter plasma levels of VLCFA. It was subsequently determined that the use of the monounsaturated fatty acid oils of oleic, 18:1o9, and erucic, 22:1 o9, combined with a low fat diet would reduce the levels of saturated VLCFA in plasma. This combination of

635

636

26

Peroxisomal disorders

glyceryl trioleate and trierucate is referred to as Lorenzo’s oil and effectively reduces VLCFA in the plasma in 4–8 weeks (Rizzo et al., 1989; Moser et al., 2005). Unfortunately, this regimen does not alter the course of the childhood cerebral form of ALD. In open, uncontrolled studies, the effects on the adult form have been uncertain but generally unimpressive (Aubourg et al., 1993; Kaplan et al., 1995). However, recently, there has been renewed interest in the use of the oil as a preventative agent in presymptomatic boys. In several series, it has been shown that the reduction of very long chain fatty acids over time correlates with a reduction in the risk of childhood cerebral disease. The results of bone marrow transplantation in mildly affected patients are encouraging (Peters et al., 2004). In patients with more advanced disease, transplantation has not resulted in halting the course and may be associated with worsening of the neurologic status immediately after the procedure and therefore is not advised (Moser et al., 1984). In those with advanced disease, neither dietary therapy nor bone marrow transplantation is effective. Boys at risk for developing the childhood form of X-ALD ( Table 26-1 (Wanders, 2004a). In fact, investigation of lipid metabolism in these disorders has broadened the understanding of several lipid metabolic pathways and identified many new enzymes. Because all are genetic diseases, alterations in metabolism are found in many or most tissues; thus, only a limited number of specific studies of the nervous system have been conducted. (Note: Symbols for genes and proteins in [square brackets] in the text and figures are those approved by the Human Genome Organization nomenclature committee.)

2.1

Elevated VLCFA in Peroxisomal Diseases

A laboratory finding of elevated VLCFA levels in plasma and tissues is pathognomonic for peroxisomal disease. Increased VLCFA are found in patients with PBD, X-ALD, and deficiencies of peroxisomal b-oxidation enzymes (Igarashi et al., 1976a; Moser et al., 1980, 1981, 1984; Watkins et al., 1995). In X-ALD brain, elevated VLCFA are associated with the inflammatory demyelinating lesions characteristic of this disease (Griffin et al., 1985). In contrast, neuronal migration defects are more characteristic of brain in PBD (Volpe and Adams, 1972). In these latter cases, metabolic defects in addition to elevated VLCFA also contribute to the pathology. In brain tissue from all patients with elevated VLCFA, the fatty acids are most abundant in cholesterol esters, gangliosides, phosphatidylcholines, and proteolipid proteins but are found in all lipid classes (Sharp et al., 1987; Theda et al., 1992). Elevated VLCFA levels could arise from either overproduction via elongation of long-chain fatty acids, or by a diminished capacity to degrade these potentially toxic fatty acids. Because elongation pathways are found in mitochondria and/or endoplasmic reticulum, and not peroxisomes, most studies have focused on peroxisomal VLCFA degradation.

2.2

Vlcfa b-Oxidation

While mitochondria efficiently degrade long-chain fatty acids by a b-oxidation pathway that is tightly coupled to energy production, they cannot catabolize VLCFA. Although peroxisomes lack an ATPgenerating electron transport chain, the matrix of the organelle contains b-oxidation enzymes capable of chain-shortening VLCFA (> Figure 26-2) (reviewed in Wanders and Waterham, 2006). In general, a VLCFA entering the peroxisome, is activated to its CoA thioester, and undergoes four sequential enzymatic reactions (oxidation, hydration, dehydrogenation, and thiolytic cleavage) to yield acetyl-CoA plus a fatty acyl-CoA that is two carbons shorter than the original VLCFA.

637

↓ Normal Normal Normal

↓

Normal

Normal

Normal

↓

↓

↓

↓

↑

↑

↑

Acyl-CoA oxidase deficiency ↑ Normal Normal Normal

X-linked adrenoleukodystrophy ↑ Normal Normal Normal

Peroxisome biogenesis disorders ↑ Normal or ↑b Normal or ↑b ↑

Normal

↓

↓

↓

↑

D-bifunctional protein deficiency ↑ Normal ↑ ↑

b

The C26:0 level in a single case of SCPX deficiency was just above the upper limit of normal (REF) Normal in infants. Increased in older children, depending on dietary intake of ruminant fats c Increased in RCDP Type 1. Normal in RCDP Types 2 and 3

a

Biochemical parameters Plasma VLCFA (C26:0) Plasma phytanic acid Plasma pristanic acid Plasma bile acid synthesis inter-mediates (THCA; DHCA) Fibroblast VLCFA (C26:0) Fibroblast VLCFA betaoxidation Fibroblast phytanic acid a-oxidation Fibroblast pristanic acid b-oxidation Fibroblast plasmalogen synthesis Normal

Normal

?

↓

Normal

Normal

↓

Normal

Normal

↓

Refsum disease Normal ↑ Normal Normal

SCPx thiolase deficiency Borderlinea ↑ ↑ ↑

↓

Normal

↓

↓

↓ or Normald Normal

?

Normal

a-Methylacyl-CoA racemase deficiency Normal ↑ ↑ ↑

Normal

Normal

Rhizomelic chondrodysplasia punctata Normal ↑ or Normalc Normal Normal

26

. Table 26-1 Biochemical abnormalities in peroxisomal disorders

638 Peroxisomal disorders

Peroxisomal disorders

26

. Figure 26-2 Peroxisomal fatty acid b-oxidation pathways. Left, degradation of VLCFA and straight-chain fatty acids. Details of the enzymatic reactions are found in the text. The products of one cycle of VLCFA b-oxidation are acetyl-CoA and a VLCFA-CoA that has been shortened by two carbons (n-2 VLCFA-CoA); the latter product undergoes several additional cycles of b-oxidation until a chain length of 8-10 carbons (medium-chain fatty acyl-CoA) has been reached. Carnitine octanoyltransferase [CROT] and carnitine acetyltransferase [CRAT] convert mediumchain acyl-CoA and acetyl-CoA to their respective carnitine derivatives for export from peroxisomes. Right, degradation of pristanic acid, the a-methyl branched-chain fatty acid product of phytanic acid a-oxidation. Pristanoyl-CoA undergoes three cycles of peroxisomal b-oxidation. The first and third cycles cleave a 3-carbon CoA derivative, propionyl-CoA, whereas the second cycle yields acetyl-CoA. The medium-chain degradation product, dimethyl-nonanoyl-CoA, is converted to its carnitine derivative by CROT for export. The pathway on the right is also required for the conversion of cholesterol to bile acids (see > Figure 26-4). Enzymes are designated by their Human Genome Organization approved symbols

The process by which a VLCFA enters the peroxisome and is esterified to CoA remains controversial. It has not been firmly established whether it is the free fatty acid or the CoA derivative that enters the organelle, or whether a very long-chain acyl-CoA synthetase (ACSVL1; [SLC27A2]) that activates VLCFA resides inside or outside the peroxisome (Lageweg et al., 1991; Steinberg et al., 1999).

639

640

26

Peroxisomal disorders

One hypothesis proposes that VLCFA enter the organelle by diffusion, at which point they are activated by an intraperoxisomal ACSVL1. An alternative hypothesis suggests that VLCFA are activated extraperoxisomally, and VLCFA-CoA are transported into the organelle by the adrenoleukodystrophy protein [ABCD1] (Wanders and Waterham, 2006). Once inside the peroxisome, VLCFA-CoAs are oxidatively degraded by four reactions catalyzed by three distinct enzymes. Acyl-CoA oxidase 1 [ACOX1], a flavin-linked oxygen-dependent enzyme, catalyzes the a,b unsaturation of the VLCFA (Osumi et al., 1980). Hydrogen peroxide, also a product of this reaction, is detoxified by the peroxisomal enzyme catalase. The second and third steps of b-oxidation – enoyl-CoA hydratase and hydroxyacyl-CoA dehydrogenase – are catalyzed primarily by DBP; [HSD17B4]) (Jiang et al., 1996a, b). The product of these reactions is a b-ketoacyl-CoA that is thiolytically cleaved in a CoAdependent reaction catalyzed by peroxisomal thiolase [ACAA1] (Miyazawa et al., 1980, 1981). This process is then repeated several times until the carbon chain length has been reduced to 8–10 carbons. The b-oxidation products, acetyl-CoA and medium-chain acyl-CoA, are then converted to carnitine derivatives by carnitine acetyltransferase [CRAT] and carnitine octanoyltransferase [CROT], respectively (Markwell et al., 1976; Farrell et al., 1984). The carnitine compounds are then exported into the cytoplasm for subsequent metabolism in mitochondria. In addition to DBP and thiolase, L-bifunctional protein (LBP; [EHHADH]) and sterol carrier protein-X (SCPx) thiolase [SCP2], respectively, may contribute to the VLCFA chain-shortening process (Palosaari and Hiltunen, 1990; Wanders et al., 1997). As expected, patients with PBD who fail to import essentially all of the above enzymes into nascent organelles have profoundly impaired VLCFA b-oxidation. This process is also severely impaired in patients with DBP deficiency, suggesting that LBP does not contribute significantly to VLCFA catabolism (van Grunsven et al., 1999). Patients with X-ALD have a less severe reduced capacity to degrade VLCFA. The peroxisomal membrane protein defective in X-ALD, ABCD1, is a member of the ATP-binding cassette transporter protein family and has been proposed to translocate VLCFA or VLCFA-CoA into the organelle, thus providing an explanation for the b-oxidation defect;(Wanders and Waterham, 2006) this awaits more rigorous experimental verification. Interestingly, the b-oxidation impairment in ACOX1-deficient patients is more similar to that in X-ALD than in PBD or DBP-deficiency, but the reason for this observation is unclear (Watkins et al., 1995). Although documented ACAA1 thiolase deficiency is not known, peroxisomes from patients with RCDP (see below) fail to import this enzyme, yet contain the other b-oxidation enzymes. Fibroblasts from RCDP patients have normal degradation of VLCFA, suggesting that SCPx thiolase can fulfill this function equally well (Hoefler et al., 1988a). Shortly after it was reported that VLCFA levels were increased in X-ALD, a few studies supporting the notion of increased VLCFA synthesis were published (Tsuji et al., 1981, 1984). However, when it became apparent that X-ALD was a peroxisomal disorder, focus shifted away from the non-peroxisomal process of chain elongation. However, the development of sensitive mass spectrometry methods for investigating elongation led some investigators to re-address this issue. It now seems clear that, in addition to a decreased rate of b-oxidation in X-ALD, elevated VLCFA levels result from increased synthesis via elongation of palmitic acid (C16:0) (Kemp et al., 2005).

2.3

Docosahexaenoic Acid Synthesis

An important function of the peroxisomal b-oxidation pathway is the synthesis of docosahexaenoic acid (DHA; C22:6o3). High concentrations of this o3 fatty acid are found in nervous tissue, and many studies have demonstrated the importance of DHA in brain and retina development, learning, memory, and behavior (reviewed in Innis, 2005). While the brain efficiently takes up preformed DHA from the circulation, astrocytes (and likely some neurons) are capable of robust synthesis of DHA (Williard et al., 2001; Kaduce et al., 2008). Essential dietary o3 fatty acids such as a-linolenic acid (C18:3o3) are taken up from the circulation by brain capillary endothelial cells and transported to astrocytes. Two cycles of the microsomal chain elongation pathway, along with insertion of double bonds by D6- and D5-desaturases, convert linolenic

Peroxisomal disorders

26

acid to docosapentaenoic acid (DPA; C22:5o3). For many years, it was hypothesized that a D4-desaturase converted DPA to DHA by insertion of the sixth double bond; however, evidence for this activity was inconclusive. It is now established that mammals utilize an alternative strategy for DHA synthesis (Voss et al., 1991; Moore et al., 1995). DPA is converted to tetracosapentaenoic acid (C24:5o3) by an additional elongation cycle, after which the D6-desaturase inserts the sixth double bond, forming tetracosahexaenoic acid (C24:6o3). One cycle of peroxisomal b-oxidation then chain-shortens this fatty acid, yielding DHA. The b-oxidation enzymes that degrade VLCFA are those required for DHA synthesis (> Figure 26-2). This was investigated mainly in skin fibroblasts from patients with PBD, RCDP, and deficiencies of peroxisomal b-oxidation enzymes. As expected, DHA synthesis from C24:6o3 was decreased in PBD, ACOX1 deficiency, and DBP deficiency, but not in RCDP (Su et al., 2001). DHA synthesis is also normal in X-ALD, suggesting that ABCD1 deficiency does not affect b-oxidation of C24:6o3 (Su et al., 2001).

2.4

Branched-Chain Fatty Acid Metabolism

Phytanic acid (3,7,11,15-tetramethylhexadecanoic acid) degradation is impaired in Refsum disease, leading to elevated tissue levels that are thought to contribute to the observed pathologic findings (reviewed in Wanders et al., 2001). Cerebellar ataxia, peripheral polyneuropathy, retinitis pigmentosa, and sensorineural deafness are among the primary neurological deficits in this disease. Partial reversal of many symptoms can be achieved by dietary restriction of this fatty acid. Phytanic acid is a b-methyl branched-chain fatty acid derived from the phytol side-chain of chlorophyll. Mammals cannot synthesize this fatty acid, but ingest it in the diet (Masters-Thomas et al., 1980a, b). Bacteria in the rumen of cows and related species hydrolyze chlorophyll, releasing the phytol that is then oxidized to phytanic acid and incorporated into complex lipids. Ruminant meats and dairy products are thus the primary sources of dietary phytanic acid in humans. The presence of the b-methyl group in phytanic acid precludes its degradation via b-oxidation unless the carboxyl carbon is first removed via a-oxidation (> Figure 26-3) (reviewed in Wanders et al., 2003; van den Brink and Wanders, 2006). The subcellular location of the latter pathway was controversial for many years, but it is now established that it resides in peroxisomes. Phytanic acid is thought to be activated to its CoA thioester extraperoxisomally; however, similar to the situation with VLCFA, how activated phytanic acid enters the organelle remains unclear. The peroxisomal enzyme phytanoyl-CoA a-hydroxylase [PHYH] (Jansen et al., 2004) is a dioxygenase that requires ferrous iron and an a-ketoglutarate co-substrate, and yields a-hydroxyphytanoyl-CoA (Mihalik et al., 1995). The product is then cleaved by a unique thiamine phosphate-dependent lyase [HACL1], yielding a one-carbon CoA compound, formyl-CoA, and an aldehyde, pristanal (Foulon et al., 1999; Van Veldhoven et al., 1999). Formyl-CoA degrades non-enzymatically to formate and CoA; formate likely exits the peroxisome for subsequent metabolism in mitochondria. Pristanal is then oxidized to pristanic acid (2,6,10,14-tetramethylpentadecanoic acid) via a splice variant of fatty aldehyde dehydrogenase [ALDH3H2] found in peroxisomes (Ashibe et al., 2007). Because the position of the methyl group has now shifted, pristanic acid can be degraded by peroxisomal b-oxidation. This requires activation to the CoA derivative by a peroxisome-associated acyl-CoA synthetase, e.g., ACSL4 or SLC27A2 (> Figure 26-3) (Steinberg et al., 1999; Lewin et al., 2002). The peroxisomal b-oxidation pathway for pristanic acid and other a-methyl-branched-chain fatty acids differs somewhat from that for VLCFA (> Figure 26-2) (reviewed in Wanders et al., 2003). Genes encoding two branched-chain acyl-CoA oxidases [ACOX2, ACOX3] are found in humans, but only ACOX2 protein is expressed under normal conditions (Casteels et al., 1990; Van Veldhoven et al., 1992). DBP [HSD17B4] catalyzes the second and third reactions. Because of the a-methyl branch, the thiolytic cleavage reaction catalyzed by SCPx thiolase [SCP2] yields the three-carbon compound propionyl-CoA and 4,8,12-trimethyltridecanoyl-CoA. The latter product can undergo additional cycles of b-oxidation, alternatively cleaving 2- and 3- carbon fragments. Like b-oxidation enzymes, a-oxidation enzymes are not imported into peroxisomes of patients with PBD. However, since phytanic acid is solely of dietary origin, the toxic burden of this fatty acid is dependent

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. Figure 26-3 Peroxisomal a-oxidation of phytanic acid. The methyl group on carbon-3 of phytanic acid precludes its degradation by b-oxidation; removal of a single carbon via the a-oxidation pathway yields pristanic acid, which can be degraded by b-oxidation (see > Figure 26-2). Details of the enzymatic reactions are found in the text. In addition to pristanal, the reaction catalyzed by HACL1 yields a one-carbon CoA compound, formyl-CoA (not illustrated). Enzymes are designated by their Human Genome Organization approved symbols

. Figure 26-4 Peroxisomal reactions in the synthesis of bile acids. Reactions catalyzed by enzymes found in the endoplasmic reticulum and mitochondria convert cholesterol to bile acid precursors THCA and DHCA; the latter compound lacks the 12-hydroxyl group. The side chain of THCA and DHCA resembles an a-methyl fatty acid and undergoes one cycle of b-oxidation in peroxisomes via the pathway shown in > Figure 26-2, right-hand side. The products of chain shortening of THCA and DHCA are the CoA derivatives of cholate and chenodeoxycholate, respectively. BAAT catalyzes the conjugation of these bile acyl-CoAs to taurine (primarily) or glycine, with release of CoA

Peroxisomal disorders

26

upon the age and diet of the patient. Nonetheless, impaired degradation of both phytanic and pristanic acids can be demonstrated in vitro in fibroblasts from PBD patients (Singh et al., 1990; Verhoeven et al., 1997; Verhoeven et al., 1998). Defects in PHYH are the primary cause of Refsum disease (Jansen et al., 1997a, b, c; Mihalik et al., 1997), where symptoms presenting in the first decade of life are unusual. Pristanic acid b-oxidation is not impaired in Refsum disease. Patients with RCDP, in addition to their failure to import peroxisomal thiolase, also do not import PHYH and have defective a-oxidation (Mihalik et al., 1997; Jansen et al., 1997a; Hoefler et al., 1988a). While these patients usually present in infancy, before ingesting significant quantities of phytanic acid, some individuals are only mildly affected and survive to adulthood where their clinical presentation is phenotypically indistinguishable from that of Refsum disease (van den Brink et al., 2003; Horn et al., 2007). Because DBP is essential for the degradation of branchedchain fatty acids, deficiency of this enzyme results in elevated levels of both pristanic and phytanic acids (Ferdinandusse et al., 2006a), as well as VLCFA and bile acid intermediates (see below). Phytanic acid, pristanic acid, and bile acid intermediates, but not VLCFA, were elevated in the one reported case of SCPx thiolase deficiency (Ferdinandusse et al., 2006b).

2.5

Bile Acid Synthesis

Bile acids are synthesized from cholesterol in a multi-step process involving several organelles, including peroxisomes. Defects in bile acid synthesis and accumulation of precursors in the biosynthetic pathway are observed in several peroxisomal diseases. Although synthesis of bile acids occurs mainly in liver, there are anecdotal observations suggesting that the brain may participate in this process. Levels of protein-bound chenodeoxycholate were found in normal rat brain that were 30-fold higher than serum levels; smaller concentrations of cholate and deoxycholate were also detected (Mano et al., 2004). Brain xanthomas in patients with cerebrotendinous xanthomatosis (deficiency of the mitochondrial enzyme sterol 27-hydroxylase) contained high concentrations of bile acid precursor derivatives (Panzenboeck et al., 2007). The brain converts cholesterol to 24-hydroxycholesterol, which is efficiently converted into bile acids (Lutjohann, 2006). Despite these observations, direct demonstration of brain bile acid synthesis is lacking. The initial steps of bile acid synthesis involve modifications (hydroxylation; oxidation/reduction) of the sterol nucleus that occur in the endoplasmic reticulum. The side-chain of cholesterol is then oxidized to a carboxylic acid by the sequential action of sterol 27-hydroxylase (mitochondria) and alcohol/aldehyde dehydrogenases (mitochondria or cytoplasm) (reviewed in Bove et al., 2004). The two products resulting from these reactions, di- and trihydroxycholestanoic acid (DHCA, THCA), are then converted to the primary bile acids chenodeoxycholate and cholate, respectively, in peroxisomes (> Figure 26-4). The 27-carbon precursors, DHCA and THCA, differ only by the absence or presence of a a-hydroxyl group on carbon 12. The side chains of both compounds resemble a-methyl-branched-chain fatty acids such as pristanic acid. To produce the mature, 24-carbon bile acids, therefore, the precursors are shortened by undergoing one cycle of peroxisomal b-oxidation. THCA and DHCA are first activated by one of two members of the very long-chain acyl-CoA synthetase family, ACSVL1 [SLC27A2] or ACSVL6 [SLC27A5] (Mihalik et al., 2002). While both enzymes are found in the endoplasmic reticulum, ACSVL1 is also found in peroxisomes (> Figure 26-4). The same peroxisomal b-oxidation enzymes and reactions that degrade pristanic acid (see > Figure 26-3), ACOX2, DBP [HSD17B4], and SCPx thiolase [SCP2], then cleave a three-carbon fragment from the sterol side chain (as propionyl-CoA). The 24-carbon products of b-oxidation are choloyl-CoA (from THCA) or chenodeoxycholoyl-CoA (from DHCA). The peroxisomal enzyme bile acid-CoA:amino acid N-acyltransferase [BAAT] produces glycine or taurine conjugates of the bile acids by the formation of an amide linkage (Pellicoro et al., 2007). The mechanism(s) of entry of precursors (or their CoA derivatives) into peroxisomes, and the exit of conjugated primary bile acids from peroxisomes, is unknown. Defective bile acid synthesis, and accumulation of precursors DHCA and THCA, are thus found in individuals with PBD, as well as with patients lacking DBP and SCPx thiolase.

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Peroxisomal disorders

a-Methylacyl-CoA Racemase

Naturally occurring phytanic acid is a mixture of two diastereoisomers (3S,7R,11R and 3R,7R,11R), and the a-oxidation pathway degrades both to pristanic acid equally well. However, the subsequent b-oxidation pathway is specific for the S-conformer (Croes et al., 1999). The bile acid precursors THCA and DHCA also exist as 25R and 25S stereoisomers. The enzyme a-methylacyl-CoA racemase, found both in peroxisomes and mitochondria, is thus needed for the complete degradation of branched-chain fatty acids and for efficient bile acid synthesis. 3R-Pristanic acid and 25R-DHCA/THCA accumulates in patients with deficiency of this enzyme, and have been associated with neurological defects including peripheral neuropathy and retinitis pigmentosa (Ferdinandusse et al., 2000, 2001).

2.7

Plasmalogen Biosynthesis

Decreased levels of ether-linked phospholipids, which normally comprise 18% of membrane phospholipids, are found in tissues of patients with PBD and RCDP (reviewed in Wanders, 2004b). Two enzymes essential for plasmalogen synthesis are found in peroxisomes, and a third is associated with the exterior of peroxisomes (> Figure 26-5). The peroxisomal enzyme dihydroxyacetonephosphate acyltransferase (DHAPAT) catalyzes the acylation of carbon 1 of dihydroxyacetone phosphate (DHAP) (Webber and Hajra, 1993). Outside the peroxisome, the enzyme fatty acyl-CoA reductase (FAR) catalyzes the NADPHdependent conversion of a fatty acyl-CoA to the corresponding fatty alcohol, which then enters the

. Figure 26-5 Peroxisomal reactions in plasmalogen synthesis. Synthesis of plasmalogens (ether-linked phospholipids) requires two peroxisomal matrix enzymes (GNPAT and AGPS) and enzyme(s) associated with the cytoplasmic face of the peroxisome membrane (FAR1,2). Details of the enzymatic reactions are found in the text. GNPAT and AGPS are symbols approved by the Human Genome Organization; the symbols FAR1 and FAR2 have not yet been approved. DHAP, dihydroxyacetone phosphate; EtN, ethanolamine

Peroxisomal disorders

26

organelle by an unknown mechanism (Cheng and Russell, 2004). Once inside the peroxisome, the fatty alcohol displaces the fatty acid on DHAP and forms an ether linkage, catalyzed by peroxisomal alkyl-DHAP synthase (ADHAPS; [AGPS]) (Zomer et al., 1993). Subsequent enzymatic steps, including reduction of alkyl-DHAP to alkyl-glycerophosphate, addition of an acyl group on carbon -2, desaturation of the alkyl group to an alkenyl group, and addition of choline or ethanolamine head groups, occur extraperoxisomally (Brites et al., 2004). Low plasmalogen levels are thought to contribute significantly to the neural pathology in both PBD and classical RCDP (Type 1) (Wanders, 2004b). While developmental delay and profound mental retardation are common to both disorders, neuronal migration defects are observed only in PBD, suggesting that the latter is not caused by plasmalogen deficiency. Patients with single enzyme deficiencies of DHAPAT (RCDP Type 2) and alkyl-DHAP synthase (RCDP Type 3) have low plasmalogens but, unlike Type 1, have normal phytanic acid metabolism (Wanders et al., 1992, 1994). These individuals have similar neurologic deficits as seen in Type 1, suggesting a common etiology of plasmalogen deficiency.

2.8

Cholesterol Synthesis

Nearly all cells are capable of synthesizing cholesterol. In the brain, astrocytes produce large amounts of this essential sterol for use by other neural cells (Pfrieger, 2003). Many early steps in cholesterol biosynthesis are now known to reside in peroxisomes (> Figure 26-6) (reviewed in Kovacs et al., 2002). The fundamental precursor of cholesterol, acetyl-CoA, is generated intraperoxisomally via the b-oxidation of VLCFA (see > Figure 26-2). Peroxisomal thiolase [ACAA1], working in the reverse direction from that in b-oxidation, condenses two molecules of acetyl-CoA to form acetoacetyl-CoA. b-Hydroxy-b-methylglutaryl-CoA (HMG-CoA) is formed by the condensation of acetoacetyl-CoA with another acetyl-CoA molecule via HMG-CoA synthase [HMGCS1] (Ashmarina et al., 1994). These two enzymatic activities are also found in the cytoplasm. HMG-CoA is then converted to mevalonate by HMG-CoA reductase [HMGCR], an enzyme found both in peroxisomes and endoplasmic reticulum (Kovacs et al., 2001). Five enzymes that act sequentially to convert mevalonate to farnesyl pyrophosphate are found exclusively in peroxisomes. These include mevalonate kinase [MVK], phosphomevalonate kinase [PMVK], diphosphomevalonate decarboxylase [MVD], isopentenyl pyrophosphate isomerase [IDI1], and farnesyl diphosphate synthase [FDPS] (Biardi and Krisans, 1996). The final product of these reactions, farnesyl pyrophosphate, leaves the peroxisome and traffics to endoplasmic reticulum membranes. There, two 15-carbon farnesyl pyrophosphate molecules condense to form squalene, which subsequently cyclizes to form the sterol nucleus. Many other enzymatic reactions associated with the endoplasmic reticulum are required to complete the synthesis of cholesterol. Despite ample evidence that peroxisomes are essential for cholesterol synthesis, the degree of biochemical abnormality measured in peroxisomal disease patients has been somewhat variable. Plasma cholesterol levels are generally lower than normal in patients with PBD (Kovacs et al., 2002), but two studies found that synthesis of cholesterol from radiolabeled acetate in PBD skin fibroblasts was not reduced (Malle et al., 1995). Activities of all peroxisomal enzymes involved in the conversion of HMG-CoA to farnesyl pyrophosphate were lower in liver samples from PBD patients (Krisans et al., 1994); MVK activity was also reduced in liver samples from two RCDP (Type 1) patients (Wanders and Romeijn, 1998). No studies of brain cholesterol synthesis have been reported.

3

Molecular Aspects of Peroxisomal Disorders

Molecular testing is possible for all of the peroxisomal disorders reported in > Table 26-1. In most cases molecular diagnosis is available in at least one international clinical laboratory and is more broadly available for the more common disorder X-linked adrenoleukodystrophy. Gene Tests (http://www. genetests.org/) is an excellent resource to identify laboratories currently offering clinical and research

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. Figure 26-6 Peroxisomal reactions in cholesterol biosynthesis. In addition to being exported from the peroxisome as acetylcarnitine, acetyl-CoA produced during the b-oxidation of VLCFA (see > Figure 26-2) can serve as a cholesterol precursor. Although the first eight enzymes required for cholesterol synthesis are found in peroxisomes, ACAA1, HMGCS1 and HMGCR are also in other cellular compartments. Reactions subsequent to the formation of farnesyl-pyrophosphate are extraperoxisomal. Enzymatic reactions are described in the text. Enzyme symbols are those approved by the Human Genome Organization. HMG, hydroxymethylglutaryl

testing for peroxisomal disorders and many other inherited disorders. Molecular testing is useful for confirmation of a clinical and biochemical diagnosis. This information has great utility in the genetic counseling of families and for family planning, including prenatal testing and preimplantation genetic diagnosis.

Peroxisomal disorders

3.1

26

X-Linked Adrenoleukodystrophy

ABCD1 encodes the adrenoleukodystrophy protein and is localized to Xq28. It is a 10 exon gene. Ancient duplication events have resulted in several pseudogenes that can confound analysis of the 30 end of the gene. It is critical to account for these pseudogenes when creating a sequence based test. The test described by Boehm et al. (Boehm et al., 1999) is highly reliable. In most cases the clinical and biochemical phenotypes of affected males is sufficient for diagnosis; molecular testing for this X-linked disorder is most useful for the testing of at-risk females, because biochemical testing detects only about 80% of these individuals. Sequence analysis of the ABCD1 coding exons of a hemizygote and obligate or biochemically confirmed female heterozygotes identifies a mutation in 93% of cases. If Southern blot analysis is performed to search for deletions and other rearrangements, test sensitivity approaches 99%. An excellent resource for the interpretation of ABCD1 sequence variations is the X-linked adrenoleukodystrophy database (http://www. x-ald.nl/). ABCD1 mutations identified in 310 patients tested in the DNA Diagnostic Lab at Johns Hopkins Hospital is summarized in > Table 26-2. There are not any common mutations, but some mutations are recurrent and some of these can be accounted for by CpG dinucleotides.

3.2

Peroxisome Biogenesis Disorders, Zellweger Spectrum

Mutations in 12 different PEX genes have been reported in patients with ZSD. These are summarized in > Table 26-3. About 15–20% of patients suspected to be in the Zellweger spectrum and who have elevated plasma VLCFA actually have defects in D-bifunctional protein (HSD17B4) or acyl-CoA oxidase (ACOX1). Biochemical analysis in blood and fibroblasts can distinguish most ZSD patients from those with a single protein deficiency. Several algorithms have been reported for the diagnosis of these disorders, including biochemical and molecular analysis (Wanders et al., 2001; Steinberg et al., 2006). Some take the approach of performing complementation analysis to identify the putative gene defect first and others to perform sequence analysis of select exons initially. This latter approach is possible because PEX1 is defective in about 70% of ZSD patients due to the presence of two common alleles: Nt2098insT (I700fs) and Nt2528G>A (G843D). At least one of these alleles is present in about 50% of all ZSD patients. The second reason that this approach is tenable is because six of the 12 PEX gene defects account for greater than 90% of all ZSD patients. Four of these genes are not large and mutations tend to be concentrated in a limited number of exons. The database dbPEX (www.dbpex.org) catalogs PEX gene mutations that are reported in the literature and submitted by individual contributors. A recent report of complementation analysis in over 500 ZSD patients found that all of these patients had mutations in one of the 12 PEX genes previously known to be associated with this disease spectrum (Bruneel et al., 2007). Thus, although there are more than 12 known

. Table 26-2 Types of ABCD1 Mutations identified in 310 patients tested by sequence analysis and Southern blot ABCD1 mutation type Missense Nonsense Frameshift Splice site Deletion, small & in frame Deletion, large Insertion, in frame No mutation detected

% of patients 63.5 10.3 11.3 4.5 3.2 5.5 0.6 1

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. Table 26-3 PEX gene defects in Zellweger Spectrum Disorders (ZSD) Complementation groups

Gene PEX1

Proportion of ZSD Patients (%) 70

PEX2

3

Clinical phenotypes ZS NALD IRD ZS

PEX3

A), and Nt370–396del27bp. The majority of the common PEX7 alleles are located in exons 7 and 9, and thus these exons can be sequenced initially in patients suspected to have RCDP1. In at least 5% of RCDP1 patients only a single heterozygous allele was identified by sequence analysis of the coding region; since these are patients with biochemical evidence of a PEX7 deficiency, it is likely that their second mutation is due to a gene rearrangement/large deletion or a mutation in non-coding DNA that is less easily identified. Although the genes encoding the enzymes responsible for RCDP2 and RCDP3 are both known and clinical testing is available for both, very little has been reported in the literature about mutations in these genes. Ofman et al. reported mutation analysis of the GNPAT gene encoding the DHAP-AT enzyme in 12 patients (Ofman et al., 2001). A variety of mutations, including clearly deleterious splice site and small insertion or deletions causing frameshift, were identified in all 12 patients. Of interest, all except one patient was apparently confirmed to be homozygous. In 1998 deVet et al. reported a missense mutation (R419H) in the ADHAPS gene in an RCDP3 patient (de Vet et al., 1998). A recombinant enzyme harboring this mutation had no enzyme activity.

3.4

D-Bifunctional Protein Deficiency

The D-bifunctional protein (736 amino acids) is encoded by a 24 exon gene. The most comprehensive analysis reported to date is a series of 110 patients who had cDNA sequence analysis (Ferdinandusse et al., 2006). Two alleles were identified in relatively high frequencies: G16S 24% and N457Y 11%. In addition, there was evidence of a genotype-phenotype correlation. Overall this is a very severe disorder, though, and in the Fernandusse cohort G. In addition, three ARD patients have been reported in the literature that have PEX7 gene mutations and thus represent the mild end of the PEX7 gene mutation biochemical and clinical spectrum.

3.7

SCPx Deficiency

This disorder has been recognized in only one patient. The patient was homozygous for a 1 nucleotide insertion (Ferdinandusse et al., 2006b).

Peroxisomal disorders

3.8

26

a-Methylacyl-CoA Racemase (a-MACR) Deficiency

Gene mutations in the 5 exon gene encoding the a-MACR enzyme have been reported in the literature on a very limited number of patients. Although the majority of patients have an adult onset disorder with a predominantly neurological phenotype, one early onset case has been reported with predominantly hepatic dysfunction (Setchell et al., 2003). This patient was homozygous for the same mutation (S52P) identified in two adult onset a-MACR deficiency patients with neurological presentations (Ferdinandusse et al., 2000). Thus, although only recognized in a small number of patients so far, there does not appear to be a straightforward genotype-phenotype correlation.

4

Pathology

Peroxisomal disorders exhibit systemic lesions that involve several organ systems: adrenal cortex (> Figure 26-7), ears, eyes, kidney, liver, and skeleton, in addition to the central and peripheral nervous systems (CNS, PNS). It is beyond the purview of this section to discuss the non-neural lesions of peroxisomal disorders. The reader, instead, is referred to several reviews (Powers, 1995, 2008; Powers and Moser, 1998). Here, focus will be only on those lesions that impact the nervous system. Abnormal elevations in fatty acids, particularly saturated very long chain (VLCFA), are characteristic of peroxisomal disorders and are proposed to be key pathogenic elements (see below). The neuropathologic lesions of human peroxisomal disorders consist of three major types: (1) abnormalities in neuronal migration or positioning, (2) abnormalities in the formation or maintenance of myelin, and (3) postdevelopmental neuronal degenerations (Powers and Moser, 1998).

4.1

Abnormalities in Neuronal Migration

The first group of neuropathologic abnormalities are those of neuronal migration or positioning which are almost restricted to the PBDs (Group 1 disorders). Abnormalities in neuronal migration are . Figure 26-7 Ballooned, striated adrenocortical cells of ALD patient. The same is seen in those with AMN. Hematoxylin-eosin, original magnification 225

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most prominent in ZS, which is characterized by a unique combination of parasylvian or centrosylvian pachygyria-polymicrogyria. Pachygyria affects medial gyri, while polymicrogyria is disposed more laterally and extends into the lateral frontal lobe and parieto-temporo-occipital region. Other foci of cortical dysgenesis, such as frontal polar pachygyria, have been observed. On coronal sectioning of the cerebrum, the abnormal gyri are seen as a thickened cortex with either excessive superficial plications or large subcortical heterotopias, the latter being more typical of pachygyric foci. Polymicrogyric cortex typically demonstrates fusion of the molecular layers with medium to large pyramidal cells destined for the deep cortex admixed with decreased numbers of layer II and III neurons in the outer cortex (> Figure 26-8). Many layer II and III neurons are . Figure 26-8 Polymicrogyric cortex of ZS. Nissl, original magnification 10

. Figure 26-9 Pachygyric cortex (left) and subcortical heterotopias of ZS. Nissl, original magnification 10

Peroxisomal disorders

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. Figure 26-10 Lamellar discontinuity and peripheral palisading of nuclei in inferior medullary olive of ZS. Phosphotungstic acid-hematoxylin, original magnification 20

located in the deep cortex and within heterotopias of subcortical white matter. The pachygyric cortical plate demonstrates similar, but more severe, architectonic changes, including the subcortical heterotopias (> Figure 26-9). Migration of all neuronal classes appears to be affected, but particularly those destined for the outer layers (Evrard et al., 1978). Less severe cerebral migratory abnormalities, usually in the form of polymicrogyria, are seen in NALD and D-bifunctional protein deficiency as diffuse, focal, or multifocal lesions that may be associated with subcortical heterotopias (reviewed in Powers, 2008). Cerebral neuronal migration problems have not been identified to date in the other peroxisomal disorders. RCDP has lacked evidence of cerebral neuronal migration lesions, except for one case that was reported prior to the era of biochemical or genetic confirmation. In this putative RCDP patient, there was a large focus of pachygyria of the posterior frontal lobe in the Rolandic region and a focus of microgyria of the frontal pole. Minor neuronal migration problems, apparently asymptomatic, are found in the form of heterotopic Purkinje cells or clusters of disordered granule-Purkinje cells (heterotaxias) in cerebellar white matter in ZS, NALD, and D-bifunctional protein deficiency. Two cases of IRD have displayed Purkinje cells abnormally distributed in the molecular layer (reviewed in Powers and Moser, 1998). Defects in neuronal positioning or terminal migration are common and usually involve the principal nuclei of the inferior medullary olives and less frequently the dentate nuclei and claustra (Evrard et al., 1978). Dysplastic inferior medullary olives, not heterotopic as in some other types of pachygyria, have been reported in ZS, RCDP, and D-bifunctional protein deficiency. They exhibit a simplification of, or discontinuities in, the normal convolutional pattern, sometimes with an apparent reduction in neuronal number and their alignment in a single row along the periphery of the olive (> Figure 26-10). A milder peripheral palisading of olivary neurons without convolutional abnormalities has been reported in IRD. Dysplastic dentate nuclei, usually seen as a simplification of the normal serpiginous pattern, has occurred in ZS and D-bifunctional protein deficiency. Neuronal loss has been reported in the dentate and olivary nuclei of two NALD patients and one D-bifunctional protein deficiency, but not dysplasias (reviewed in Powers and Moser, 1998). Fetuses at risk to develop ZS disclose neocortical migratory defects in the form of micropolygyric ripples and subtle subcortical heterotopias. Thin abnormal cortical plates with more obvious subcortical heterotopias occur later in gestation (22 weeks postmenstrual estimated gestational age). Astrocytes, neuroblasts, immature neurons, radial glia, and PAS-positive macrophages contain abnormal pleomorphic cytosomes; these include membranous cytoplasmic bodies that are electron opaque (> Figure 26-11). The

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. Figure 26-11 Fetal cerebrum from ZS (at risk). Radial glial cell from centrosylvian area contains pale, lamellated cytoplasmic inclusions and cytofilaments in a fetal ZS of 22 weeks EGA. Uranyl acetate-lead citrate, original magnification 46,500

latter seem to be typical of ZS and may represent gangliosides containing saturated VLCFA. Some neurites also exhibit lamellae and lipid profiles. Dysplastic alterations of the inferior olivary and dentate nuclei are present (Powers et al., 1989). These fetal lesions confirm the prediction that the insult (presumably metabolic) causing the neocortical migration defect is operating throughout the entire neocortical neuronal migratory period (Evrard et al., 1978). Immunohistochemical deficiencies of the cell adhesion molecule L1 (Tsuru et al., 1997) and doublecortin (Qin et al., 2000) have been reported in fetal Zellweger brains. Tissue elevations in saturated VLCFA are proposed to be a key pathogenic element through their incorporation into migrating cell membranes and acylated adhesion molecules (Powers, 1995; Powers et al., 1989). The stereotypic centrosylvian localization and the combination of both pachygria and polymicrogyria also suggest that regional tissue constraints, such as the density of radial glia and interweave of axons, act in concert with the biochemical defect to impede neuronal migration (Powers et al., 1989). Both PBDs and D-bifunctional protein deficiency display abnormalities in neuronal migration and both demonstrate increases in saturated VLCFA. However, ALD/AMN and acyl-CoA oxidase also have elevated saturated VLCFA, yet do not demonstrate the migratory lesions. Consequently, it is necessary to implicate a threshold effect in which the highest elevations in VLCFA should have the most severe and consistent neuropathologic lesion. ZS does have both the highest elevations and the most severe pachygyriamicropolygyria; hence, this postulate gains some support (Powers and Moser, 1998). Alternatively, it is also necessary to implicate the bile acid precursors that are common to the PBDs and some patients with D-bifunctional protein deficiency (Faust et al., 2005).

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26

Abnormalities in Myelin

Abnormalities of myelin, usually of CNS and particularly cerebral, are seen in diseases of both Groups 1 and 2. Adult Refsum disease is an exception in that it primarily exhibits a hypertrophic (onion bulb) demyelinating peripheral neuropathy. The lesions in CNS white matter vary greatly between these diseases, but can be divided into three basic types: (1) a nonspecific reduction in myelin volume or myelin-staining with or without a mild reactive astrocytosis (hypomyelination); (2) noninflammatory dysmyelination; and (3) inflammatory demyelination. Dysmyelination (Greek: dys – ill) refers to a genetic disorder of myelin in which either the myelin is biochemically abnormal or the myelin-forming cells have a molecular abnormality that affects either the formation or maintenance of myelin. The neuropathologic distinctions between delayed and arrested types of hypomyelination and between simple hypomyelination and hypomyelinative dysmyelination are imprecise. Demyelination (Latin: de – after) refers to the destruction, usually inflammatory and immune-mediated, of myelin that has already formed and is usually both morphologically and biochemically normal. In the Zellweger spectrum and RCDP (Group 1), as well as deficiencies of acyl-CoA oxidase or D-bifunctional protein (Group 2), the fundamental myelin problem appears to be mainly a lack or delay of CNS myelin formation, that is hypomyelination, which is seen as myelin pallor microscopically (> Figure 26-12) and reduced volume of white matter grossly (reviewed in Powers, 2008). A cerebellar preference seems to exist for D-bifunctional protein deficiency (van der Knaap and Valk, 2005). Other microscopic features may include a slight reactive astrocytosis without a macrophage or classical inflammatory (e.g., lymphocytic) response. In NALD, however, true inflammatory demyelination also occurs, as in ALD (see below), but on a more limited scale. Patients with AMN (even ‘‘pure’’ without clinical or radiological involvement of CNS white matter), either acyl-CoA oxidase or D-bifunctional protein deficiencies, and ‘‘normal’’ white matter in ALD display dysmyelinative foci (reviewed in Powers, 2008). Magnetic Resonance Spectroscopy (MRS) confirms abnormalities in ‘‘normal’’ white matter of ALD in the form of increases in choline (demyelination) and decreases in N-acetyl aspartate (axonal and/or oligodendrocytic loss) (Dubey et al., 2005). Neuropathologically, these foci consist of myelin pallor, variable axonal and oligodendrocytic loss, periodic acid-Schiff (PAS)-positive macrophages, a few reactive astrocytes, and few or no detectable lymphocytes (> Figure 26-13) (Powers et al., 2000). On the other hand, some prolonged or clinically progressive cases of acyl-CoA oxidase or D-bifunctional protein deficiencies . Figure 26-12 A paucity of myelin (blue linear staining) in cerebral white matter of ZS. The blue linear staining that is present is largely erythrocytes within microvessels. Luxol fast blue-PAS, original magnification 35

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. Figure 26-13 Focus of dysmyelination in cerebral white matter of AMN. The same can be seen in ‘‘normal’’ white matter of ALD. Mild myelin pallor and decreased number of oligodendroglial nuclei with a few weakly PAS-positive macrophages (arrows)

have had imaging findings more consistent with demyelination, including inflammation due to the finding of contrast enhancement. On personal examination of the slides from a 6-year-old patient with acyl-CoA deficiency (courtesy of P. Schwartz and E. Monuki), the patient showed as florid an inflammatory demyelination as ALD and two patients showed D-bifunctional protein deficiency of the same complementation group: the 6-month old showed only hypomyelination/dysmyelination, while the 14-month old also showed inflammatory demyelination. The possible cause of hypomyelination and dysmyelination in the PBDs and the pseudo-PBDs might be related to the abnormal elevations of fatty acids, particularly the saturated VLCFA. The causes of the cerebral hypomyelination in ZS is probably multifactorial and also due to: (1) severe neuronal migration abnormalities with a consequent reduction in the normal complement of axons, (2) marked deficiencies in plasmalogens, (3) a gliopathy due to the accumulation of abnormal unidentified lipids, (4) abnormal gangliosides lacking the proper signaling characteristics for myelination, and (5) superimposed hypoxia/ischemia/acidosis. The dysmyelinative foci mentioned above appear to be the initial myelin lesion of AMN and ALD (Powers et al., 2000). It is postulated that they are due to the incorporation of saturated VLCFA into myelin, which destabilizes it and leads to its spontaneous breakdown. Free saturated VLCFA are extremely insoluble, particularly at normal body temperature; they adversely affect the viscosity of erythrocyte and adrenocortical cell membranes, disrupt model membranes, and are toxic to a number of cell types (reviewed in Moser et al., 2001; Powers et al., 1980; Powers and Moser, 1998). In tissue their presence, when in excess, is detected as linear clearings or striations with the light microscope and clear, fine acicular or mildly curved clefts (thinner and more pliant-appearing than those of cholesterol) and lamellae ultrastructurally (> Figure 26-14), because they are removed during processing by non-polar lipid solvents such as xylene (Powers and Schaumburg, 1974; Powers, 1985). In some settings they appear to originate as slit-like dilatations of rough endoplasmic reticulum (Gordon, 1977). The toxicity of free fatty acids varies directly with their length and degree of saturation. Most of the emphasis in ALD has been on C26:0 and C24:0, but these patients accumulate a considerable amount of saturated VLCFA with chain lengths>C26 (Igarashi et al., 1976b). In tissue culture, at least, even C18:0 is cytotoxic (Gordon, 1977). The sources for the VLCFA in ALD are both endogenous and exogenous. Whereas a mild to moderate excess of VLCFA is present in all brain lipids, the greatest excess occurs in ganglioside, PLP, cholesterol ester, and

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. Figure 26-14 Ultrastructural appearance of lamellae and clear lipid profiles or clefts in adrenocortical cells of ALD. Similar to identical structures are seen in Schwann and Leydig cells, some macrophages and rare oligodendrocytes of ALD and AMN. Uranyl acetate-lead citrate, original magnification 12,000

phosphatidylcholine fractions, the latter even in ‘‘normal’’ white matter. The cholesterol esters are not found in ‘‘normal’’ white matter, but in macrophages of actively demyelinating areas (reviewed in Moser et al., 2001), which indicates that they are secondary players in the dysmyelination. VLCFA in any of the other three myelin components would be reasonable candidates to destabilize the myelin sheaths, once a certain threshold is reached. PLP may be the most appealing candidate, both for the dysmyelination and particularly for the transition to inflammatory demyelination (see below) (Powers, 1985, 1995; Ito et al., 2001). Some AMN dysmyelinative lesions remain indolent, while others elicit a mild inflammatory response and others (about 20–25%) progress to the classical confluent inflammatory demyelination of ALD (AMN-ALD) (Powers et al., 2000). An autosomal modifier gene has been believed to be responsible for the phenotypic divergence of AMN and ALD, both of which can be caused by the identical genetic defect in ABCD1 (reviewed in Moser et al., 2001). If that is true, then what factor converts AMN to AMN-ALD? There is some evidence that brain trauma may be one such factor. The inflammatory demyelination of ALD or AMN-ALD is distinctive. Neither its presence nor age of onset appears to correlate with the plasma or fibroblast levels of VLCFA or with adrenal or testicular insufficiency. Confluent and bilaterally symmetrical loss of myelin typical of most leukodystrophies involves the cerebral and, to a lesser degree, cerebellar white matter. The cerebral lesions usually begin in the parietooccipital regions, specifically the posterior limbs, and splenium, and progress more asymmetrically toward the frontal and temporal lobes. A topographic progression has not been recognized in the cerebellum, and the cerebellar lesions generally lag behind the cerebral lesions. Arcuate fibers are generally spared, except in chronic cases. The loss of myelin exceeds that of axons, but there is always substantial axonal loss. The earliest change at the advancing edge appears to be enlargement of the extracellular space (edema) with a few astrocytes and macrophages at the ultrastructural level; scant vacuoles and mild myelin swelling, reactive

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. Figure 26-15 Lymphocytic (arrow) and macrophage (arrowheads) infiltrations in cerebral demyelinative lesion of ALD. Normal white matter (WM). Hematoxylin-eosin, original magnification 20

astrocytosis, and microglial/macrophage infiltration can be appreciated at the light microscopic level. Perivascular lymphocytes, and much less frequently plasma cells, are generally found behind this advancing edge where numerous lipophages (macrophages with myelin debris) are identified (> Figure 26-15). This localization, as well as the presence of a lymphocytic perivasculitis within the corticospinal tract degeneration secondary to cerebral axonal loss, led to a pathogenetic proposal long ago: first myelin breaks down (dysmyelination), then the dominant second phase of inflammatory demyelination ensues (Schaumburg et al., 1975; Powers, 1985). The predominant cells that participate in the demyelination of ALD are reactive astrocytes, microglia/macrophages and, to a lesser extent and probably subsequently, T lymphocytes. The T cells are predominantly CD8 cytotoxic lymphocytes (CTLs) with the a/b receptor; most CD8 CTLs are also CD44 positive. They appear to kill oligodendrocytes by lysis via the granule exocytosis pathway (Powers et al., 1992; Ito et al., 2001). Oxidative damage also occurs in the lesion; more importantly, we have provided evidence for a TH-1 response as well as the presence of peroxynitrite and 4hydroxynonenal (HNE), the latter being a toxic by-product of lipid peroxidation. Both peroxynitrite and HNE can also rapidly kill cells via a nonapoptotic, lytic mechanism. Lipid peroxidation is a self-propagating process and, therefore, could be one explanation for the confluent nature of the demyelinative lesion (Powers et al., 2005). Antigenic determinant spreading and superantigens may also be operative (Ito et al., 2001). The demonstration of CD1b and CD1c molecules on astrocytes and macrophages suggests that lipid antigen presentation is a significant pathogenetic event in ALD and makes the CD1 gene a plausible candidate for a modifier gene. Both ALD lipids (cholesterol esters, ganglioside and phospholipids) and PLP contain the abnormal VLCFA and, hence, would be suitable molecules for CD1 presentation and determinant spreading (reviewed in Moser et al., 2001; Ito et al., 2001). Perivascular cells and macrophages express MHC class 1 and class 2 molecules as well as a number of chemokines and cytokines, particularly tumor necrosis factor (TNF)a. Reactive hypertrophic astrocytes in the same area can be even more immunoreactive for TNF-a than macrophages (Powers et al., 1992). While frank vasogenic edema usually is not detected microscopically, small blood vessels demonstrate upregulation of intercellular adhesion molecules (I-CAM) (Powers et al., 1992), and increased permeability of the blood brain barrier can be inferred from the characteristic contrastenhancement in the imaging studies. Within the demyelinative lesion these immunoreactivities gradually diminish and interstitial lipophages move to a perivascular location. Oligodendrocytes are reduced around the advancing edge. At the end stage of the lesion there is almost total loss of myelin and oligodendrocytes

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with a significant loss of axons. A few interstitial or perivascular lipophages may persist with isomorphic or anisomorphic fibrillary astrogliosis (Schaumburg et al., 1975; Powers, 1985). Ultrastructurally, lamellae and lipid profiles, highly similar to those in adrenocortical and Leydig cells, have been identified rarely within cells consistent with oligodendrocytes; lamellar inclusions are commonly observed within CNS macrophages containing myelin debris (Powers and Schaumburg, 1974). Spicular or trilaminar inclusions, more typical of the PBDs, also may be found within CNS macrophages. The lamellae have been postulated to be bilayers of free VLCFA, while the lipid profiles are the abnormal cholesterol esters (reviewed in Powers, 2008). To summarize, the inflammatory demyelination of ALD and AMN-ALD are the same and appears to involve an initial innate immune response to the insoluble lipids that may simulate a bacterial pathogen, in which macrophages and astrocytes produce cytokines, particularly TNF-a; this promotes a compromise of the blood-brain-barrier and lymphocytic influx. Following that, an adaptive immune response occurs in which several pathogenic elements seem to participate: a MHC-dependent TH-1 response, MHCunrestricted CD1 lipid presentation, CD8 CTLs-probably unconventional, and oxidative damage by peroxynitrite and HNE, with resultant widespread oligodendroglial lysis and loss of myelin. The free saturated VLCFA are believed to play a pivotal role and their incorporation into PLP may be the most defining alteration. Despite biochemical and ultrastructural evidence for the involvement of brain, peripheral nerve, adrenal cortex and testis, the only inflammatory site in ALD or AMN-ALD is the brain. Hence, a CNSspecific antigen, such as PLP, has a particular pathogenic appeal (Powers, 1985). How or where the genetic deficiency of ABCD1 fits into the pathogenetic scheme is still unknown, as is its precise physiological role.

4.3

Postdevelopmental Neuronal Degeneration

The next group of neuropathologic lesions shared by disorders of both Group 1 and 2 are post-developmental, non-inflammatory neuronal degenerations, of which the myeloneuropathy of AMN is the most common and most studied. The myeloneuropathy of AMN, and perhaps also that of D-bifunctional protein deficiency, displays the pattern of a central-peripheral distal (dying-back) axonopathy. The severity of the myeloneuropathy also does not seem to correlate with the duration or severity of endocrine dysfunction. The spinal lesions display equivalent losses of axons and myelin sheaths, most commonly in the gracile and corticospinal tracts (> Figure 26-16). The most severe losses are usually observed in the cervical gracile tracts and the lumbar lateral corticospinal tracts; that is, in the most distal parts of the axons furthest from the nutritive parent cell body (a ‘‘dying-back’’ pattern). The spinal lesions are bilateral, usually symmetrical, exhibit a prominent microglial infiltration, but lack reactive astrocytes or lymphocytes. Peripheral nerve lesions are milder, more variable, and more non-specific than the myelopathy, except for the diagnostic lamellar and lipid inclusions in Schwann cells that are highly similar to identical to those in adrenocortical and testicular interstitial cells (Powers and Schaumburg, 1974; Schaumburg et al., 1977; Powers, 1985; Powers et al., 2000). The largest myelinated fibers in peripheral nerve are the most severely affected. A comparable loss of the large myelinated fibers has been reported in a 4-year-old with D-bifunctional protein deficiency, both in sural and trigeminal nerves (Schroder et al., 2004). At autopsy, the lumbar dorsal root ganglia (DRG) in AMN does not exhibit appreciable neuronal loss, necrosis, apoptosis, or obvious atrophy. Nodules of Nageotte are inconspicuous. Morphometric analysis, however, reveals neuronal atrophy with a decrease in the number of the largest neurons and a corresponding increase in neurons less than 2,000 mm2, especially in the 1,500–1,999 mm2 range; this correlates with the loss of the largest myelinated fibers in the PNS. No consistent immunohistochemical differences in the size or number of specific cell types are observed. Mitochondria in the AMN DRG neurons contain lipidic inclusions at the ultrastructural level raising the possibility that, in addition to the well-known peroxisomal defect, impaired mitochondrial function may contribute to the myelopathy through a failure of ATP-dependent axoplasmic transport in AMN spinal tracts with consequent ‘‘dying-back’’ axonal degeneration (Powers et al., 2001). Another postdevelopmental neuronal degeneration is cerebellar atrophy in a few protracted RCDP cases in the age group of 4, 9 and 11 years and one 4-year-old with D-bifunctional protein deficiency and seizures. The RCDP patients did not have a significant seizure or hypoxic history. These patients have

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demonstrated diffuse cortical atrophy due to severe losses of Purkinje and granule cells with focal losses of basket cells (> Figure 26-17). The distal and superficial folia were more severely affected, and the vermis was more affected in one patient. Surviving Purkinje cells show a variety of progressive nuclear and cytoplasmic degenerative changes characterized predominantly by contraction and distortion of their nuclei. Study of this lesion with immunohistochemical stains reveals that both Purkinje and internal granule cells are lost, at least in part, by an apoptotic mechanism. Surviving Purkinje cells demonstrate prominent decreases in . Figure 26-16 AMN spinal cord. Cross section of lower cervical cord showing loss of myelin staining (loss of myelinated axons) in midline gracile tracts. Luxol fast blue-PAS, original magnification 10

. Figure 26-17 Cerebellar atrophy due to a marked loss of Purkinje cells and granular neurons in chronic RCDP. Hematoxylineosin, original magnification 20

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parvalbumin as well as other proteins involved in calcium homeostasis and energy production (Powers et al., 1999). Cerebellar atrophy probably also occurred in a patient with IRD, but the authors interpreted it as a hypoplasia (Torvik et al., 1988). Another postdevelopmental neuronal degeneration is a progressive loss of hearing that has been classified as sensorineural in type. It can be seen in ZSD, ARD, RCDP, acyl-CoA oxidase deficiency, and D-bifunctional protein deficiency. One case of NALD showed severe atrophy of the sensory epithelium and the tectorial membrane, a single case of IRD showed again severe atrophy of the sensory epithelium and the stria vascularis, and a 4-year-old patient with D-bifunctional protein deficiency exhibited a reduction of nerve fiber density in the cochlear nerve. No morphologic abnormalities have been reported in the relevant central neurons and pathways. Another postdevelopmental lesion is retinal pigmentary degeneration, which has been reported in ZSD and ARD. A few cases of ZS and NALD have been studied thoroughly and have revealed ganglion cell loss, gliosis of the nerve fiber layer and optic nerve, optic atrophy, extensive loss of outer and inner segments of photoreceptor cells, and thinning of the nuclear layer. D-bifunctional protein deficiency also may demonstrate optic atrophy. Ultrastructurally, spicular inclusions were seen in retinal macrophages, and ganglion cells contained some electron-opaque membranous cytoplasmic bodies. A single case of IRD showed comparable light microscopic changes. Therefore, in the sensorineural deafness and pigmentary retinopathy the lesion appears to reside in specialized sensory neurons (reviewed in Powers and Moser, 1998). The final postdevelopmental degenerative lesion is restricted to the neurons of the dorsal nuclei of Clarke and the lateral cuneate nuclei, second order sensory neurons, which demonstrate striated perikarya due to the accumulation or storage of lamellar and lipid profiles of abnormal cholesterol esters containing VLCFA (> Figure 26-18). This is accompanied by lamellae and lipid clefts within neighboring axonal spheroids. This CNS neuronal lesion has been reported only in ZS. It is postulated that the fundamental lesion in AMN is axonal degeneration, again perhaps related to the saturated VLCFA, particularly in gangliosides, that might interfere with normal axonal membrane-trophic factor interations (Powers et al., 2000). Electrophysiologic and, most recently, MRS data confirm the fundamental axonopathic nature of AMN (Dubey et al., 2005). It is also proposed that the cerebellar degeneration in RCDP involves the apoptotic cell death of Purkinje and granule cells due to abnormalities in calcium homeostasis that might be precipitated by the excess phytanic acid in the cerebrospinal fluid of these patients, perhaps in concert with a deficiency in tissue plasmalogens. Recently, experimental evidence . Figure 26-18 Several striated neurons (arrows) and one undergoing phagocytosis (arrowhead) in dorsal nucleus of Clarke in ZS. Luxol fast blue-PAS, original magnification 75

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has been generated that supports this proposal: phytanic acid can affect calcium homeostasis, depolarize mitochondria, generate reactive oxygen species, and kill rat hippocampal astrocytes (Kahlert et al., 2005).

Acknowledgments This Chapter is dedicated to Professor Hugo Moser, who promoted interdisciplinary research in peroxisomal disorders. Dr. Moser was initially involved in the preparation of this chapter but painfully passed away at the beginning of 2007. Ann, Hugo’s wife, was so kind and alert to have some of Hugo’s coworkers writing the different sections of the Chapter. This very nice Ann’s trait is most gratefully appreciated. Our thanks also go to Annette Snitcher.

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long-chain fatty acids: Therapeutic application in adrenoleukodystrophy. Am J Clin Nutr 40(2): 277-284. van Geel BM, Bezman L, Loes DJ, Moser HW, Raymond GV. 2001. Evolution of phenotypes in adult male patients with X-linked adrenoleukodystrophy. Ann Neurol 49(2): 186-194. van Grunsven EG, van Berkel E, Mooijer PA, Watkins PA, Moser HW, et al. 1999. Peroxisomal bifunctional protein deficiency revisited: Resolution of its true enzymatic and molecular basis. Am J Hum Genet 64(1): 99-107. Van Veldhoven PP, Mannaerts GP, Casteels M, Croes K. 1999. Hepatic alpha-oxidation of phytanic acid. A revised pathway. Adv Exp Med Biol 466: 273-281. Van Veldhoven PP, Vanhove G, Assselberghs S, Eyssen HJ, Mannaerts GP. 1992. Substrate specificities of rat liver peroxisomal acyl-CoA oxidases: Palmitoyl-CoA oxidase (inducible acyl-CoA oxidase), pristanoyl-CoA oxidase (non-inducible acyl-CoA oxidase), and trihydroxycoprostanoyl-CoA oxidase. J Biol Chem 267(28): 20065-20074. Verhoeven NM, Schor DS, Roe CR, Wanders RJ, Jakobs C. 1997. Phytanic acid alpha-oxidation in peroxisomal disorders: Studies in cultured human fibroblasts. Biochim Biophys Acta 1361(3): 281-286. Verhoeven NM, Schor DS, Roe CR, Wanders RJ, Jakobs C. 1998. Pristanic acid beta-oxidation in peroxisomal disorders: Studies in cultured human fibroblasts. Biochim Biophys Acta 1391(3): 351-356. Volpe JJ, Adams RD. 1972. Cerebro-hepato-renal syndrome of Zellweger: An inherited disorder of neuronal migration. Acta Neuropathol 20(3): 175-198. Voss A, Reinhart M, Sankarappa S, Sprecher H. 1991. The metabolism of 7,10,13,16,19-docosapentaenoic acid to 4,7,10,13,16,19-docosahexaenoic acid in rat liver is independent of a 4-desaturase. J Biol Chem 266(30): 19995-20000. Wanders RJ. 2004a. Metabolic and molecular basis of peroxisomal disorders: A review. Am J Med Genet A 126 (4): 355-375. Wanders RJ. 2004b. Peroxisomes, lipid metabolism, and peroxisomal disorders. Mol Genet Metab 83(1–2): 16-27. Wanders RJ, Boltshauser E, Steinmann B, Spycher MA, Schutgens RB, et al. 1990. Infantile phytanic acid storage disease, a disorder of peroxisome biogenesis: A case report. J Neurol Sci 98(1): 1-11. Wanders RJ, Dekker C, Hovarth VA, Schutgens RB, Tager JM, et al. 1994. Human alkyldihydroxyacetonephosphate synthase deficiency: A new peroxisomal disorder. J Inherit Metab Dis 17(3): 315-318. Wanders RJ, Denis S, Wouters F, Wirtz KW, Seedorf U. 1997. Sterol carrier protein X (SCPx) is a peroxisomal branched-chain beta-ketothiolase specifically reacting with 3-oxo-pristanoyl-CoA: A new, unique role for SCPx

in branched-chain fatty acid metabolism in peroxisomes. Biochem Biophys Res Commun 236(3): 565-569. Wanders RJ, Jansen GA, Lloyd MD. 2003. Phytanic acid alphaoxidation, new insights into an old problem: A review. Biochim Biophys Acta 1631(2): 119-135. Wanders RJ, Jansen GA, Skjeldal OH. 2001. Refsum disease, peroxisomes and phytanic acid oxidation: A review. J Neuropathol Exp Neurol 60(11): 1021-1031. Wanders RJ, Romeijn GJ. 1998. Differential deficiency of mevalonate kinase and phosphomevalonate kinase in patients with distinct defects in peroxisome biogenesis: Evidence for a major role of peroxisomes in cholesterol biosynthesis. Biochem Biophys Res Commun 247(3): 663-667. Wanders RJ, Schumacher H, Heikoop J, Schutgens RB, Tager JM. 1992. Human dihydroxyacetonephosphate acyltransferase deficiency: A new peroxisomal disorder. J Inherit Metab Dis 15(3): 389-391. Wanders RJ, Schutgens RB, Schrakamp G, Bosch van den H, Tager JM, et al. 1986. Infantile Refsum disease: Deficiency of catalase-containing particles (peroxisomes), alkyldihydroxyacetone phosphate synthase and peroxisomal beta-oxidation enzyme proteins. Eur J Pediatr 145(3): 172-175. Wanders RJ, Waterham HR. 2006. Biochemistry of mammalian peroxisomes revisited. Annu Rev Biochem 75: 295-332. Wanders RJA, Barth PG, Heymans HSA. 2001. Single peroxisomal enzyme deficiencies. Scriver CR, Beaudet AL, Sly WS, Valle D, editors. The metabolic and molecular bases of inherited disease, Eighth Edition. New York: McGraw Hill. pp. 3219-3256. Warren DS, Morrell JC, Moser HW, Valle D, Gould SJ. 1998. Identification of PEX10, the gene defective in complementation group 7 of the peroxisome-biogenesis disorders. Am J Hum Genet 63(2): 347-359. Warren DS, Wolfe BD, Gould SJ. 2000. Phenotype-genotype relationships in PEX10-deficient peroxisome biogenesis disorder patients. Hum Mutat 15(6): 509-521. Watkins PA, Chen WW, Harris CJ, Hoefler G, Hoefler S, et al. 1989. Peroxisomal bifunctional enzyme deficiency. J Clin Invest 83(3): 771-777. Watkins PA, McGuinness MC, Raymond GV, Hicks BA, Sisk JM, et al. 1995. Distinction between peroxisomal bifunctional enzyme and acyl-CoA oxidase deficiencies. Ann Neurol 38(3): 472-427. Webber KO, Hajra AK. 1993. Purification of dihydroxyacetone phosphate acyltransferase from guinea pig liver peroxisomes. Arch Biochem Biophys 300(1): 88-97. Weleber RG, Tongue AC, Kennaway NG, Budden SS, Buist NR. 1984. Ophthalmic manifestations of infantile phytanic acid storage disease. Arch Ophthalmol 102(9): 1317-1321.

Peroxisomal disorders Weller S, Cajigas I, Morrell J, Obie C, Steel G, et al. 2005. Alternative splicing suggests extended function of PEX26 in peroxisome biogenesis. Am J Hum Genet 76(6): 987-1007. Williard DE, Harmon SD, Kaduce TL, Preuss M, Moore SA, et al. 2001. Docosahexaenoic acid synthesis from n-3 polyunsaturated fatty acids in differentiated rat brain astrocytes. J Lipid Res 42(9): 1368-1376. Wilson GN, Holmes RG, Custer J, Lipkowitz JL, Stover J, et al. 1986a. Zellweger syndrome: Diagnostic assays, syndrome delineation, and potential therapy. Am J Med Genet 24(1): 69-82.

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Wilson GN, Holmes RG, Custer J, Lipkowitz JL, Stover J, et al. 1986b. Zellweger syndrome: Diagnostic assays, syndrome delineation, and potential therapy. Am J Med Genet 24(1): 69-82. Zomer AW, de Weerd WF, Langeveld J, Bosch van den H. 1993. Ether lipid synthesis: Purification and identification of alkyl dihydroxyacetone phosphate synthase from guineapig liver. Biochim Biophys Acta 1170(2): 189-196.

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S. L. Hoops . T. Kolter . K. Sandhoff

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 672

2 2.1

Sphingolipid Structure, Function, and Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 673 Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 674

3 3.1 3.2 3.2.1 3.2.2 3.2.3

Glycosphingolipid Catabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 675 Topology of Endocytosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 675 Mechanism of Lysosomal Glycosphingolipid Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 676 Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 676 Sphingolipid Activator Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 678 Vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 681

4 4.1 4.2

Pathogenesis of Sphingolipidoses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 681 Cell‐Type Specific Expression of Glycosphingolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 682 Residual Activity of the Degrading System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 683

5 5.1 5.2 5.2.1 5.2.2 5.2.3 5.2.4 5.3 5.4 5.5 5.6 5.7 5.8

Sphingolipidoses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 683 GM1‐Gangliosidosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 683 GM2‐Gangliosidoses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 684 B‐Variant of GM2‐Gangliosidoses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 684 B1‐Variant of GM2‐Gangliosidosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 685 0‐Variant of GM2‐Gangliosidosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 685 AB‐Variant of GM2‐Gangliosidosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 685 Fabry’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 686 Niemann‐Pick Disease, Types A and B . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 687 Metachromatic Leukodystrophy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 688 Gaucher’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 689 Krabbe’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 690 Farber’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 691

6

Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 691

7

Therapeutic Approaches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 692

G. Tettamanti & G. Goracci (eds.), Neural Lipids, DOI 10.1007/978-0-387-30378-9_27, # Springer ScienceþBusiness Media, LLC 2009

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Abstract: The sphingolipidoses are a group of inherited lysosomal storage diseases, which are caused by a defect in one or more sphingolipid degradation steps. The subsequent accumulation of nondegradable material in one or more organs leads to the expression of the disease. Sphingolipids are integral parts of the plasma membrane of eukaryotic cells, where they form characteristic patterns. They contribute to the glycocalix of the cell and are believed to play a role in cell adhesion phenomena, in the barrier function of the skin, in the immune system, in signal transduction processes, and during embryogenesis. After their biosynthesis at intracellular membranes, they reach the plasma membrane, where they contribute to membrane function. The constitutive degradation of glycosphingolipids takes place on the surface of intraendosomal and intralysosomal membrane structures by the action of specific acid exohydrolases, sphingolipid activator proteins (SAPs), and anionic phospholipids. The deficiency of one of the proteins involved in sphingolipid degradation can cause sphingolipidosis. The storage of nondegradable compounds, the nature of the storage material, the cell‐type specific expression of glycosphingolipids, and the residual activity of the degrading system are significant factors that contribute to the clinical manifestations of sphingolipidoses. However, the pathogenesis of these diseases is poorly understood until now. In this chapter, we summarize the characteristics of GM1‐Gangliosidosis, GM2‐Gangliosidoses (including B‐variant or Tay‐Sachs disease; 0‐variant, or Sandhoff ’s disease, and AB‐variant), Fabry’s disease, Niemann‐Pick disease (NPD) (types A and B), Metachromatic Leukodystrophy (MLD), Gaucher’s disease, Krabbe’s disease, and Farber’s disease, which are all sphinglipidoses affecting the central nervous system. Most sphingolipidoses are as yet incurable diseases. Both, the ratio of substrate influx into the lysosomes and the degradation capacity can be addressed by therapeutic approaches. The current strategies for restoration of the defective degradation capacity within the lysosome are enzyme replacement therapy (ERT), cell‐mediated therapy (CMT) including bone marrow transplantation (BMT) and cell‐mediated ‘‘cross correction,’’ gene therapy, and enzyme‐enhancement therapy with chemical chaperones. The reduction of substrate influx into the lysosomes can be achieved by substrate deprivation therapy. Patients suffering from adult forms of Gaucher’s disease and Fabry’s disease have been successfully treated by ERT. List of Abbreviations: ASA, Arylsulfatase A (ASA); ASM, Acid sphingomyelinase; BMP, Bis-(monoacylglycero)-phosphate; BMT, Bone marrow transplantation; Cer, Ceramide (N-Acylsphingosine); CMT, Cellmediated therapy; CNS, Central nervous system; Da, Dalton; ER, Endoplasmic reticulum; ERT, Enzyme replacement therapy; Gal, D-Galactose; GalNAc, N-Acetylgalactosamine; GA1, Galβ1,3GalNAcβ1,4Galβ1,4Glcβ1Cer; GA2, GalNAcβ1,4Galβ1,4Glcβ1Cer; Glc, D-Glucose; GlcCer, β-Glucosylceramide; GM1, Galβ1,3GalNAcβ1,4(NeuAcα2,3)Galβ1,4Glcβ1Cer; GM2, GalNAcβ1,4(NeuAcα2,3)Galβ1,4Glcβ1Cer; GM2AP, GM2 activator-protein; GSL, Glycosphingolipids; HSV, Herpes simplex viral vector; LacCer, Lactosylceramide; m, Meter; MDR, Multiple drug resistance; MLD, Metachromatic Leukodystrophy; mN, milliNewton; NeuAc, N-Acetylneuraminic acid; NPC, Niemann-Pick disease, type C; NPD, Niemann-Pick disease; SAP, Sphingolipid activator protein; Sap, Saposin; SPC, Sphingosylphosphocholine; TIM, Triosephosphate isomerase

1

Introduction

The sphingolipidoses are a group of inherited diseases, which are caused by mutations in genes encoding proteins involved in the lysosomal degradation of sphingolipids. The subsequent accumulation of nondegradable material in one or more organs leads to the expression of the disease. Together with mucopolysaccharidoses, mucolipidoses, glycoprotein‐, and glycogen storage diseases, sphingolipidoses belong to the lysosomal storage diseases (Gieselmann, 1995; Suzuki and Vanier, 1999; Platt and Walkley, 2004; Winchester, 2004). With a collective frequency of 1 in 8,000 live births, the lysosomal storage diseases are rare disorders, of which about 40 genetically different forms are known (Meikle et al., 1999). The endosomes and the lysosomes are the organelles, in which cellular components and foreign material are degraded by hydrolytic enzymes into their building blocks. These building blocks leave the lysosomes via diffusion or via transport by specialized proteins. Defects in any of the enzymes or cofactors

Sphingolipid‐inherited diseases of the central nervous system

27

(such as SAPs, Sandhoff et al., 2001) involved in the degradation process can lead to a lysosomal storage disease. In addition, defects in transport systems (Aula and Gahl, 2001; Gahl et al., 2001), the protective protein (d’Azzo et al., 1982, 2001), or incorrect posttranslational modifications of lysosomal proteins (I cell‐ disease/Mucolipidosis II: Kornfeld and Sly, 2001; Austin’s disease: Hopwood and Ballabio, 2001) can give rise to a lysosomal storage disease. Drug‐induced lysosomal storage has also been reported, so is sphingomyelin, stored in the brain of patients treated with tricyclic antidepressants over long time periods due to the drug‐induced degradation of acid‐sphingomyelinase (Hurwitz et al., 1994). Sphingolipid storage diseases are a subgroup of the lysosomal storage diseases. Defects in nearly every step in sphingolipid and glycosphingolipid catabolism can cause a human disease. Onset, development, and symptoms vary widely between different sphingolipidoses, but can also differ drastically within one and the same disease. In general, onset and severity of the sphingolipidoses correlate with the residual activity of the deficient degrading system. In the so‐called pseudodeficiencies, there are no visible disease symptoms, because of the high residual activity of the degrading system. Sphingolipid storage as secondary effect of impaired lysosomal function occurs in NPD, type C (Patterson et al., 2001).

2

Sphingolipid Structure, Function, and Biosynthesis

Most sphingolipids contain the amino alcohol D‐erythro‐sphingosine as a structural component, which is usually N‐linked to a fatty acid to form ceramide. In sphingomyelin and in the glycosphingolipids, a hydrophilic head group is attached to the terminal hydroxyl group of ceramide to form amphiphilic molecules (> Figure 27-1). Together with cholesterol and the glycerolipids, sphingolipids form the lipid bilayer of biological membranes (Kolter and Sandhoff, 1999).

. Figure 27-1 Ganglioside GM1, an acidic glycosphingolipid of the ganglio‐series. Structure and partial structures are given

In the plasma membrane, glycosphingolipids are believed to segregate into lipid rafts (Munro, 2003). These membrane domains are additionally enriched in glycosylphosphatidylinositol‐anchored proteins, sphingomyelin, and cholesterol, and constitute the physiological surroundings of many membrane proteins (Brown and London, 2000; van Meer and Lisman, 2002). More than 400 different glycosphingolipid structures have been characterized from natural sources. They result not only from the combination

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Sphingolipid‐inherited diseases of the central nervous system

of different sugar residues within the oligosaccharide chain, but also from differences in fatty acid chain length and degree of hydroxylation and desaturation of the ceramide backbone. The structures of glycosphingolipids can be classified into series, which are characteristic for a group of evolutionary related organisms (Kolter and Sandhoff, 1999). On the surface of eukaryotic cells, glycosphingolipids form characteristic patterns, which are species‐ and cell‐type specific and which can change with cell growth, differentiation, viral transformation, ontogenesis, and oncogenesis (Hakomori, 1981). Little is known about the specific function of individual sphingolipid species, but the conservation of their structure during evolution indicates an essential function in multicellular organisms (Kolter et al., 2000). Until now, sphingolipids are believed to play a role in the following processes:  As an integral part of the glycocalix, glycosphingolipids protect biological membranes from degradation and uncontrolled fusion.  Cell adhesion phenomena result from the interaction of glycosphingolipid‐linked carbohydrate residues with soluble lectins or with lectins on the surface of neighboring cells. Physiological examples are the blood group antigens of the ABO‐system, the extravasation of leukocytes via selectin‐glycosphingolipid interactions during inflammation, and the interaction of myelin with neuronal axons, where gangliosides on the surface of neuronal cells function as binding sites for the myelin‐associated glycoprotein (Yang et al., 1996). Glycosphingolipids can also serve as ligands for viruses (Markwell et al., 1981), bacteria (Karlsson, 1989), and toxins (Varki et al., 1993; Sandvig and van Deurs, 2002).  Ceramides of the epidermis are essential for the barrier function of the human skin (Wertz et al., 1998). In free form (Robson et al., 1994), or covalently linked to proteins of the cornified envelope (Marekov and Steinert, 1998), ceramides of the stratum corneum contribute to the water permeability barrier.  In the immune system, glycosphingolipids play not only a role as antigens (ABO‐System, Forssman), they can also stimulate the formation of autoantibodies in peripheral neuropathies like the Guillain‐ Barre´ Syndrome. SAPs are required for the presentation of lipid antigens in CD1 molecules (see later).  Sphingolipids appear to be involved in the transduction of extracellular signals into the interior of cells (Huwiler et al., 2000). Many substances such as vitamin D3, tumor necrosis factor a, g‐interferon, or interleukin 1 can cause the release of ceramide from sphingomyelin. In most cell types, ceramide mediates antimitogenic responses such as cell differentiation, cell cycle arrest, cell senescence, or apoptosis (Ruvolo, 2003). In addition, other intermediates of sphingolipid metabolism such as sphingosine, sphingosine‐1‐phosphate, and ceramide‐1‐phosphate are regarded as signal substances (Liu et al., 1999; Brindley et al., 2002).  Glycosphingolipids might play a role in embryogenesis: Genetically modified mice deficient in glucosylceramide synthase die as early as 7.5th day of embryogenesis (Yamashita et al., 1999), indicating a role of glucosylceramide or higher glycosphingolipids in this process.

2.1

Biosynthesis

The biosynthesis of glycosphingolipids occurs at intracellular membranes of the endoplasmic reticulum (ER) and the Golgi apparatus (Kolter and Sandhoff, 1999). The first steps in ceramide formation occur at the cytoplasmic face of the ER membrane (Merrill, 2002). Glucosylceramide is formed on the cytosolic face of the Golgi membrane and is subsequently translocated to the luminal face, eventually by the multiple drug resistance pump (MDR1) in the biosynthesis of neutral glycosphingolipids (De Rosa et al., 2004). Subsequent glycosylation reactions are catalyzed by glycosyltransferases in the Golgi apparatus (Maccioni et al., 2002; Giraudo et al., 2003). In the case of gangliosides, the limited specificity of some of the transferases gives rise to complex patterns on the cell surface within a combinatorial biosynthetic pathway (Kolter et al., 2002). After their biosynthesis, glycosphingolipids reach the plasma membrane through vesicular exocytotic membrane flow. With the exception of serine palmitoyltransferase deficiency (Bejaoui et al., 2002), no human diseases are known that are caused by defects in glycosphingolipid biosynthesis.

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27

Glycosphingolipid Catabolism

The constitutive process of membrane digestion requires endocytosis of parts of the plasma membrane and their subsequent transport to the lysosomal compartment. There, membrane components including glycosphingolipids are degraded into their building blocks by lysosomal hydrolases in combination with protein cofactors, the SAPs. The resulting cleavage products, monosaccharides, sialic acid, fatty acids, and sphingosine, are able to leave the lysosome.

3.1

Topology of Endocytosis

Glycosphingolipids, together with other components of the plasma membrane, are transported to the lysosomal compartment via endocytotic vesicular flow. Parts of the membrane enriched in glycosphingolipids bud into coated pits, are internalized, uncoated, and fuse with early endosomes. Subsequently, those parts of the endosomal membranes enriched in components derived from the plasma membrane invaginate and bud off into the endosomal lumen. These intraendosomal vesicles reach the lysosol after successive processes of membrane fission and fusion along the endocytic pathway (Fu¨rst and Sandhoff, 1992) and become intralysosomal vesicles or other intralysosomal membrane structures (> Figure 27-2).

. Figure 27-2 Models of endocytosis and lysosomal digestion of glycosphingolipids (GSL) in the plasma membrane (Sandhoff and Kolter, 1996). (a) Conventional model: the degradation of the GSLs of plasma membrane occurs selectively within the lysosomal membrane. (b) The alternative model: plasma membrane GSLs incorporated into the membrane of intraendosomal vesicles (multivesicular bodies) during endocytosis. The vesicles reach the lysosomal compartment when late endosomes are fused transiently with primary lysosomes and are degraded there. PM: plasma membrane

This model of membrane digestion (> Figure 27-2b; Fu¨rst and Sandhoff, 1992) was developed to explain the digestion of membrane components without alteration of the integrity of the lysosomal membrane. This perimeter membrane is covered by a thick glycocalix (Eskelinen et al., 2003), which

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protects the membranes from the attack by enzymes present in the lysosol. A series of studies (Sandhoff and Kolter, 1996; Mo¨bius et al., 1999; Mo¨bius et al., 2002) confirms this model. For example, intraendosomal and intralysosomal vesicles of this type have been observed in cells from patients with sphingolipid storage diseases such as GM1 gangliosidosis (Suzuki and Chen, 1968) or a combined SAP deficiency (Harzer et al., 1989), where they accumulate as multivesicular storage bodies.

3.2

Mechanism of Lysosomal Glycosphingolipid Degradation

The constitutive degradation of glycosphingolipids takes place on the surface of intraendosomal and intralysosomal membrane structures by the action of specific acid exohydrolases. It starts with the sequential cleavage of monosaccharide residues from the nonreducing end of the oligosaccharide part of the glycosphingolipids. Degradation of glycosphingolipids with less than four sugar residues (Wilkening et al., 2000) requires the presence of SAPs, which mediate the interaction between the membrane bound lipid substrate and the water‐soluble enzyme. In vivo, enzymatic hydrolysis of most membrane‐bound sphingolipids is also stimulated by the anionic lysosomal phospholipid bis‐(monoacylglycero)‐phosphate (BMP), which concentrates in the inner membranes of lysosomes (Mo¨bius et al., 2003).

3.2.1 Enzymes An overview of lysosomal sphingolipid degradation is shown in > Figure 27-3. Selected enzymes are described in this section. GM1‐b‐Galactosidase is a protein of 64 kDa, which is derived from an 88‐kDa precursor (review: Suzuki et al., 2001). This enzyme can occur as a homodimer (Hubbes et al., 1992) or as a ternary complex with a sialidase and the so‐called protective protein (d’Azzo et al., 1982, 2001). GM1‐b‐galactosidase catalyzes the hydrolysis of ganglioside GM1 to GM2 in the presence of either the GM2‐activator protein or Sap‐B (Wilkening et al., 2000). b‐Hexosaminidases are a group of three dimeric isoenzymes formed by a combination of two chains (a and b), differing in their substrate specificity. b‐Hexosaminidase A (ab) cleaves off terminal b‐glycosidically linked N‐acetylglucosamine‐ and N‐acetylgalactosamine residues from negatively charged and uncharged glycoconjugates. b‐Hexosaminidase A can cleave glycolipid substrates on membrane surfaces only if they extend far enough into the aqueous phase. Therefore, the degradation of ganglioside GM2, the natural substrate of b‐hexosaminidase A, occurs only in the presence of the GM2‐activator protein. The enzyme has two active sites, one on the a‐chain, and the other on the b‐chain (Kytzia and Sandhoff, 1985). b‐Hexosaminidase B (bb) predominantly cleaves uncharged substrates like glycolipid GA2 and oligosaccharides with terminal N‐Acetylhexosamine residues. The crystal structure of b‐hexosaminidase B (Maier et al., 2003; Mark et al., 2003) has been solved. b‐Hexosaminidase S (aa) is of secondary significance for GM2 degradation. It contributes to the degradation of glycosaminoglycans and sulfated glycolipids (Hepbildikler et al., 2002). a‐Galactosidase A is a homodimer of 50 kDa subunits (Desnick et al., 2001). It catalyzes the lysosomal hydrolysis of globotriaosylceramide to lactosylceramide and galactose. Glucosylceramide‐b‐Glucosidase, also called glucocerebrosidase, has a molecular weight of about 65 kDa in its glycosylated form and contains 497 amino acids (Beutler and Grabowski, 2001). It is a water‐soluble lysosomal enzyme that can associate to membranes. The X‐ray structure of this enzyme has been recently published (Dvir et al., 2003). It comprises three domains; the catalytic site is contained in a (ba)8 triosephosphate isomerase (TIM) barrel, which is characterized by eight parallel b‐strands that form a closed cylinder stabilized by hydrogen bonds. The enzyme catalyzes the hydrolysis of glucosylceramide to ceramide and glucose. It can be allosterically activated by Sap‐C (Berent and Radin, 1981; Fabbro and Grabowski, 1991) and by negatively charged phospholipids (Vaccaro et al., 1997), of which the lysosomal bis(monoacylglycero)phosphate should be of physiological relevance (Wilkening et al., 1998).

. Figure 27-3 Degradation of selected sphingolipids in the lysosomes of the cells (Kolter and Sandhoff, 1998). The eponyms of individual inherited diseases (in frame) are given. Activator proteins required for the respective degradation step in vivo are indicated. Variant AB, AB variant of GM2‐gangliosidosis (deficiency of GM2‐activator protein); Sap, saposin

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Arylsulfatase A (ASA) (von Figura and Gieselmann, 2001) belongs to the group of the sulfatases, which contain a formylglycine residue as a posttranslational modification required for the cleavage of sulfuric acid esters. ASA catalyzes the conversion of sulfatide into galactosylceramide and sulfate. For this process, the assistance of Sap‐B is required (Mehl and Jatzkewitz, 1964). Galactosylceramide‐b‐Galactosidase, also called galactocerebrosidase, is a membrane‐associated protein with a molecular weight of about 50 kDa (Wenger et al., 2001). It hydrolyzes galactosylceramide into ceramide and galactose. Sap‐A and Sap‐C are able to stimulate this degradation step in vivo. For example, mice carrying a mutation in the Sap‐A domain of the Sap‐precursor protein show accumulation of galactosylceramide and the late‐onset form of Krabbe’s disease (Matsuda et al., 2001). Acid Sphingomyelinase (ASM) (Schuchman and Desnick, 2001) is a monomeric glycoprotein with a molecular weight of 70 kDa. It has a modular structure, which includes a Sap‐like domain and a catalytic domain. Within the lysosomes, ASM catalyzes the degradation of sphingomyelin to ceramide and phosphorylcholine. In vivo, the stimulation of the enzyme by SPAs is not necessary. However, in vitro results indicate that ASM can be stimulated by lysosomal lipids and SAPs (Linke et al., 2001a). Acid Ceramidase (Moser et al., 2001) is a heterodimeric enzyme composed of an a‐subunit of 13 kDa and a b‐subunit of 40 kDa (Bernardo et al., 1995). They originate from a common 55 kDa precursor that is processed within late endosomes and lysosomes (Koch et al., 1996). Acid ceramidase catalyzes the degradation of ceramide to sphingosine and fatty acid in the lysosomes, in the presence of Sap‐C (Linke et al., 2001b) or Sap‐D (Klein et al., 1994).

3.2.2 Sphingolipid Activator Proteins To date, five SAPs are known: the GM2‐activator protein and the four saposins (Saps) ‐A, ‐B, ‐C, and ‐D, which are proteolytically derived from the sphingolipid activator protein precursor (pSap or prosaposin). The GM2‐activator is a 162 amino acid glycoprotein with a molecular weight of 17.6 kDa in its deglycosylated form (Sandhoff et al., 2001). It acts as an essential cofactor in the in vivo degradation of ganglioside GM2 by b‐hexosaminidase A (Conzelmann and Sandhoff, 1979). The inherited deficiency of the protein leads to the AB variant of GM2‐ gangliosidoses, in which lipid accumulation in neuronal cells leads to the early death of the patients (Sandhoff et al., 2001). According to the X‐ray structure (Wright and Rastinejad, 2000), the GM2‐activator contains a hydrophobic cavity to accommodate the ceramide portion of GM2, which is lined by surface loops and a single short a helix. To present ganglioside GM2 or related glycosphingolipids (e.g., GM1, Wilkening et al., 2000) to the active site of the degrading enzyme, the GM2‐activator has to insert into the bilayer of intralysosomal lipid vesicles and to lift the glycolipid out of the membrane. It can be regarded as a weak detergent with high selectivity, which forms a stoichiometric, water‐soluble glycolipid–protein complex. In vitro, it acts as a lipid transfer protein carrying lipids from donor to acceptor liposomes (Conzelmann et al., 1982). A detailed model of this mechanism based on structural information (Wright et al., 2003) and photoaffinity labeling (Wendeler et al., 2004) is shown in > Figure 27-4. The Saps or saposins A–D are four acidic, not enzymatically active, heat‐stable, and protease‐resistant glycoproteins consisting of approximately 80 amino acids (Sandhoff et al., 2001). Their molecular weights are 8–11 kDa and their isoelectric points on about pH 4.3. They contain six highly conserved cysteine residues that form three disulfide bridges (Vaccaro et al., 1995b), and one conserved N‐glycosylation site (Kishimoto et al., 1992), with the exception of Sap‐A, which has an additional one. The Saps belong to a family of saposin‐like proteins that includes NK‐Lysin, from which the three‐ dimensional structure is known (Liepinsh et al., 1997); ASM, the pore forming peptide of Entamoeba histolytica, the solution structure of which has been recently solved (Hecht et al., 2004); the small subunit of human acyloxyacylhydrolase; and the three Sap‐like domains of pulmonary surfactant protein B precursor (Munford et al., 1995). The proteins of this group carry out diverse functions, but they also share lipid binding and membrane perturbing properties. Although the four Saps share a high degree of homology (Ponting, 1994) and similar properties, they act differently and show different specificity.

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. Figure 27-4 Model for GM2‐activator stimulated hydrolysis of ganglioside GM2 by human b‐hexosaminidase A (Wendeler, 2004, unpublished). The GM2‐activator protein contains a hydrophobic cavity, dimensions of which are appropriate to accommodate the ceramide portion of GM2, lined by surface loops and a single short helix. The most flexible of the loops contains the substrate‐binding site and controls the entrance to the cavity, so that two conformations are possible: one open and one closed. The open empty activator binds to the membrane by using the hydrophobic loops and penetrates into its hydrophobic region of the bilayer. Then the lipid recognition site of the activator can interact with the substrate and its ceramide portion could move inside the hydrophobic cavity. At this point, the conformation of the full activator may change to the closed one, the complex could in this way leave the membrane, and the GM2 could be exposed to the water‐soluble enzyme to be degraded

In vivo, Sap‐A is required for the degradation of galactosylceramide by galactosylceramide‐b‐galactosidase. Mice carrying a mutation in the Sap‐A domain of the Sap‐precursor accumulate galactosylceramide and suffer from a late‐onset form of Krabbe’s disease (Matsuda et al., 2001). To date, no human disease, which is caused by the isolated defect of Sap‐A, is known. Sap‐B is the first activator protein identified (Mehl and Jatzkewitz, 1964). It mediates the degradation of sulfatide by ASA. In addition, Sap‐B is required for the degradation of globotriaosylceramide and digalactosylceramide in vivo, as demonstrated in patients with Sap‐B deficiency, where these substrates accumulate in the urine (Li et al., 1985). It is also required for the degradation of ganglioside GM3 and lactosylceramide, as reported by studies in cultured human skin fibroblast of Sap‐B deficient patients (Conzelmann et al., 1988). Like the GM2‐activator, Sap‐B can be regarded as a physiological detergent, which shows a broader specificity than the GM2‐activator. It is able to stimulate the degradation of 20 glycolipids in the presence of human, plant, and bacterial enzymes (Li et al., 1988). The crystal structure of unglycosylated human recombinant Sap‐B has been solved (Ahn et al., 2003). A shell‐like homodimer encloses a large hydrophobic cavity; the monomers are composed of four amphipathic a‐helices arranged in a long hairpin that is bent into a simple V‐shape. Like in the GM2‐activator, there are two possible conformations of the Sap‐B dimers. In addition, a similar mechanism for its action has been proposed: the open conformation should interact directly with the membrane, promote a reorganization of the lipid alkyl chains, and extract the lipid substrate accompanied by a change to the closed conformation. Thus, the substrate could be exposed to the enzyme in a water‐soluble activator–lipid complex, consistent with the previous observation that Sap‐B can act as a lipid‐transport protein (Vogel, 1991). The inherited defect of Sap‐B leads to an atypical form of MLD, with late infantile or juvenile onset (Kretz et al., 1990). The disease is characterized by accumulation of sulfatides, digalactosylceramide, and globotriaosylceramide (Sandhoff et al., 2001).

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Sap‐C has been first isolated from spleen of patients with Gaucher’s disease (Ho and O’Brien, 1971). It is required for the lysosomal degradation of glucosylceramide by glucosylceramide‐b‐glucosidase (Ho and O’Brien, 1971) and for ceramide by acid ceramidase (Klein et al., 1994). In addition, Sap‐C renders protease‐resistance to glucosylceramide b‐glucosidase inside the cell (Sun et al., 2003). Recently, the solution structure of Sap‐C has been solved (de Alba et al., 2003). This structure consists of five a‐helices, which are tightly packed to form the half of a sphere. All charged amino acids, predominantly negative ones, are solvent‐exposed, whereas the hydrophobic residues are contained within the protein core. In contrast to the mode of action of the GM2‐activator and of Sap‐B, Sap‐C performs a direct activation of the enzyme glucosylceramide‐b‐glucosidase (Ho and O’Brien, 1971; Berent and Radin, 1981; Fabbro and Grabowski, 1991). In vitro studies show that Sap‐C also supports the interaction of the enzyme with the substrate, which was embedded in vesicles containing anionic phospholipids, and that Sap‐C is able to destabilize these vesicles (Wilkening et al., 1998). Binding of Sap‐C to phospholipid vesicles is a pH‐controlled, reversible process (Vaccaro et al., 1995a). Sap‐C deficiency leads to an abnormal juvenile form of Gaucher’s disease and an accumulation of glucosylceramide (Christomanou et al., 1986; Schnabel et al., 1991). Sap‐D stimulates lysosomal ceramide degradation by acid ceramidase in cultured cells (Klein et al., 1994) and in vitro (Linke et al., 2001b). Moreover, it activates ASM, but this does not seem to be necessary for the in vivo degradation of sphingomyelin (Morimoto et al., 1988; Linke et al., 2001a). Nevertheless, the physiological function and the mode of action of Sap‐D remain unclear. It has been observed that it is able to bind to vesicles containing negatively charged lipids and to solubilize them at an appropriate pH (Ciaffoni et al., 2001). This finding may account for a more general function of Sap‐D. Inherited diseases based on the isolated defect of this cofactor are unknown to date. All four Saps are derived from a single protein, the Sap‐precursor, or prosaposin, which is proteolytically processed to the mature activator proteins in the late endosomes and lysosomes (Fu¨rst et al., 1988; O’Brien et al., 1988; Nakano et al., 1989). Prosaposin is a 70kDa glycoprotein detected mainly uncleaved in brain, heart, and muscle, whereas mature saps are mainly found in liver, lung, kidney, and spleen. The Sap‐ precursor also occurs in body fluids such as milk, semen, cerebrospinal fluid, bile, and pancreatic juice. The Sap‐precursor is either intracellular targeted to the lysosomes via mannose‐6‐phosphate receptors or sortilin (Lefrancois et al., 2003), or secreted and reendocytosed by mannose‐6‐phosphate receptors, LRP, or mannose receptors (Hiesberger et al., 1998). Until now, two different mutations in four human patients have been reported. These mutations lead to a complete deficiency of the whole Sap‐precursor protein, and consequently of all four Saps. A patient who died at the age of 16weeks, and his fetal brother carried homoallelic mutation of the start codon (Schnabel et al., 1992; Bradova et al., 1993). In the remaining patients, who belong to two unrelated families of the same district of eastern Slovakia, homoallelic deletion within the Sap‐B domain leads to a frame‐shift and a premature stop codon (Hulkova et al., 2001). The Sap‐precursor deficiency is a fatal infantile storage disorder, characterized by hepatosplenomegaly and severe neurological symptoms. In all human patients, and also in the Sap‐precursor knockout mouse (Fujita et al., 1996), there is simultaneous storage of many sphingolipids, including ceramide, glucosylceramide, lactosylceramide, ganglioside GM3, galactosylceramide, sulfatides, digalactosylceramide, and globotriaosylceramide, accompanied by an accumulation of intralysosomal membranes. This storage can be completely reversed by treatment with human Sap‐precursor, as demonstrated in prosaposin deficient fibroblasts (Burkhardt et al., 1997). Besides their function as enzyme cofactors, SAPs play an important role in lipid antigen presentation. It is now established that CD1 molecules present lipid antigens to T cells. These lipids have to be first removed from the membranes where they are embedded, to allow the loading of CD1 molecules. There is evidence that SAPs participate in this loading process. Antigen presentation in human CD1b (Winau et al., 2004) as well as human (Kang et al., 2004) and mouse CD1d (Zhou et al., 2004) have been studied: Human CD1b apparently requires especially Sap‐C to present different types of antigens. On the contrary, the glycolipid antigen bound to mouse CD1b determines whether this molecule associates with Sap‐A or the GM2‐activator. The state of our knowledge in this regard is incomplete. Further research is required to reach a better understanding of the mechanisms involved in this process.

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3.2.3 Vesicles Additional requirements for the degradation of glycolipids are given by the size, the lateral pressure, and the composition of intralysosomal vesicles. The diameter of intralysosomal vesicles has been determined in tissues from sphingolipid activator protein‐deficient patients to be in the range from 50 to 100nm (Bradova et al., 1993). The lateral surface pressure of most biological membranes is in the range between 30 and 35mNm–1 (Marsh, 1996; Maggio et al., 2002). This high lateral pressure seems to contribute to the protection of limiting membranes against inappropriate degradation, because in vitro experiments showed that the GM2‐ activator protein is only able to penetrate into a phospholipid monolayer, when the lateral pressure is below a critical value of 15–25 mNm1, depending on the lipid composition (Giehl et al., 1999). Even if no data are available on the lateral pressure of intralysosomal vesicles, the combination of size and composition can be expected to lower this pressure beyond this critical value. The lipid composition of the acidic compartments of the cell differs considerably from that of plasma membrane or limiting membranes of cellular organelles. BMP is a characteristic anionic phospholipid of the acidic compartments of the cell. It is biosynthetically formed during the degradation of phosphatidylglycerol and cardiolipin (Amidon et al., 1996; Brotherus and Renkonen, 1977), presumably on the surface of intralysosomal vesicles. It has an unusual sn1, sn10 ‐configuration, which accounts for its higher resistance to the action of phospholipases than normal phospholipids (Matsuzawa and Hostetler, 1979). Other anionic lipids like phosphatidylinositol (Kobayashi et al., 1998) and dolichol phosphate (Chojnacki and Dallner, 1988), albeit in smaller amounts than BMP, are also found within the lysosomal compartment. In addition, differences in lipid composition among the compartments of the endocytic pathway have been observed in recent experiments on human B‐lymphocytes, using an immuno‐electron microscopic method (Mo¨bius et al., 2003). On the one hand, about 80% of the cholesterol detected in the endocytic pathway is present in the recycling compartments and in internal membrane vesicles of early and late endosomes, whereas it is almost absent in lysosomes. On the other hand, the percentage of BMP increases starting from late endosomes to lysosomes, where the maximal amount is found. Moreover, BMP was shown to be mainly present in internal membranes, therefore it distinguishes these membranes from the perimeter membrane. High amounts of BMP and low amounts of cholesterol in internal lysosomal membranes appear to be required for the degradation of glycosphingolipids. The presence of BMP in these vesicles increases the ability of the GM2‐activator to solubilize lipids (Werth et al., 2001). In addition, negatively charged lysosomal lipids drastically stimulate the interfacial hydrolysis of membrane‐bound ganglioside GM1 by GM1‐b‐galactosidase (Wilkening et al., 2000), of ganglioside GM2 by b‐hexosaminidase A (Werth et al., 2001), and of the sulfated gangliotriaosylceramide SM2 by b‐hexosaminidases A and S (Hepbildikler et al., 2002) in the presence of the GM2‐activator protein. Furthermore, in the presence of Sap‐C, a drastic enhancement of glucosylceramide degradation by glucosylceramide‐b‐glucosidase produced by negatively charged model lipids such as phosphatidylserine, phosphatidylglycerol, and phosphatidic acid (Berent and Radin, 1981; Sarmientos et al., 1986; Salvioli et al., 2000) has been reported.

4

Pathogenesis of Sphingolipidoses

The sphingolipidoses are a group of inherited human diseases caused by a defect in one or more sphingolipid degradation steps, in which the nondegradable lipid substrates accumulate in different organs. The resulting diseases are usually named according to the identity of the storage material in these sphingolipidoses (Kolter and Sandhoff, 1998; Suzuki and Vanier, 1999; Winchester, 2004). The mode of inheritance is autosomal recessive with the exception of Fabry’s disease (Desnick et al., 2001). Defects for almost every step in glycosphingolipid degradation have been described. This is due to the strictly sequential degradation pathway of these compounds in humans, where the use of alternative pathways is not possible during a metabolic blockage. Lactosylceramide is an exception in this regard, because it can be degraded by two different enzyme/activator systems (Zschoche et al., 1994). Therefore, no single enzyme defect

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is known that leads to its isolated storage. Lactosylceramide accumulates together with other sphingolipids when various cofactors are absent, for example, in Sap‐precursor deficiency (Bradova et al., 1993). Most enzymes and cofactors deficient in the sphingolipidoses have been characterized and their genes have been cloned (Gieselmann, 1995). This allows the correlation between genotype and phenotype and often facilitates the diagnosis (Suzuki, 1987) of these diseases. Many mutations have been identified (Gieselmann, 1995) and animal models of most sphingolipidoses have been created by targeted disruption of the respective genes in mice ( Suzuki and Vanier, 1998; Suzuki etal., 1998). In contrast to the biochemical bases and primary causes of sphingolipidoses, which are well known, the pathogenesis of this group of diseases is poorly understood and often a correlation of genotype and phenotype is difficult to establish. For example, mutations within different genes can lead to similar clinical pictures, as in the GM2‐gangliosidoses, which can be caused by defects of genes coding the b‐hexosaminidase a‐chain, the b‐chain, or the GM2‐activator protein. On the contrary, mutations within the same structural gene can give rise to different clinical manifestations. Even patients with identical mutations within the gene that encodes ASA (Penzien et al., 1993) can show different clinical forms, due to a differing genetic background. However, a number of significant factors are known that influence the pathogenesis of sphingolipidoses. Storage of nondegradable compounds. A direct consequence of the metabolic blockage is the accumulation of the corresponding enzyme substrates. The amphiphilic storage compounds are not excreted by the affected cells and the growing amount of accumulating material might initially lead to a mechanical damage of the cell, and subsequently to apoptosis (e.g., Taniike et al., 1999; Finn et al., 2000). Nature of the storage material. The degradation disorder can be accompanied by the formation of toxic or morphologically active substances. A highly cytotoxic substance, galactosylsphingosine (psychosine), accumulates in cells of patients suffering from Krabbe’s disease. Galactosylsphingosine has also been recognized as a ligand of an orphan receptor (Im et al., 2001) and might also act by uncoupling mitochondrial respiratory chain (Tapasi et al., 1998). The most affected oligodendrocytes, which express glycosphingolipids of the gala‐ series, are destroyed before a manifestation of galactosylceramide storage can develop (Miyataki and Suzuki, 1972). Other examples of the formation of toxic substances are glucosylsphingosine in Gaucher’s disease (Nilsson and Svennerholm, 1982) or lysosulfatide in MLD (Toda et al., 1990). As a consequence of the formation of morphologically active substances, neuronal dysfunction develops (Walkley, 1998). In patients and in the mouse model of Sandhoff ’s disease (Wada et al., 2000), as well as in animal models of other gangliosidoses (Jeyakumar et al., 2003), neuroinflammatory responses, e.g., as demonstrated by microglia activation, have been reported. A molecular cause for this effect might be the rab‐activation by cholesterol, which secondarily accumulates in most sphingolipid storage diseases (Sillence and Platt, 2003).

4.1

Cell‐Type Specific Expression of Glycosphingolipids

Within sphingolipidoses, a great heterogeneity regarding the affected organs is observed. The brain is affected in many of these disorders, albeit the function of the skin or of visceral organs can be impaired in others. This is due to the cell‐type specific expression of glycosphingolipids: Lipid storage in lipidosis patients occurs especially in those cells and organs, in which the lipid substrates of the corresponding deficient degrading system are prevalently synthesized or taken up by endocytosis. For example, complex gangliosides are predominantly formed in neuronal cells (Ledeen and Salsman, 1965), so that an altered catabolism of these glycolipids leads initially to damage of the central nervous system, as can be observed in the gangliosidoses. In the case of Gaucher’s disease, storage is especially manifest in macrophages, which have large amounts of sphingolipids to degrade after phagocytosis of cells. Ceramide is also essential for normal skin function and contributes to the formation of water permeability barrier. Human patients with a complete deficiency of the degrading enzyme glucosylceramide‐b‐glucosidase in rare cases of Gaucher’s disease (collodian babies) exhibit a transepidermal water loss and die shortly after birth (Lui et al., 1988). Finally, galactosylceramide and sulfatide are characteristic lipids of myelin, so that in both Krabbe’s disease and MLD, the myelin‐forming cells are primarily affected.

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Residual Activity of the Degrading System

In most variants of a particular sphingolipidosis, there are forms with onset in early childhood and death within the first years of life, and also chronic forms with adult onset. The decisive factor in this respect is the residual activity of the gene product in the lysosomes (Conzelmann and Sandhoff, 1983/1984; Leinekugel et al., 1992). The residual activity correlates directly with the onset and severity of the disease: a complete deficiency of a lysosomal enzyme leads to an early onset and a severe course of the disease, whereas only a few percent of residual activity can be sufficient to delay the onset of the disease and cause a mild course. A critical value of enzyme activity can be calculated using the so‐called threshold theory, which has been derived from enzyme‐kinetics (Conzelmann and Sandhoff, 1983/1984). It predicts that only the decrease of the variant enzyme activity below this threshold leads to a reduced turnover rate and subsequent accumulation of the substrate. The theory has been confirmed for most sphingolipidoses (Kolter and Sandhoff, 1998). It predicts that slight elevations of enzyme activity can drastically improve the conditions of the patients, which is of relevance for the therapeutic approaches that are currently developed (see > Section4.7; Desnick and Schuchman, 2002).

5

Sphingolipidoses

5.1

GM1‐Gangliosidosis

The GM1‐gangliosidosis is caused by an inherited deficiency of the lysosomal enzyme GM1‐b‐galactosidase (Suzuki et al., 2001). According to the substrate specificity of the variant enzyme, the defect can alternatively lead to another disorder called Morquio type B disease. Storage of galactosylceramide‐b‐galactosidase substrates, galactosylceramide, and lactosylceramide, does not occur. Three clinical forms of GM1‐gangliosidosis can be distinguished. In Infantile (type 1) GM1‐Gangliosidosis, symptoms appear immediately after birth or in early infancy. Developmental arrest is followed by progressive deterioration of the nervous system. Exaggerated startle response to sounds, macular cherry‐red spots, rigospasticity associated with seizures, hepatosplenomegaly, and generalized skeletal dysplasia are usually present. Patients die within the first 2 years of life. Late infantile/juvenile form (type 2) or adult/chronic form (type 3) are characterized by progressive neurologic symptoms in children or in young adults. In the case of the late infantile/juvenile form, there are heterogeneous phenotypic expressions and no specific neurologic manifestations are known. In adults, extrapyramidal signs frequently presenting as dystonia are the most common neurologic manifestations. Dysmorphic changes are less prominent or absent in these clinical forms. Apart of ganglioside GM1, a b‐galactosidase defect can lead to accumulation of other substances such as glycolipid GA1 (Jatzkewitz and Sandhoff, 1963), oligosaccharides from glycoproteins, and intermediates of keratansulfate degradation. These substances are stored in different organs, according to their specific site of biosynthesis. GM1 storage in neurons causes the degeneration of the nervous system. Meganeurites and ectopic dendrogenesis have been observed in GM1‐gangliosidosis. This could be explained by the antineurotoxic, neuroprotective, and neurorestorative properties that have been attributed to ganglioside GM1. On the other hand, a neuroinflammatory response has been implicated in the pathogenesis of the disease (Jeyakumar et al., 2003). The severity and progression of this disease correlate with the residual enzymatic activity in cells and body fluids (Yoshida et al., 1991). Gene mutations identified in GM1 gangliosidosis and Morquio B disease include missense/nonsense mutations, duplications/insertions, and insertions causing splicing defects. Neither the type nor the location of mutations in the gene could be correlated to the phenotype of the patients. Also, mutations within the protective protein can indirectly cause accumulation of ganglioside GM1 (d’Azzo, 2001). An engineered mouse model resembling the neurological phenotype of human GM1‐gangliosidosis is available (Hahn et al., 1997). Spontaneous animal models of this disease, such as feline and canine, are also known (Suzuki et al., 2001). A treatment of this disease is not available to date, but a chemical chaperone therapy for the brain pathology has been suggested (Matsuda et al., 2003).

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Morquio type B disease is clinically a mild phenotype of Morquio A disease. In the latter one, keratansulfate accumulates as a consequence of the deficiency of the enzyme N‐acetylgalactosamine‐6‐ sulfatase. Like GM1‐gangliosidosis, Morquio type B is due to the inherited defect of GM1‐b‐galactosidase. This disease is characterized by the predominant storage of keratansulfate and oligosaccharides with terminal galactose residues. Patients show generalized skeletal dysplasia. Involvement of the nervous system or hepatosplenomegaly is not observed. Schindler’s disease (Desnick and Schindler, 2001) is an autosomal recessive inherited disease caused by deficient activity of a‐N‐acetylgalactosaminidase, also known as a‐galactosidase B. This exoglycosidase is of particular interest because it is a recently recognized member of the lysosomal complex containing a‐neuraminidase, b‐galactosidase, and the ‘‘protective protein.’’ As a consequence of the enzyme defect, sialylated and asialoglycopeptides, as well as glycosphingolipids and oligosaccharides with a‐N‐acetylgalactosaminyl residues accumulate. Clinically, three phenotypically distinct variants of the disease can be distinguished: type I is an infantile neuroaxonal dystrophy; type II is a milder, adult‐onset disorder characterized by angiokeratoma, corporis diffusum, mild intellectual impairment, and peripheral neuroaxonal degeneration; and type III is an intermediate and variable form with manifestations such as epilepsy and mental retardation in infancy or autistic behavior evident in early childhood. Characterization of the mutations in the a‐N‐acetylgalactosaminidase gene causing the three subtypes revealed a nonsense and several missense mutations.

5.2

GM2‐Gangliosidoses

The name GM2‐gangliosidoses (Sandhoff et al., 1989; Gravel et al., 2001) refers to a group of inherited diseases caused by a defect in degradation of ganglioside GM2 and related glycolipids and their subsequent lysosomal accumulation, especially in neuronal cells. The catabolism of these lipids requires the action of b‐hexosaminidases together with a protein cofactor, GM2‐activator protein. The inborn deficiency of the GM2‐activator as well as the deficiency of the a‐ or b‐chain of the b‐hexosaminidase isoenzymes leads to one of the three different variants of this disease. The names of the GM2‐gangliosidosis variants correspond to the isoenzyme remaining intact. The so‐called B‐variant is due to an a‐chain deficiency, which gives rise to deficient activity of b‐hexosaminidases A and S, with normal hexosaminidase B. Its infantile form is usually called Tay–Sachs disease. The 0‐variant, or Sandhoff ’s disease, is caused by the deficiency of the b‐chain and the resulting deficient activity of both b‐hexosaminidases, A and B. Finally, the AB‐variant is a consequence of mutations in the GM2‐activator gene and is characterized by normal b‐hexosaminidase A, b‐hexosaminidase B, and b‐hexosaminidase S activities in detergent‐containing enzyme assays, but with deficiency of the GM2‐activator.

5.2.1 B‐Variant of GM2‐Gangliosidoses From a clinical point of view, this variant can be subclassified into infantile, juvenile, chronic, and adult‐ onset forms. Infantile Tay–Sachs Disease. This disease has a higher prevalence among Ashkenazi Jews with a heterozygote frequency of 1:27. The symptoms of this disease were first described by the ophthalmologist Warren Tay in 1881. Years later, the neurologist Berhard Sachs described its morphological characteristics. Affected children are normal at birth and show first symptoms, such as mild motor weakness and increased startle reaction between 3 and 6 months of life. Weakness, hypotonia, poor head control, and decreasing attentiveness are observed and visual symptoms appear. A common finding in all cases is the so‐called cherry‐red spot in the retina of the patients. Motor, mental, and visual capabilities decline rapidly after about 10 months of age and the infant becomes less responsive. Macrocephaly and seizures are common in the second year. Further deteriorations lead to a vegetative state. Death often occurs between the second and fourth year of life and is frequently preceded by bronchopneumonia. Juvenile Form. Between 2 and 6 years, the first motor symptoms are noted and death occurs by the age of 10 to 15 years. At the end of the first decade, loss of speech, increasing spasticity, seizures, loss of vision, and

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progressive dementia are common manifestations. A vegetative state is finally reached and the patients die frequently as a consequence of intercurrent infections. Chronic Form. The onset of symptoms occurs between 2 and 5 years of age. These include abnormalities in gait and posture. With advanced age, neurological symptoms appear. Mental and verbal intelligence as well as sensory modalities remain intact. The patients can reach an age of 40 years. Adult Onset. The symptoms of this group of patients are very heterogeneous. Neurological disorders such as spinal muscular atrophy and psychoses may occur. Nevertheless, intelligence and visual capability are not affected.

5.2.2 B1‐Variant of GM2‐Gangliosidosis This variant (Kytzia et al., 1983; Suzuki and Vanier, 1991) differs enzymatically from the B‐variant by an altered substrate specificity of the mutated b‐hexosaminidase A. Although no activity is detected toward the natural substrate ganglioside GM2 and negatively charged synthetic substrates, synthetic uncharged substrates used for diagnosis are cleaved. The function of the active site of the a‐chain is defective, whereas subunit association, enzyme processing, and the activity of the b‐chain are not impaired. Homozygous patients with the B1‐mutation show the course of the juvenile disease. The late infantile form was found within the compound heterozygotes with a null allele.

5.2.3 0‐Variant of GM2‐Gangliosidosis This variant of the GM2‐gangliosidosis is also known as Sandhoff ’s disease. It was the first gangliosidosis, in which the underlying enzymatic defect was identified. In a postmortal tissue of a patient diagnosed as Tay– Sachs disease, the analyzed storage substances showed not only the presence of the negatively charged glycolipids characteristic of Tay–Sachs disease, but also of uncharged glycolipids such as glycolipid GA2 in the brain and globoside in visceral organs (Sandhoff et al., 1971). All these storage lipids had a terminal b‐ glycosidically bound N‐acetylgalactosamine residue in common. The underlying defect could be discovered during studies on the degradation of the radiolabeled storage compounds: in all tissues investigated, an almost total loss of b‐hexosaminidase A and B activity was found (Sandhoff et al., 1968). Later, the defect of both enzymes was attributed to mutations on the gene of the common b‐chain. Clinically, following forms have been reported: Infantile Sandhoff ’s Disease. In addition to the clinical and pathological manifestations present in Tay–Sachs disease, organomegaly and bone deformations also occur in this case (> Figure 27-5). Juvenile Sandhoff ’s Disease. First symptoms such as slurred speech, cerebellar ataxia, and psychomotor retardation appear at 3 to 10 years. Vision is normal in this case. Spasticity increases, and mental function deteriorates gradually. Adult Sandhoff ’s Disease. The onset of symptoms is delayed into late adult life (chronic variants). The clinical manifestations of patients with this variant are also similar to those of Tay–Sachs disease variants of corresponding ages, with the exception of additional accumulation of uncharged enzyme substrates.

5.2.4 AB‐Variant of GM2‐Gangliosidosis In this variant, the deficiency of the GM2‐activator protein (Conzelmann and Sandhoff, 1978) with normal b‐hexosaminidase A, B, and S activity leads to the accumulation of glycolipids GM2 and GA2. The clinical picture resembles that of Tay–Sachs disease with a delayed appearance of symptoms. The GM2‐gangliosidoses are histopathologically characterized by the presence of swollen neurons with massive accumulation of storage material in lysosomes (membranous cytoplasmic bodies) throughout the central and peripheral nervous system. The pathogenesis of the GM2‐gangliosidosis is not completely understood. Due to the metabolic blockage, ganglioside GM2 accumulates and precipitates together with other membrane lipids and proteins within the lysosomes of cells (Sandhoff et al., 1989), particularly in neuronal cell bodies. The resulting

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. Figure 27-5 Patient with infantile 0‐variant of GM2‐gangliosidosis (Kolter and Sandhoff, 2000)

storage granules fill the entire cytoplasm and may interfere with intracellular transport and other activities. Although the storage compounds themselves are normal and nontoxic, the toxic derivate lysoganglioside GM2 was found in patients with GM2‐gangliosidosis (Neuenhofer et al., 1986; Kobayashi et al., 1992). It is not clear, however, whether this phenomenon contributes to the pathogenesis of the disease. The formation of misconnections in the nervous tissues might be one of the causes of the neurological phenotype. Preliminary evidence suggests that the undegraded storage material can, to some extent, be recycled and reach, for example, Golgi apparatus and the plasma membrane. Such an effect might lead to changes in the pattern of gangliosides on neuronal surfaces, and to the interference with the establishment of proper connections. Besides the storage of glycolipids, other possible mechanisms have been proposed. BMT experiments suggest a complex pathogenetic mechanism that may well involve lytic compounds, for example, in the blood circulation and/or cytokines generated in the brain (Norflus et al., 1998). The severity of the disease correlates with the residual activity of the defective enzymes (Leinekugel et al., 1992). Mutations in the genes encoding the three polypeptide chains have been identified (Gravel et al., 2001). Naturally occurring animal models of GM2‐gangliosidoses have been reported in dogs, cats, and pigs. In addition, murine models of Tay–Sachs disease, Sandhoff ’s disease, and GM2‐activator deficiency have been generated using gene‐targeting techniques (Gravel et al., 2001). Unlike in the human GM2‐ gangliosidoses, the phenotypes of these murine models differ greatly from each other. These differences among the models are the result of different ganglioside degradation pathways in mice and humans. In mice, the defective step can be bypassed by the action of a sialidase (Sango et al., 1995). An inducible mouse model of the late‐onset B‐variant has been recently established (Jeyakumar et al., 2002). To date, a therapy attenuating the development or reversing the clinical manifestations of the GM2‐ gangliosidosis is not available. Treatment is restricted to supportive care and appropriate management of the intervening problems.

5.3

Fabry’s Disease

Fabry’s disease is an inborn deficiency of lysosomal b‐galactosidase A, which catalyzes the degradation of globotriaosylceramide. It is a panethnic, X‐chromosomal‐linked, inherited disorder with an estimated frequency of 1:117,000 (Meikle et al., 1999) to 1:40,000 birth (Desnick et al., 2001). The disease was first described in 1898 by dermatologists Fabry and Anderson independently from each other. It is characterized

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by the accumulation of enzyme substrates with terminal a‐glycosidically bound galactose residues, especially globotriaosylceramide. Hemizygous males have extensive deposition of this substance in the lysosomes of endothelial, perithelial, and smooth‐muscle cells of blood vessels. Many cell types in the heart, kidneys, eyes, cornea, and the autonomous nervous system are also affected. The first clinical symptoms usually occur during childhood or adolescence. These include severe pain in the extremities, vascular cutaneous lesions, hypohidrosis, and corneal and lenticular opacities. The disease develops to renal, cardiac, and/or cerebral complications of the vascular disease. These complications are the most common causes of death in the fourth or fifth decade of life. A variant of the disease characterized by a milder progression and a primary impairment of the heart muscle has been attributed to enhanced residual activity of the defective enzyme of more than 5% of normal. Heterozygous females are generally asymptomatic or have an attenuated form of this disease, although they can rarely be as severely affected as hemizygous males. Such variable phenotype is expected in females heterozygous for X‐linked diseases as a result of random X inactivation. The observed symptoms result from accumulating glycolipids in the affected tissues, the blockage of blood vessels, or both simultaneously. For example, in the kidney, the lesions are caused by glycosphingolipid accumulation in several cell types, but renal blood vessels are progressively and often extensively involved. On the other hand, in the nervous system, vascular involvement is the predominant cause of the affection. About 180 different mutations causing Fabry’s disease have been identified including partial gene rearrangements, splice‐junction defects, and point mutations (Desnick et al., 2001). An animal model of this disease has been created (Ohshima et al., 1997), allowing studies of pathogenesis and evaluation of therapeutic strategies. The treatment of Fabry’s disease by ERT is currently possible (Desnick and Schuchman, 2002). Recombinant a‐galactosidase A derived from human skin fibroblasts (Schiffman et al., 2001) or CHO‐ cells (Eng et al., 2001) are being currently used. A chemical chaperone approach with galactose was successful in a male patient of the cardiac variant of the disease (Frustaci et al., 2001), and improvements of this approach have been discussed (Fan et al., 1999; Fan, 2003).

5.4

Niemann‐Pick Disease, Types A and B

Niemann‐Pick disease (NPD), of types A and B, is a group of lysosomal storage disorders caused by the inherited deficiency of the lysosomal enzyme ASM and the accumulation of its substrate, sphingomyelin (Schuchman and Desnick, 2001). It is a panethnic disease with a higher frequency among Ashkenazi Jews (1:60 for heterozygotes). The eponym ‘‘Niemann‐Pick disease’’ refers to a heterogeneous group of lysosomal storage disorders, originally defined in terms of histology. According to the clinical manifestations, this heterogeneous disease has been classified into four subtypes, Niemann‐Pick types A–D (NPA–NPD, Crocker and Farber, 1958), and subsequently into types A–C (Crocker, 1961). In turn, on the basis of molecular and biochemical criteria, they can be grouped into two classes: those with a primary deficiency in ASM activity (types A and B), and those with defective intracellular processing and transport of LDL‐ derived cholesterol (type C). Niemann‐Pick type C disease (Patterson et al., 2001; Patterson, 2003) is a panethnic, fatal autosomal recessive, neurovisceral lipid storage disorder. It is caused by NPC1 or NPC2 mutations. In both cases, trafficking of endocytosed cholesterol is altered. This results in massive accumulation of cholesterol. Secondarily, several sphingolipids including sphingomyelin also accumulate. Type A NPD is a fatal disorder of infancy with life expectancy of 2 to 3 years. At birth, affected newborns appear to be normal, but in the first few months of life, symptoms such as hepatosplenomegaly, moderate lymphadenopathy, hypotonia, and muscular weakness appear. The observed decrease in linear growth and body weight is a consequence of feeding difficulties and splenomegaly. Other common manifestations in later stages of the disease are microcytic anemia, decreased platelet count, osteoporosis, brownish‐yellow color of the skin, and cherry‐red maculae revealed by ophthalmologic examination. Psychomotor retardation becomes evident by 6 months of age and gradually increases over the years. At an advanced state, the patient loses contact with the environment.

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Type B NPD is a phenotypically variable disorder that is usually diagnosed in childhood. Patients can reach adulthood. Commonly, enlargement of liver and/or spleen occurs, but the central nervous system is little or not involved. Progressive pulmonary infiltration causes the major disease complications in more severely affected patients. As in other sphingolipidoses, the different residual enzymatic activities of ASM accounts for the phenotypic differences between types A and B of the disease. Patients with type B show a higher residual enzyme activity than that of type A patients (Graber et al., 1994). In both types, the histologic hallmark is the formation of pathologic ‘‘foam cells’’ or ‘‘Niemann‐Pick cells.’’ Their presence is characteristic for NPD, although patients with other pathologies may have histologically similar cells. The formation of these histiocytic cells is caused by storage of sphingomyelin and other lipids in the monocyte‐macrophage system. The accumulation of sphingomyelin results from abnormal turnover of cell membranes due to ASM deficiency. Because cells of the monocyte‐macrophage system, particularly in spleen and lymph nodes, actively phagocytose sphingomyelin‐rich membranous material, they accumulate high amounts of sphingomyelin. Storage in liver, brain, kidney, and lungs also occurs. In the central nervous system of type B NPD patients, there is little or no lipid storage. The involvement of sphingolipid‐induced alterations in signal transduction is also currently discussed as a possible pathogenic mechanism of sphingolipidoses. Much interest in this field has focused on the role of ceramide as potential signaling substance in these pathways (van Blitterswijk et al., 2003). In response to extracellular stimuli, ceramide can be generated by hydrolysis of sphingomyelin, catalyzed by different sphingomyelinases with different topologies. Thus, neutral sphingomyelinase as well as ASM might play a role in the activation of the so‐called sphingomyelin pathway. Several observations are of particular relevance for NPD. ASM appears to be necessary for FAS‐induced apoptosis (De Maria et al., 1998). On the other hand, lymphoblasts of patients with NPD and also ASM knockout mice showed no ceramide formation and no apoptosis in response to irradiation (Santana et al., 1996). These results suggest that NPD patients may have subtle abnormalities in various signaling pathways and that these abnormalities could be exacerbated by stress. There is currently no clinical evidence in humans supporting this hypothesis. However, in mice deficient in ASM, a severe impairment in early host defense against Listeria monocytogenes has been demonstrated (Utermo¨hlen et al., 2003). Another factor that may contribute to the pathogenesis of the disease is the formation of sphingosylphosphocholine (SPC), a potent mitogen that can induce neurite outgrowth, and that has been shown to accumulate in type A NPD (Strasberg and Callahan, 1988). Different mutations in the ASM gene that cause types A and B NPD have been described. Three mutations, R496L, L302P, and fsP330, account for about 92% of the mutant alleles in Ashkenazi Jewish type A NPD patients. A common mutation in type B patients is the single lesion dR608 (Schuchman and Desnick, 2001). Two mouse models of NPD have been constructed by using gene‐targeting strategies (Horinouchi et al., 1995; Otterbach and Stoffel, 1995). Even if the precise targeting events differed in the two animals, they show essentially identical phenotypes. The NPD ‘‘knockout’’ mice develop features of both types A and B NPD, and are excellent models for evaluation of various therapeutic strategies. To date, there is no specific treatment available for NPD. If performed early in life, BMT has shown to have a positive effect on the clinical course of severely affected type B NPD patients (Vellodi et al., 1987).

5.5

Metachromatic Leukodystrophy

Metachromatic Leukodystrophy (MLD) is caused by the inherited deficiency of the lysosomal enzyme ASA and the subsequent accumulation of sulfatide in several tissues. The storage material accounts for the observed metachromatic staining (von Figura and Gieselmann, 2001; Gieselmann, 2003). The disease can be classified into a late infantile, a juvenile, and an adult form. The late infantile form begins between 6 months and 4 years of age, and death usually occurs about 5 years later. It begins with hypotonia, unsteady gait, and mental regression. Several abilities progressively deteriorate. Common symptoms are loss of speech, blindness, quadriparesis, peripheral neuropathy, and seizures. In the final stage before death, the child is bedridden, in a decerebrate state, and loses all contact with its surroundings.

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The juvenile form of MLD is characterized by an onset ranging from 4 to 16 years and death before 20 years. The adult form can begin after puberty up to the sixth decade of life and may extend for a few years or for decades. This form is less frequent than the previous two. In both cases, patients show gradual deterioration in school or job performance, with emotional and behavioral disturbances or psychiatric symptoms in the adult form. Other clinical manifestations are gait clumsiness, incontinence, and optic atrophy. During the final stages of this disease, the patient reaches a vegetative state. The biochemical defect in all forms of MLD is a deficiency in the enzymatic hydrolysis of sulfatides. These sulfated glycolipids occur mainly in the myelin sheaths in the white matter of the brain, in the peripheral nervous system, and in the kidney tissue. The clinical and histopathologic manifestations of MLD are fundamentally caused by a demyelination process. This phenomenon appears to be secondary to sulfatide‐ induced changes within the cells responsible for myelin maintenance, namely the oligodendrocytes in the central nervous system and the Schwann cells in the peripheral nervous system. Additionally, lysosulfatide, a cytotoxic sulfatide derivate that occurs in tissues of the patients, seems to play a role in the pathogenesis of this disease (Toda et al., 1990). More than 60 different mutations in the ASA gene are associated with the MLD phenotype. Some mutations produce a complete loss of enzyme activity and patients homozygous for this allele develop the late infantile form of the disease. On the other hand, low but definite amounts of ASA activity result in milder forms of the disease (Polten et al., 1991; Barth et al., 1993). In addition, the inherited deficiency of Sap‐B, the cofactor required for sulfatide cleavage by ASA in vivo, leads to a clinical picture similar to MLD caused by ASA deficiency. In this case, the activity of ASA toward soluble, synthetic substrates is normal (Schlote et al., 1991). A mouse model of MLD that resembles the late infantile form of the human disease has been created (Hess et al., 1996). However, it shows only a low extent of neurological and neuropathological changes, but was of value for characterization of defects in acoustic perception. To date, there is no causal therapy for MLD. The disease is invariantly lethal. In a related disease, multiple sulfatase deficiency, mucosulfatidosis or Austin’s disease, the activities of all known sulfatases are strongly reduced. This defect is due to an erroneous posttranslational modification, which is necessary for sulfate ester hydrolysis. Cells from these patients show a deficient transformation of a cysteine into a formylglycine residue (Schmidt et al., 1995). The phenotype of this disease can be described as a combination of symptoms of MLD and a mucopolysaccharidosis (Hopwood and Ballabio, 2001). Nine mutations in seven patients have been identified (Dierks et al., 2003).

5.6

Gaucher’s Disease

Gaucher’s disease is the most common of the sphingolipidoses (Beutler and Grabowski, 2001; Zhao and Grabowski, 2002). This disease is caused by the inborn deficiency of glucosylceramide‐b‐glucosidase, also called glucocerebrosidase (Brady et al., 1965; Patrick, 1965) and the subsequent accumulation of its substrate, glucosylceramide. The disease was initially described in 1882 by Gaucher, and the identification of the stored material was achieved by Aghion in 1934. Three different types of Gaucher’s disease are distinguished: Gaucher’s Disease Type I, which is the adult form, has a nonneuropathic course and is the most frequent form of this disease. It has an incidence of 1:50,000–200,000 births with a higher one among Ashkenazi Jewish population (1:1,000). The life expectancy of these patients ranges between 6 and 80 years. Gaucher’s Disease Type II, the acute form, is a very rare panethnic disease characterized by the involvement of the nervous system with early onset and a life expectancy of below 2 years. Gaucher’s Disease Type III, the subacute or juvenile form, is an intermediate variant of the other two types, with prevalence in the northern Swedish population. In this case, the neurological symptoms have a later onset and a slower development than in form II; the survival of the patients is between a few years and four decades.

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In all variants, patients may show hepatosplenomegaly, anemia, thrombocytopenia, and bone damage. The severity of these symptoms differs widely within each type and is inversely correlated with the residual enzyme activity determined in skin fibroblasts of Gaucher patients (Meivar‐Levy et al., 1994). A partial defect in glucosylceramide‐b‐glucosidase is not associated with visible skin phenotype in humans, but complete enzyme deficiency leads to the ‘‘collodion baby’’ phenotype with a severe impairment of skin function (Lui et al., 1988). Even if the enzyme activity is reduced in all cell types, the phenotype is predominantly manifested in macrophages of the reticuloendothelial system, because these cells have to degrade large amounts of glycolipids derived from the phagocytosis of erythrocytes. Due to the stored material, macrophages acquire a typical morphology, which is characterized by the enlargement of the cell and the occurrence of cytoplasmic linear inclusions. Not only the appearance of these so‐called Gaucher cells in the affected tissues, but also the inflammatory response that they induce by releasing cytokines might account for the hypertrophy of the affected organs, for cortical bone loss, and bone marrow disease. The CNS disease affecting types II and III is characterized by a progressive loss of neuronal cells. This might be caused by the accumulation of glucosylsphingosine, which apparently produces neuronal toxicity (Nilsson and Svennerholm, 1982; Orvisky et al., 2000), and might also be able to induce imflammatory response (Mizukami et al., 2002). Together with glucosylceramide accumulation, an elevation of the activity of chitotriosidase in plasma occurs. This can be used for diagnostic purposes (Hollak et al., 1994). Approximately 200 mutations at the glucosylceramide‐b‐glucosidase locus have been found in patients with Gaucher’s disease, four of which account for about 86% of the cases in the Jewish population and for 68% in non‐Jewish population. Two cases of Gaucher’s disease are known, where the cause is the absence of a sphingolipid activator protein, Sap‐C (Christomanou et al., 1986; Schnabel et al., 1991; Rafi et al., 1993). An animal model of type II of the disease has been created by targeted disruption of the glucosylceramide‐b‐glucosidase gene in mice (Tybulewicz et al., 1992). Recently, additional viable models of Gaucher’s disease have been developed by introduction of the point mutations N370S, V394L, D409H, or D409V into the mouse glucosylceramide‐b‐glucosidase locus (Xu et al., 2003). Due to the pioneering work of R. O. Brady, a causal therapy of the adult form of Gaucher’s disease (type I) is available (Brady et al., 1965). It consists of the use of glucosylceramide‐b‐glucosidase purified from placenta or recombinantly expressed, which has been modified in the carbohydrate part to contain targeting information for the mannose receptor on macrophages (Barton et al., 1990). After treatment of the patients, a normalization of the blood parameters, as well as a reduced weight of liver and spleen can be observed. For treatment of the infantile form of the disease, BMT has to be considered.

5.7

Krabbe’s Disease

Krabbe’s disease, which is also called globoid cell leukodystrophy (GCL) (Suzuki and Suzuki, 1970; Wenger et al., 2001), is caused by an inherited deficiency of the lysosomal enzyme galactosylceramide‐b‐galactosidase, also called galactocerebrosidase. In the case of Krabbe’s disease, the enzyme defect is not accompanied by substrate accumulation due to the rapid loss of galactosylceramide synthesizing cells. Together with MLD, this disorder belongs to the classical myelin disease, as the degradation of glycolipids of the white matter of the brain, sulfatide, and galactosylceramide is impaired. Clinically, this disease has two variants: infantile and late‐onset GCL. Infantile GCL usually begins between 3 and 6 months of life, the survival of the patients is less than 2 years. Clinical and pathological manifestations are exclusively observed in the nervous system. First symptoms often are irritability or hypersensitivity to external stimuli, but within a short time, severe mental and motor deteriorations occur. Commonly, patients become blind, deaf, flaccid, and hypotonic. The symptoms of Late‐Onset GCL can appear at any time after the patients are able to walk; onset ranges from a few years up to 73 years of age have been reported. Common clinical manifestations are psychomotor retardation, blindness, spastic paraparesis, and dementia. Like the infantile form, the late‐onset form is also fatal.

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The most characteristic histopathological changes are an extensive demyelination, loss of oligodendroglia, gliosis, and presence of numerous multinucleated globoid cells. These cells are hematogenous macrophages containing undigested galactosylceramide. The pathogenesis of this disease can be attributed to a combination of two phenomena: the impaired degradation of galactosylceramide, which leads to globoid cell infiltration, and the accumulation of its cytotoxic derivative galactosylsphingosine (psychosine), which causes oligodendroglial cell destruction. As a substrate of the deficient enzyme, psychosine is not degraded and reaches toxic levels. Consequently, oligodendroglial cells are destroyed. As myelin is an extension of the oligodendroglial cell membrane, rapid myelin breakdown occurs. Several mutations on the galactosylceramide‐b‐galactosidase gene have been identified. One of the most frequent ones, which is associated with the infantile form of the disease, is a deletion of exons 11–17 (Rafi et al., 1995; Selleri et al., 2000). An authentic mouse model for Krabbe’s disease, the twitcher mouse, is known. The molecular defect in these animals is a premature stop codon within the coding sequence (Sakai et al., 1996). In addition, two transgenic mouse strains, which serve as models for GCL have been created. One of them carries a mutation on the galactosylceramide‐b‐galactosidase gene leading to low enzyme activity (Luzi et al., 2001). In the other case, a mutation within the Sap‐A domain of the Sap‐precursor protein causes a deficiency of the mature activator (Matsuda et al., 2001). To date, treatment of this disease is limited to BMT in patients with only minimal neurologic involvement.

5.8

Farber’s Disease

Farber’s disease (Moser et al., 2001) is a rare disorder of sphingolipid metabolism caused by an inherited deficiency of lysosomal acid ceramidase and the storage of ceramide in the lysosomes. Usually, the symptoms of the disease appear few months after birth and death occurs within the first years of life, although patients with milder forms of the disease can reach adulthood. The most characteristic clinical manifestation is the development of painful and progressive joint deformations, subcutaneous nodules, and progressive hoarseness. Granulomas and lipid‐laden macrophages are present in these tissues. Other frequently affected organs are liver, spleen, lung, and heart. Neuronal accumulation of ceramide and gangliosides has also been reported. A classification into seven subphenotypes has been suggested. Type 1 is the classic Farber’s disease, types 2 and 3 are intermediate and mild forms, type 4 is the severe neonatal– visceral form, and type 5 is the neurologic progressive variant. Type 6 has been described in one patient and represents a coincidental combination of two distinct genetic disorders: Farber’s disease and Sandhoff ’s disease. Type 7 is caused by a lack of prosaposin, which, in turn, leads to the deficiency also of the activators Sap‐C and ‐D, which are required for ceramide degradation (Schnabel et al., 1992). The biochemical findings show combined characteristics of Farber’s disease, glucosylceramide, and galactosylceramide lipidoses. The disorder has been reclassified into diseases caused by sphingolipid activator protein deficiencies. The important role played by ceramide for skin function accounts for the striking involvement of subcutaneous tissues in Farber’s disease. Nevertheless, other manifestations in Farber’s disease remain unexplained. Even if ceramide appears to modulate a large variety of cellular functions including apoptosis, the accumulated lysosomal ceramide in Farber’s disease does not apparently cause apoptosis or any other response expected of this signaling substance. The clinical course of this disease correlates with the residual acid ceramidase activity (Levade et al., 1995). Point mutations on the acid ceramidase gene in patients of Farber’s disease have been identified (Koch et al., 1996; Ba¨r et al., 2001).

6

Diagnosis

The diagnosis of sphingolipidoses (Suzuki, 1987) is based on the evaluation of clinical symptoms and characteristic pathological manifestations, analysis of storage compounds, and especially the measurement

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of enzyme activities of individual hydrolases. Enzyme sources such as serum, leukocytes, cultured skin fibroblasts, or biopsy material can be employed. Prenatal diagnosis is carried out in amnion cells or chorion villi. The use of serum as enzyme source is common because of the easy and painless obtainment of the sample. However, not every hydrolase can be effectively determined due to the low enzyme activity present in this medium compared with the activity present in cultured cells. Moreover, variant proteins may have a lower stability in serum. Lymphocyte samples have the advantage that they are more homogenous and allow the determination of results with low dispersion. Leukocytes and skin fibroblasts are also appropriated samples. Natural or synthetic substrates with fluorogenic or chromogenic properties are frequently employed (Suzuki, 1987). The analysis of substrate accumulation can be demonstrated in cultured cells of patients by using radioactive labeled substrates or catabolic substrate precursors. For example, for Farber’s disease, radiolabeled ceramide, sulfatide, or sphingomyelin can be used. The diagnosis of sphingolipidoses can also be achieved by biosynthetic labeling of the sphingolipids in cells derived from patients and in normal cells. After a long chase period, the substrate, degradation of which is impaired, will show a higher radioactivity than in control cells. The advantage of this method is that proteins and lipids are tested in their natural and topological correct environment. Furthermore, the intensity of the radioactivity detected correlates directly with the extent of the enzyme defect. In addition, this method also allows the identification of sphingolipidoses caused by the absence of activator proteins (Klein et al., 1994).

7

Therapeutic Approaches

The theoretical basis for the therapeutic approaches toward sphingolipidoses is the threshold theory (Conzelmann and Sandhoff, 1983/1984). According to this theory, the ratio of substrate influx into the lysosomes and the degradation capacity determines the onset and severity of the diseases. Both parameters can be addressed by therapeutic approaches. The objective of some of the causal therapies of sphingolipidoses is the restoration of the defective degradation capacity within the lysosome. The current strategies for causal treatment (Kolter and Sandhoff, 1999; Desnick and Schuchman, 2002) are ERT, CMT including BMT and cell‐mediated ‘‘cross‐correction,’’ gene therapy, and enzyme‐enhancement therapy with chemical chaperones (Fan et al., 1999; Desnick and Schuchman, 2002; Fan, 2003; Kolter and Wendeler, 2003). An additional strategy for the treatment of sphingolipidoses consists in the reduction of substrate influx into the lysosomes. This can be achieved by substrate deprivation therapy (Platt et al., 1997). ERT. The aim of ERT (Neufeld, 2004) is to diminish substrate storage by the exogenous supply of the defective lysosomal enzyme. This was demonstrated for many lysosomal storage diseases in cultured cells, and in part also in animal models, where the proteins are taken up by cells by receptor‐mediated endocytosis. The enzymes are generally targeted for uptake by the mannose‐6‐phosphate receptor system, present in nearly all cells, or the mannose receptor, present in cells of the macrophage lineage. To date, ERT of sphingolipidoses is successfully applied in patients suffering adult forms of Gaucher’s disease and Fabry’s disease (Desnick and Schuchman, 2002). The blood–brain barrier prevents therapeutic enzymes from reaching neural cells and thus limits the application of this therapeutic method for the treatment of CNS diseases. CMT. In this case, cells are used as therapeutic agents (Dobrenis, 2004). There are two fundamental forms in which therapeutic benefits can be obtained. The first is to replace or compensate the defective cell population with normal equivalents, so that normal tissue or organ function might be restored. BMT and the use of neural progenitor cells are examples of this method. The second way can be through ‘‘cross‐ correction,’’ where cells release enzymes for uptake by deficient cells. This mechanism may be complemented using gene overexpression and a receptor‐mediated uptake system. Currently, there is evidence that CMT significantly alleviates pathologic manifestations of the CNS in lysosomal storage diseases. For example, in animal models, direct implantation of cells and use of BMT to

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deliver microglial/brain macrophage precursors have both resulted in cross‐correction. BMT in animal models of lysosomal storage diseases (Hoogerbrugge et al., 1988; Birkenmeier et al., 1991) has led to an improvement of the neurological symptoms and the regression of neuronal injury. Gene Therapy. This approach (Sands, 2004) is based on the insertion of a functional copy of the mutated gene into cells, which in turn produce the deficient protein. The gene therapy approach takes advantage of the process of cross‐correction. The deficient enzyme should be stably overexpressed by a few cells, secreted in high levels, and thus correct the phenotype of adjacent cells. At the cell culture level, transduction experiments have been carried out with retrovirally mediated galactosylceramide‐b‐galactosidase cDNA (Gama Sosa et al., 1996). In the mouse model of another lysosomal storage disease, Sly syndrome (mucopolysaccharidosis VII), the visceral pathology was corrected by adenovirus‐mediated gene transfer (Ohashi et al., 1997; Stein et al., 1999). Gene therapy is expected to be an effective approach for the treatment of lysosomal storage diseases with CNS involvement. Two approaches have been successfully carried out in the animal model of Tay–Sachs and Sandhoff’s diseases. The direct injection of a replication‐defective herpes simplex viral vector (HSV‐TOaHex), which efficiently transfers and expresses the b‐hexosaminidase a‐subunit cDNA, into the brain internal capsule of the Tay–Sachs mouse model has been shown to restore the b‐hexosaminidase A activity and to reduce the GM2‐ganglioside storage (Martino et al., unpublished). In the murine model of Sandhoff’s disease, the intracerebral injection of a recombinant adenoviral vector encoding the b‐subunit of b‐hexosaminidase A resulted in nearly normal levels of enzymatic activity in the entire brain. The addition of hyperosmotic concentrations of mannitol allowed an enhancement of vector diffusion (Bourgoin et al., 2003). In humans, the major difficulty in the employment of this gene therapy approach for the treatment of CNS diseases is to efficiently deliver a gene therapy vector to the brain through a systemic route. This and other obstacles remain to be overcome before this treatment can be successfully applied to affected humans. Enzyme‐Enhancement Therapy. This recently developed method relies on the use of chemical chaperones. These substances enhance the fraction of the functional variant protein. As a consequence, the degradation capacity in the lysosomes significantly increases. Substrate analogs and enzyme inhibitors have been used to stabilize the variant proteins defective in Fabry’s disease (Fan et al., 1999), Gaucher’s disease (Sawkar et al,. 2002), GM1 gangliosidosis (Matsuda et al., 2003), and GM2‐gangliosidosis (Tropak et al., 2004). Substrate Deprivation Therapy. The pathological accumulation of a substance in the lysosome occurs as long as biosynthesis continues. Using inhibitors of sphingolipid biosynthesis (Kolter and Sandhoff, 1999), the influx of substrate into the lysosomes may be reduced. The substrate deprivation approach (Platt et al., 1997) has been investigated in a genetic model. Mice suffering from Sandhoff ’s disease were crossbred with mice defective in the anabolically acting GM2/GD2‐synthase (Liu et al., 1999). The lifespan of these animals was much longer, but they developed a late‐onset neurological disease due to the accumulation of oligosaccharides. Inhibitors of glycosphingolipid biosynthesis (Kolter and Sandhoff, 1999) are suitable to reduce substrate influx into the lysosomes. For example, the ceramide glucosyltransferase inhibitor N‐butyldeoxynojirimycin has been initially investigated in the animal model of Tay–Sachs disease (Platt et al., 1997). Its efficacy could be demonstrated in a clinical trial for the treatment of human patients of Gaucher’s disease, type I (Cox et al., 2000). Substrate deprivation with N‐butyldeoxynojirimycin and BMT led to an increased survival of the mouse model of Sandhoff ’s disease. Substrate deprivation therapy, applied to patients with some residual enzymatic activity or combined with methods that restore this activity, is expected to be helpful in the treatment of sphingolipidoses.

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Mouse Models with Gene Deletions of Enzymes and Cofactors Involved in Sphingolipid Synthesis and Degradation

R. Jennemann . H.-J. Gro¨ne . H. Wiegandt . R. Sandhoff

1 1.1 1.1.1 1.1.2 1.1.3 1.1.4 1.1.5 1.1.6 1.1.7 1.1.8 1.2 1.2.1 1.2.2 1.3 1.4

Mouse Models Leading to Restricted Sphingolipid Expression in Brain . . . . . . . . . . . . . . . . . . . . . . 705 Glycosphingolipid Synthesizing Enzymes, Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 705 b1,4-GalNAc Transferase Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 708 GD3 Synthase Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 709 GM3 Synthase Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 710 Double Deficiency of b1,4-GalNAc Transferase and GD3-Synthase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 711 Double Deficiency of the b1,4-GalNAc Transferase and GM3-Synthase . . . . . . . . . . . . . . . . . . . . . . . . 712 Cell-Specific Deletion of the Glucosylceramide Synthase Gene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 713 Galactosylceramide Synthase Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 714 Cerebroside Sulfotransferase Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 715 Sphingosine Kinase Deficiencies, Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 717 Sphingosine Kinase Sphk1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 717 Sphingosine Kinase Sphk2 and Double-Deficient Mice for Sphk1/Sphk2 . . . . . . . . . . . . . . . . . . . . . . 717 Serine Palmitoyl-CoA Transferase Subunit Gene Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 718 3-Phosphoglycerate Dehydrogenase Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 719

2 2.1 2.2 2.2.1 2.2.2 2.2.3 2.3 2.3.1 2.3.2 2.3.3 2.3.4 2.3.5 2.3.6 2.3.7 2.4 2.5 2.6 2.6.1 2.6.2 2.7

Mouse Models with Deletions of Sphingolipid Degrading Enzymes, Introduction . . . . . . . . . . . 719 Deficiency of b-Galactosidase 1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 721 Deficiencies of Isoforms of b-Hexosaminidase, Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 722 HexA Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 722 HexB Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 723 HexA and HexB Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 723 Lysosomal Activator Proteins, Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 724 GM2-Activator Protein Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 724 Prosaposin Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 725 SAP-A Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 725 SAP-B Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 726 SAP-C Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 726 SAP-D Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 726 Combined SAP-C and SAP-D Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 726 a-Galactosidase A Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 727 Galactosylceramidase Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 727 b-Glucosidase Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 728 Acid b-Glucosidase, Lysosomal Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 728 b-Glucosidase, Endoplasmic Reticulum Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 730 Arylsulfatase A Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 730

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2.8 2.8.1 2.8.2 2.9 2.10

Sphingomyelinase Deficiency, Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 731 Acidic Sphingomyelinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 732 Neutral Sphingomyelinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 732 Acid and Neutral Ceramidase Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 733 Neuraminidase 1 Deficiency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 733

3

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 734

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Abstract: Sphingolipids are constituents of the cell membrane. They are believed to play critical roles in many cellular events such as signaling, in modulation of cell adhesion, and as receptor molecules in cell recognition. They may also be involved in cell differentiation, cancer development, and intracellular transport. The core constituent of sphingolipids is a sphingosine base. Acylation of the aminogroup leads to ceramide. Furthermore, carbohydrates, a phosphate, or phosphorylcholine may be linked to ceramide resulting into glycosphingolipids, ceramide-1-phosphate, and sphingomyelin. Linkage of a phosphate group to sphingosine leads to sphingosine-1-phosphate. Mouse models with gene deletions of enzymes necessary for sphingolipid synthesis and degradation have been generated. These models provide insights into the cellular functions of sphingolipids. List of Abbreviations: BMP, bis(monoacylglycero)phosphate; CNS, central nervous system; CST, Cerebroside sulfotransferase; FD, Farber Disease; GD, Gaucher disease; GLD, globoid cell leukodystrophy; GSL, Glycosphingolipids; MAG, myelin associated glycoprotein; MAL, myelin and lymphocyte protein; MBP, myelin basic protein; MBP+, myelin basic protein positive; NPD, Niemann-Pick disease; PLP, proteolipid protein; PNS, peripheral nervous system; SAP, sphingolipid activator proteins; SIMP, single insertion mutagenesis procedure; SM, sphingomyelin; SPT, Serine palmitoyl-CoA transferase

1

Mouse Models Leading to Restricted Sphingolipid Expression in Brain

The core unit of all sphingolipids is a sphingoid base. Most commonly found in mammals is the (2S,3R,4E)2-amino-octadec-4-ene-1,3-diol (D-erythro-sphingosine (d18:1)). Other sphingoid bases vary in the carbon chain length, contain additional hydroxy groups, or are saturated. Synthesis of all sphingoid bases starts with the condensation of L-serine and a long-chain acyl-CoA, the rate-limiting step of sphingolipid synthesis. Most sphingolipids contain a long-chain fatty acid linked via an amide bond, leading to ceramides. Furthermore, a monosaccharide, phosphate, or phosphorylcholine may be added to the hydroxy group in position 1 (> Figure 28‐1).

1.1 Glycosphingolipid Synthesizing Enzymes, Introduction Glycosphingolipids (GSLs) are amphipathic membrane constituents of eukaryotic cells. A major portion of the GSLs is located on the outer leaflet of the cellular plasma membrane, possibly concentrated in special domains, where they may act in the assembly of signaling molecules (Hakomori, 1990; Nagai and Tsuji, 1994; Ichikawa and Hirabayashi, 1998; van Meer, 1998), receptor activities (Yoon et al., 2006) or modulation of cell adhesion (Schnaar, 1991; Hidari et al., 1996), and differentiation (Varki, 1993). Intracellularly, GSLs may be important for protein trafficking (Sprong et al., 2001; Tamboli et al., 2005). The lipophilic ceramide anchor of GSL, integrated into the plasma membrane, is linked to a great variety of complex carbohydrate structures. Synthesis of mammalian GSLs starts either on the cytoplasmic surface of the Golgi by enzymatic transfer of UDP-activated glucose to ceramide (Futerman and Pagano, 1991; Jeckel et al., 1992) or on the luminal side of the ER with UDP-galactose (Degroote et al., 2004) (> Figure 28‐2). The resulting gluco- or galactoceramides can be elongated by enzyme-catalyzed additions of monosaccharides in the Golgi lumen. Acidic GSLs, for example, sulfatides and gangliosides, are formed by addition of sulfuric or neuraminic acid residues. Although only comparatively few GSL structures are derived from galacto-cerebroside (GalCer), hundreds of structurally different GSLs including higher sulfatides and gangliosides are known to contain the glucosylceramide core. Nervous tissue has comparably high concentrations of GSLs, in particular complex gangliosides. Thus, organs derived from neuroectoderm have been the focus of research. For decades, researchers in the GSL field have claimed that gangliosides — especially because of their high concentration in the brain — play a critical role during development and maturation of the central (CNS) and peripheral (PNS) nervous systems. Several gene disruptions — either as total deletion models or cell specific — were initiated to elucidate their function.

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. Figure 28‐1 Biosynthesis of simple sphingolipids – Membrane bound synthesis of sphingosine (Sph), Sphingosine-1-phosphate (S1P), Ceramide (Cer), ceramide-1-phosphate (C1P) and galactosylceramide (GalCer) takes place in the endoplasmic reticulum, whereas that of glucosylceramide (GlcCer) and sphingomyelin (SM) is proceeding in the Golgi apparatus (For details see recent reviews (Kolter and Sandhoff, 1999; Lahiri and Futerman, 2007)

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. Figure 28‐2 Major biosynthetic pathways of brain glycosphingolipids – The scheme was modified according to (Kolter and Sandhoff, 1999). GSL mainly expressed in brain are framed. Glycosphingolipids are abbreviated according to the recommendations of the International Union of Pure and Applied Chemistry–International Union of Biochemistry Joint Commission on Biochemical Nomenclature of Glycolipids (IUPAC–IUB Joint Commission on Biochemical Nomenclature of Glycolipids (1999) J. Mol. Biol. 286, 963-970)

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1.1.1 b1,4-GalNAc Transferase Deficiency The first genetic deletion of GSL synthesis enzymes was that of b1,4-GalNAc transferase gene (B4Galnt1), a carbohydrate transferase, responsible for the synthesis of all neuronal complex gangliosides. GalNAc transferase (EC 2.4.1.92; synonyms are B4galnt1, Galgt1, or GM2/GD2-synthase) catalyzes the transfer of UDP-activated N-acetyl-galactosamine to lactosylceramide. The gene consists of 11 exons including the noncoding exon 1 and untranslated regions 30 to exon 11. GalNAc transferase is obligatory for the synthesis of complex ‘‘brain type’’ ganglio-series gangliosides based on GA1 core structure that is highly expressed in the CNS and PNS in cells of neural origin. Gangliosides are thought to take part in various biological events such as cell adhesion, binding of bacterial toxins, and neurite outgrowth. A deletion of the gene was performed in parallel by two different groups. In order to disrupt the gene, a neomycin antibiotic selection cassette was inserted into exon 4 (Takamiya et al., 1996) or replacd exons 6, 7, and parts of exon 8 (Kawai et al., 2001). In the CNS and PNS, histopathologic reduction could not be seen. Initial investigations of the mutant mice unexpectedly detected minor neurologic abnormalities. Only a slight alteration in the conduction velocity of the tibial nerve to the somatosensory cortex was seen (Takamiya et al., 1996). However, although these mice lacked the complex gangliosides, the content of the simple gangliosides GM3 and GD3 increased in brain (> Figure 28‐3). The higher neural levels of GM3 and GD3 — present in a comparable molar quantity as the more complex gangliosides in normal brain tissue — obviously were able to compensate the absence of the complex gangliosides to a certain extent. Surprisingly examinations of these B4Galnt1/ mice revealed that males were infertile. Levels of serum testosterone in male mutant mice were reported to be very low. Multinuclear giant cells — a consequence of a dysjunction defect of spermatids — and vacuolated Sertoli cells were observed in the semiferous tubules of the testes leading to an arrest in spermatogenesis (Takamiya et al., 1998). A detailed analysis of the complex GSL pattern revealed at least 14 different testicular GSL structures that were affected by the deletion of the GalNAc transferase (Sandhoff et al., 2005a). Detailed investigations of the CNS have then shown a dysfunction of the sensory nerves measured by a reduced response to mechanical pain. In peripheral nerves, degeneration of myelinated fibers were observed. Wrapped membranous structures were found in dorsal root ganglia and sciatic nerves. In addition, numerous glial cells of the dorsal horn were enlarged and showed increased extension of . Figure 28‐3 Major biosynthetic pathways of brain gangliosides in b1,4-GaINAc transferase deficient mice – Blocks in the pathways in B4galnt1/ mice are shown. Accumulating precursor gangliosides are framed

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their processes. All observed defects were progressive. Motoric symptoms, indicated by gait disturbance, were noticed 70 weeks after birth (Sugiura et al., 2005). Furthermore, an impaired stability of paranodal junctions and disruption of ion channel clusters was associated with the absence of the complex gangliosides that colocalize with paranodal structures in PNS and CNS (Susuki et al., 2007). It has been proposed that axon/myelin contacts are stabilized by interactions preferentially between the complex gangliosides GD1a and GT1b as ligands on the axonal membranes and myelin associated glycoprotein (MAG), a member of the Siglec family of sialic acid binding lectins in the oligodendritic cell myelin membranes (Vyas et al., 2002). Disconnections between MAG and gangliosides obviously have been the cause that led to axonal degeneration of the myelinated axons and to myelination defects in peripheral nerves of B4Galnt1/ mice, along with a decrease of MAG (Sheikh et al., 1999). Phenotypic comparison of MAG/ with B4Galnt1/ mice resulted in similar neuropathological and neurologic deficits supporting the concept that MAG and complex gangliosides are essential for axon stability in the nervous system (Pan et al., 2005). All observed symptoms have been progressively with age (Chiavegatto et al., 2000; Sun et al., 2004). One important conclusion from the investigations of B4Galnt1/ mice is that simple gangliosides GM3 and GD3 cannot adequately substitute for complex gangliosides in the CNS and PNS.

1.1.2 GD3 Synthase Deficiency GD3 synthase (St8sia1; EC 2.4.99.8; also known as a-2,8-sialyltransferase (Siat8)) consists of 5 exons from which parts of exon 1 and 5 exhibit noncoding regions. The enzyme catalyzes the synthesis of the b-series gangliosides (GD3, GD2, GD1b, GT1b, and GQ1b). St8ia1 gene deletion in mice was achieved by insertion of a neomycin antibiotic-resistant cassette into exon 1 (Okada et al., 2002). St8sia1/ mice reached a normal life span, although they lacked all b-series gangliosides (> Figure 28‐4). In an initial study, hypoglossal nerves were axotomized and nerve regenerations were determined 10 weeks after cleavage. The number of surviving neurons decreased significantly and only a reduced regeneration was observed in the mutant mice as compared with the wild-type controls. These results agreed well with earlier studies that had shown a positive influence of exogenously administered ganglioside GD1b on the regeneration of hypoglossal nerves (Itoh et al., 2001a). Since mouse embryonic stem cells bearing a homozygous mutation in the St8sia1 locus undergo neuronal differentiation, b-series gangliosides might not be required for this process (Kawai et al., 1998). Additional determinations in St8sia1/ mice resulted in the description of a more distinct phenotype (Handa et al., 2005). Thus, the response to a heat stimulus or the threshold to mechanical pain showed a . Figure 28‐4 Major biosynthetic pathways of brain gangliosides in GD3 synthase deficient mice – Blocks in the pathways in St8sial/ mice are shown. Major gangliosides still expressed are framed

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significant increased sensitivity, as compared with control mice. However, formalin-induced paw edema and cFos expression, as a functional marker to identify the activity of spinal neurons, decreased significantly in St8sia1/ mice, suggesting b- and c-series gangliosides as critical mediators in the sensory nervous system in the transmission and modulation of pain. In addition, it was investigated whether b-series gangliosides represent the recognition elements for Clostridium toxins. It has been described that tetanus and botulinum toxins showed much less toxicity in B4Galnt1/ mice lacking all complex gangliosides, as compared with wild-type mice (Kitamura et al., 1999). Experiments performed in St8sia1/ mice lacking only the b-series gangliosides did not show a difference in the toxic potential of several Clostridium botulinum toxin types but revealed a high resistance against Clostridium tetani toxin suggesting that in particular the b-series gangliosides are the recognition structures for tetanus toxin.

1.1.3 GM3 Synthase Deficiency GM3 synthase, CMP-NeuAc:lactosylceramide-a2,3-sialyltransferase (St3gal5), also known as sialyltransferase 9 (Siat9), EC 2.4.99.9, is responsible for the first step in the synthesis of a-, b-, and c-series gangliosides. The gene consists of seven coding exons. To disrupt the St3gal5 gene, a neomycin selection cassette was flanked with two DNA fragments downstream from exon 4 and upstream of exon 5 of the respective gene. In this way, the targeting construct lacked the complete exon 4 (Yamashita et al., 2003). All a-, b-, and c-series gangliosides were absent in the brain of St3gal5/ mice. Synthesis of unusual gangliosides increased dramatically. They were identified as GM1b, GD1a, and GD1c, and occur only in marginal quantities in wild-type brain (Furuya et al., 1994). These three gangliosides are members of the 0-series ganglioside family with a gangliotetraosylceramide (GA1) core structure. Obviously, GA1 sialylation was still possible in St3gal5/ mice, catalyzed by different sialyltransferases that are not affected by the gene deletion (> Figure 28‐5). Phenotypic investigations in St3gal5/ mice showed neither major neurological symptoms nor histological abnormalities as compared with controls. In addition, there was no significant difference in weight gain and the animals reached a normal age. A corresponding St3gal5 point mutation detected in humans led to an infantile-onset symptomatic epilepsy syndrome, associated with stagnation in development and blindness (Simpson et al., 2004). A phenotype similar to the one observed in humans surprisingly could so far not be detected in

. Figure 28‐5 Major biosynthetic pathways of brain gangliosides in GM3 synthase deficient mice – Blocks in the pathways in St3gal5/ mice are shown. Major novel gangliosides still expressed are framed

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St3gal5/ mice. In the mutant mice, the 0-series gangliosides GM1b, GD1a, and GD1c obviously are able to substitute the a- and b-series gangliosides in their function. Presence of the 0-series gangliosides in the brain of human patients has so far not been investigated. Moreover, St3gal5/ mice displayed higher insulin sensitivity and an increase of phosphorylated insulin receptor in skeletal muscle. These observations could be well correlated with the lack of GM3 that is by far the most prominent ganglioside in muscle. This study implied that GM3 might be able to serve as a negative regulator in the insulin-signaling cascade in vivo (Yamashita et al., 2003).

1.1.4 Double Deficiency of b1,4-GalNAc Transferase and GD3-Synthase Mice with deficiencies in both genes B4Galnt1 and St8sia1 were generated in order to investigate whether neurological defects accumulate in these mice. Double mutation bearing animals were generated and investigated independently by two groups (Kawai et al., 2001; Inoue et al., 2002). Combination of B4Galnt1/St8sia1 gene deletion led to the total loss of the b-, a-, and 0-series gangliosides, except of GM3, which was the sole ganglioside in the brains of the double mutated mice (> Figure 28‐6). The life span of the double mutants was drastically reduced; almost 90% reached an age of less than 25 weeks, and these animals reacted extremely sensitively to sound stimuli that induced lethal seizures (Kawai et al., 2001). Mutant mice generated by Inoue et al. displayed significantly hypertrophic nerve axons and myelin sheaths of sciatic and trigeminal nerves. Furthermore, B4Galnt1/St8sia1/ mice developed a persistent skin injury, predominantly observed in the face and initiated by frequently scratching. Degeneration of peripheral nerves associated with reduced sensory function — also indicated by a lowered sensitivity against mechanical pain — obviously triggered self-inflicted skin wounds in these mice and led to extensive skin lesions. . Figure 28‐6 Major biosynthetic pathways of brain gangliosides in GD3 synthase and b1,4-GaINAc transferase double deficient mice – Blocks in the pathways in St8sial//B4galnt1/ mice are shown. Accumulating precursor GM3 is framed

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1.1.5 Double Deficiency of the b1,4-GalNAc Transferase and GM3-Synthase A further step toward an elucidation of functions of gangliosides in the nervous system was taken by combining B4Galnt1/ and St3gal5/ mouse models (Yamashita et al., 2005b). The double mutants were not able to synthesize common brain gangliosides, GM3 included, but surprisingly still expressed a minor proportion of glycolipids that were analyzed to belong to the sialylated lacto or neolacto series. Increase of lactosylceramide occurred and additionally SM3 the sulfate-esterified form of lactosylceramide was newly biosynthesized in brains of B4Galnt1//St3gal5/ mice (> Figure 28‐7). Close similarities in the composition of the ceramide moiety of the lactosylceramides and its sulfatester SM3 in brains of double mutated mice as compared with that of the neural gangliosides of wild-type mice suggested the presence of sulfotransferase activity not only in oligodendrocytes, but also in neuronal cells. This observation could recently be supported by Eckhardt et al. (2007) demonstrating — by using a mouse model with an overexpressed cerebroside sulfotransferase (CST) gene in combination with arylsulfatase A deficiency — that sulfatide expression in neurons is potentially possible. B4Galnt1//St3gal5/ showed a much stronger phenotype as compared with that of mice still expressing ganglioside GM3 (see earlier). Shortly after weaning, mice displayed a severe neurodegenerative disease and died early. More than 50% within the first 4 weeks after birth and only a few animals were able to survive longer than 3 months. The brain weight was significantly lower in the mutant animals after 1 month and differences increased with age. Histological investigations of the CNS showed extreme vacuolization in the white matter regions and an increased number of apoptotic cells in the cerebral cortex. Reactive astrogliosis occurred in the cortex regions adjacent to the corpus callosum, confirmed by western blot analysis showing an increased expression of the astrocyte marker GFAP. Finally, as revealed by electron microscopy, axonal degenerations, abnormal axon–glia interactions were apparent in B4Galnt1//St3gal5/ mice. Results of this study clearly implied the general importance of gangliosides for the stabilization of the CNS. . Figure 28‐7 Major biosynthetic pathways of brain gangliosides in GM3 synthase and b1,4-GaINAc transferase double deficient mice – Blocks in the pathways in St3gal5//B4galnt1/ mice are shown. Compensatory expressed glycolipids are framed

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1.1.6 Cell-Specific Deletion of the Glucosylceramide Synthase Gene Glucosylceramide synthase (Ugcg; UDP-glucose:ceramide glucosyltransferase, EC 2.4 1. 80) consists of nine exons. The enzyme catalyzes the first step of the GSL biosynthesis, the transfer of UDP-activated glucose to ceramide. In 1999, a systemic disruption of glucosylceramide synthase was achieved in mice by elimination of exon 7 (Yamashita et al., 1999). Homozygously targeted embryonic stem cells could be differentiated into endodermal-, mesodermal-, and ectodermal-like derivatives, but were not able to generate well-formed differentiated tissues in vitro. Consequently, elimination of all glucosylceramide-based glycolipids in vivo led to embryonic lethality starting at stage E7.5. It could be demonstrated that glucosylceramide-based glycolipids were vital for embryogenesis and cell differentiation. The question whether glycolipids were essential during embryonic development of the brain could not be answered. Therefore, a new approach was developed by two independent groups to generate cell-specific deletion of Ugcg and the absence of glucosylceramide-derived glycolipids using the Cre/loxP system (Jennemann et al., 2005; Yamashita et al., 2005a). In both reports, exons 6–8 of the gene were flanked with loxP sites. Mice were generated that carried the nestin gene promoter driven cre-recombinase transgene in combination with a ‘‘floxed’’ and null allele or alternatively two ‘‘floxed’’ alleles of Ugcg. Nestin — an intermediate filament protein of neuroepithelial stem cells — is expressed early in neuronal cells during embryonic development from day E9.5. Glucosylceramide-based glycolipids were almost completely absent in the brain of Ugcgnull/flox//NesCre mice already at embryonic stage E14.5 (Jennemann et al., 2005) (> Figure 28‐8). GlcCer synthase genedeficient mice showed no significant accumulation of ceramide in the brain. Sphingomyelins (SMs) increased quantitatively approximately to the extent that gangliosides were lacking. Nevertheless, shortly after birth, mice developed severe neurologic dysfunctions indicated by a shuffling gait and ataxia. Axonal branching of the Purkinje neurons in the cerebellum was significantly reduced. Peripheral nerves showed hypertrophic axons and broadened myelin sheaths, combined with disorderly arrangement of the myelin. The branching defects observed in vivo were confirmed in primary cultures of hippocampal neurons in vitro, which exhibited diminished dendritic complexity and underwent early pruning. All animals died within 3 weeks after birth (Jennemann et al., 2005). Mice with brain-specific deletion of Ugcg described by Yamashita et al. (2005a) still contained amounts of glucosylceramide-based glycolipids (approximately 25% of normal brain). Variances in the amounts of remaining glycolipid, which led to a milder phenotype, might be explained by a low activity of the . Figure 28‐8 Major biosynthetic pathways of brain gangliosides in neural GlcCer synthase deficient mice – Blocks in the pathways in Ugcgnull/flox/NesCre mice are shown. Compensatory sphingomyelin (SM) increases and is framed

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nestin-cre mice. However, also in this study, neurologic abnormal behavior and severe degenerations of the Purkinje cells in the cerebellum in particular in aged mice have been recognized. Neural-specific deletions of glycolipids, including the gangliosides, indicated that they may not serve functions essential for early brain development, but are required for neuron differentiation, brain maturation, and stability after birth. Glycolipids may be essential for differentiation of several different cell types. A recent study has demonstrated that absence of Ugcg cell specifically in the epidermis leads to the functional breakdown of the water barrier in the skin (Jennemann et al., 2007).

1.1.7 Galactosylceramide Synthase Deficiency UDP-galactose:ceramide galactosyltransferase (Ugt8a, also known as cerebroside galactosyltransferase (Cgt), EC 2.4.1.45) is the key enzyme in the biosynthesis of galactosylcerebroside and catalyzes the transfer from UDP-galactose to ceramide. The enzyme contains six exons from which exon 1 remains completely untranslated. Exon 6 contains an additional untranslated region. Synthesis of galactosylceramide (GalCer) is performed on the luminal side of the endoplasmic reticulum, which is opposite to glucosylceramide synthase acting on the cytoplasmic surface of the Golgi apparatus. GalCer and its sulfated form sulfatide (SM4s) are, in contrast to the glucosylceramide derivatives, expressed in neuronal cells in small amounts. Its main distribution is in oligodendrocytes in the CNS and in Schwann cells of the PNS (von Figura et al., 2001; Pernber et al., 2002; Molander-Melin et al., 2004). Oligodendrocytes and Schwann cells are forming the multilamellar, spirally wrapped myelin sheaths that are serving as an axonal insulator in the internodes of nerve axons. Myelin consists mainly of lipids (more than 70% of its dry weight). Major fractions of these lipids (approximately 27%) are GalCer and SM4s. This abundance has led to the assumption that GalCer and SM4s play important roles for myelinization processes of nerve axons. Deletion of the enzyme was performed by two different groups in 1996 (Bosio et al., 1996; Coetzee et al., 1996). Both groups deleted the Ugt8a gene by placing a neomycin gene selection cassette into a Kpn I site of the first coding exon. Absence of GalCer synthase activity and consequently lack of GalCer and its sulfated form SM4s have been shown in both models (> Figure 28‐9). Ugt8a/ mice expressed instead, to a lesser extent glucosylceramide (GlcCer) and its sulfated derivative, as well as SM. The ‘‘neo’’-biosynthesized sphingolipids expressed to a major proportion a-hydroxy fatty acids in their ceramide moiety. This change in the fatty acid composition might be correlated with the biophysical properties of the myelin sheaths when galactosphingolipids are lacking, which then may lead to an increased fluidity and permeability of the myelin lipid bilayer (Bosio et al., 1998). Ugt8a/ mice were of smaller size than control littermates. Although initial ultrastructural analysis of myelin demonstrated only slightly thinner sheaths in the ventral area of the spinal cord, oligodendrocytes and Schwann cells of mice had partly lost their insulator function. Velocity of nerve conduction was reduced to that of unmyelinated nerve axons (Bosio et al., 1996; Coetzee et al., 1996). Ugt8a/ mice showed a conspicuous gait pattern due to weakness of their hind and front legs progressing to a severe paralysis at the age of 3 weeks. These symptoms got worse with age and led to death of Ugt8a/ mice 25–30 days after birth (Bosio et al., 1996). In the mouse model generated by Coetzee et al. (1998), similar symptoms were described but some animals survived longer than 90 days. Therefore, adult animals could be investigated for potential defects caused by the absence of GalCer synthase. A few abnormal myelination events were observed at postnatal day P30 restricted to midbrain and hindbrain, but occurred throughout the whole CNS at P90, with a high regional variability in the extent of demyelination. Activated microglia cells and astrocytes (astrogliosis) occurred in particular in older Ugt8a/ mice. In wild-type mice an age-dependent switch of the alternatively spliced isoforms of the MAG from L-MAG to S-MAG mRNA, which increased after postnatal day 90, was observed. In contrast, Ugt8a/ mice still showed a pronounced expression of L-MAG mRNA with increasing age at comparable levels as S-MAG. This observation was explained by alterations in oligodendrocyte maturation status in Ugt8a/ mice, caused by the absence of galactosylceramide-derived glycolipids (Coetzee et al., 1998). Detailed electron

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. Figure 28‐9 Biosynthetic pathways of galactosylceramide – Blocks in the pathways in Ugt8a/ mice is shown. Compensatory expressed glycolipids are framed

microscopic investigations in the CNS of Ugt8a/ mice revealed altered nodal lengths and abundance in heminodes. In paranodal structures, absence of regularly arrayed densities known as transverse bands and occurrence of reversed lateral loops were seen. However, the PNS did not seem to be affected (Dupree et al., 1998). In summary, GalCer and its sulfated derivative SM4s, lacking in Ugt8a/ mice, appear to be essential for long-term maintenance of myelin structure and function, as well as node and paranode formation and axo–glial interactions.

1.1.8 Cerebroside Sulfotransferase Deficiency CST (EC 2.8.2.11) encodes the gene responsible for the 3-O-sulfation of galactosylceramide (Gal3st1). Sulfatide (HSO3-3-galactosylceramide, SM4s), accompanied by GalCer, is the most prominent glycolipid of oligodendrocytes in the CNS and of Schwann cells in the PNS. SM4s is additionally expressed in high amounts in kidney, gastric mucosa, lung, and endometrium of various mammals. The SM4s-related glycerolipid seminolipid (HSO3-3-monogalactosylalkylacylglycerol, SM4g) represents the major glycolipid in testicular tissue. Sulfatides have been surmised to play important roles in a variety of physiological functions in which they may act as potential binding partners for extracellular matrix proteins, chemokines (Sandhoff et al., 2005b), cellular receptors, and microorganisms. They may also be involved in blood coagulation, complement activation, and cation transport (Vos et al., 1994; Ishizuka, 1997). Gal3st1 contains multiple exons 1 and two coding exons from which exon 2 encodes the transmembrane region of the gene, and exon 3 constitutes the catalytic domain followed by an 30 untranslated region. Gene deletion was achieved by replacing parts of exons 2 and 3 by a neomycin selection cassette (Honke et al., 2002). In this report it could convincingly be shown that both CST enzyme activity and resulting products were absent (> Figure 28‐10). Moreover, not only the SM4s in the brain, but also its glycerol derivative, seminolipid, was eliminated in Gal3st1/ mice, whereby its progenitor monogalactosylalkylacylglycerol (GalEAG) accumulated. Animals deficient in CST activity were born according to Mendelian rules but

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. Figure 28‐10 Biosynthetic pathways of cerebroside sulfotransferase – Blocks in the pathways in Gal3st1/ mice is shown. Compensatory expressed progenitor glycolipid galactosylceramide (GalCer) is framed

displayed severe neurological defects after birth indicated by pronounced tremor and progressive ataxia, as well as hind and forelimb weakness. Myelin vacuolization was observed in the cerebellar white matter, diencephalon, brainstem, and the spinal anterior column. Abnormal paranodal junctions were also reported (Honke et al., 2002). More detailed investigations revealed aberrantly localized K+ ion channels on the paranodes of spinal cord. Na+ ion channel clusters were stained in the nodes of Ranvier just as in wild-type controls but exhibited significantly longer shapes with lower intensities in Gal3st1/ mice; these lesions were more pronounced with age. Although the total count of K+ and Na+ ion channel clusters in controls were almost not influenced by age, numbers of clusters in Gal3st1-deficient mice significantly decreased starting 6 weeks after birth. Still, significant demyelination processes were not described (Ishibashi et al., 2002). In contrast, recent investigations of myelin and axon structure revealed significant alterations in Gal3st1/ mice indicated by degenerating myelin sheaths and redundant and uncompacted myelin. Deteriorated paranode structures and increasing nodal length were observed. Disorganized and everted lateral loops were also found in this study (Marcus et al., 2006). Gal3st1/ male mice were infertile and the weight of their testes was almost one third of that from control males, and the epididymis contained no sperm. Histology of the testis showed that haploid cells and secondary spermatocytes were absent from the seminiferous tubules. Late spermatocytes were also affected showing nuclear condensations and formed multinucleated cells at the metaphase of the first meiotic division. In addition, an increased apoptosis was observed in germ cells (Honke et al., 2002). The kidney likewise contains major amounts of sulfatides. Investigations of kidneys from Gal3st1/ mice showed regular morphology. However, a decreased L-selectin-dependent monocyte infiltration after ureteral obstruction of the kidney was observed in Gal3st1/ mice (Ogawa et al., 2004), perhaps because of an effect of sulfatides, synthesized in epithelial cells on the adhesive properties of peritubular endothelia. Investigations on the function of sulfatides in oligodendrocyte differentiation have been performed. Using primary cultures from Gal3st1/ mice, increased oligodendrocyte maturation and a rise of positive myelin basic protein positive (MBP+) cells was demonstrated in vitro. These results in cell culture were confirmed by in situ hybridization against proteolipid protein (PLP) mRNA, which was used as marker

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protein indicating oligodendrocyte maturation status in vivo (Hirahara et al., 2004). Results of this study revealed that sulfatide (SM4s) may act as a negative regulator for oligodendrocyte differentiation. The data from Gal3st1/ mice seem to indicate that sulfatides maintain structure and function of myelin, are involved in paranodal and ion channel cluster formation, foster oligodendrocyte maturation, and are necessary for spermatogenesis.

1.2 Sphingosine Kinase Deficiencies, Introduction Sphingosine kinase (SphK, EC 2.7.1.91) catalyzes the transfer of one phosphate group to sphingosine to form sphingosine-1-phosphate (S1P; for synthesis pathway, see > Figure 28‐1). S1P is a signaling molecule involved in numerous events in the cell such as survival, differentiation, growth, migration, calcium homeostasis, and angiogenesis (Spiegel and Milstien, 2003). S1P signaling through S1P receptors, S1P1 to S1P5, a family of G protein-coupled receptors, induces activation of several intracellular signal transduction pathways. Some of the proteins involved are protein kinase C, phosphoinositide-3 kinase, the small GTP binding proteins Rho and Rac (the latter also with GTPase activity), and phospholipase C (Brinkmann, 2007). S1P receptor signaling triggers mechanisms important in particular for the development of the vascular system, heart, smooth muscle cell, as well as the immune system (Brinkmann, 2007). Two isoforms of sphingosine kinases are known to be involved in the synthesis of S1P in mammals. Gene deletions of Sphk1 and Sphk2 were initiated to investigate their potential physiological role in vivo.

1.2.1 Sphingosine Kinase Sphk1 The sphingosine kinase 1 gene (Sphk1) contains six exons from which exon 1 and parts of exon 2 and 6 display untranslated regions. Gene disruption was achieved by insertion of a neomycin cassette into the targeting construct replacing exons 3–5 and a part of exon 6. Deletion of the mRNA encoding Sphk1 was shown by northern blot analysis and real-time PCR. Although a substantial reduction of enzyme activity in several organs of Sphk1/ mice occurred, biosynthesis of sphingosine-1-phosphate was almost not affected by the gene deletion. Serum concentrations nevertheless decreased. S1P is known to regulate lymphocyte trafficking; however, in Sphk1/ mice, lymphocyte distribution in lymphoid organs remained unaffected, and physiological abnormalities could not be observed (Allende et al., 2004). Obviously, Sphk1 isoenzyme sphingosine kinase-Sphk2 could substitute for the lack of Sphk1. To provide further proof, an experiment was initiated in which Sphk1/ mice received an intravenous injection of a synthetic mimic of sphingosine, FTY720. Sphingosine-1-phosphate receptor (S1P1) is required for the exit of lymphocytes from lymphoid organs. Phosphorylated FTY720 is known to trigger inactivation of S1P1. As consequence of the injection of FTY720 in both wild-type and mutant mice, FTY720 was phosphorylated and lymphopenia occurred. These results suggested that Sphk1 is not exclusively required for the activation of the sphingosine analogue FTY720. Second, the amount of sphingosine phosphorylation in Sphk1/ mice by Sphk2 is sufficient to prevent vascular damage or changes in lymphocyte distribution in these mice. Sphingosine kinases exert obviously redundant functions. To prove this, an additional deletion of the gene encoding the isoenzyme Sphk2 was performed.

1.2.2 Sphingosine Kinase Sphk2 and Double-Deficient Mice for Sphk1/Sphk2 The Sphk2 gene consists of seven exons with the coding region located in exon 3 to 7. Deletion of the gene was achieved by replacing parts of exon 4 and exons 5–7 with a LacZ-neomycin cassette. Gene deletion was confirmed with RT-PCR analysis of Sphk1 mRNA from several organs of Sphk2/ mice (Mizugishi et al., 2005). Mutant mice had a normal life span. Histology of several organs was normal.

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Since compensatory mechanisms also obviously play an important role in Sphk2/ mice, mice with double mutation for Sphk1 and Sphk2 were generated. Not unexpectedly, Sphk1//Sphk2/ embryos were not viable. No Sphk1//Sphk2/ embryos survived beyond E13.5 (Mizugishi et al., 2005). Embryos developed until stage E9.5 normally but exhibited vascular defects indicated by cranial hemorrhage at E11.5 to E12.5. Sphk1//Sphk2/ embryos displayed severely disturbed neurogenesis as seen by neural tube closure defects. Moreover, dramatically increased apoptosis accompanied by decreased mitoses was observed in the embryonic nervous system. In the same report, it could for the first time be convincingly demonstrated that deletion of the sphingosine-1-phosphate receptor (S1P1) also induces neurologic symptoms indicated by increased apoptosis in the telencephalon and diencephalons in the developing nervous system. In addition, the finding that S1P1 colocalizes in the brain with sphingosine kinases indicated their close relationship. However, the milder neurologic symptoms observed in the receptor-S1P1 deletion model might be explained by ability of S1P to signal also through other sphingosine-1-phosphate receptors like S1P3 or S1P5. There might be not only a redundancy for sphingosine kinases but also within the sphingosine-1-phosphate receptors S1P1 to S1P5.

1.3 Serine Palmitoyl-CoA Transferase Subunit Gene Deficiency Serine palmitoyl-CoA transferase (SPT) catalyzes the transfer from L-serine to CoA-activated palmitic acid (> Figure 28‐11) and represents, therefore, the key enzyme essential for biosynthesis of sphingosine and its further derivatives sphingosine-1-phosphate, SM, ceramide, and GSLs. Two candidate genes, LCB1 and LCB2, were discovered, which led to lack of SPT activity in yeast suggesting that these two genes encode subunits of SPT. In search of respective homologue genes in mammals it could be demonstrated that SPT is a heterodimer consisting of two subunits designated Sptlc1 and Sptlc2. In order to disrupt SPT activity in mouse models completely, gene deletions of both subunits were performed (Hojjati et al., 2005). The Sptlc1 gene was deleted by elimination of exons 7/8 and the Sptlc2 gene was disrupted by deletion of exon 1. Deletion of the genes Sptlc1 and Sptlc2 were confirmed by northern, southern, and western blot analysis. Both homozygous offsprings of Sptlc1 or Sptlc2 were embryonically lethal, indicating the essential property of SPT during development. SPT enzyme activity was approximately halved in the liver of heterozygous mice. However, they appeared to be healthy and were used for lipid analysis. In this preliminary study, a decrease of liver and plasma ceramide and sphingosine levels was demonstrated. Sphingosine-1-phosphate and in particular lysosphingomyelin content in serum could be shown to be decreased. Cell-specific and, if necessary, inducible mouse models of Sptlc1 or Sptlc2 and combinations of them shall provide the opportunity for further, more detailed studies of the function of SPT for the development of the vascular system, visceral organs, and the nervous system. . Figure 28‐11 Biosynthetic pathways of sphingosine – Block of the synthesis of dehydrosphinganine and subsequent inhibition of sphingosine synthesis in Sptlc/ mice is shown

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1.4 3-Phosphoglycerate Dehydrogenase Deficiency D-3-phosphoglycerate dehydrogenase (Phgdh, EC 1.1.1.95) is reported to initiate the oxidative reaction of 3-phosphoglycerate to 3-phosphohydroxypyruvate, which then, followed by transamination and dephosphorylation, results in L-serine (> Figure 28‐12). L-serine is indispensable for the synthesis of many molecules including nucleotides, amino acids including glycine and cysteine, phospholipids such as phosphatidyl serine and phosphatidyl ethanolamine, as well as sphingolipids. A mouse model was generated in which exons 4 and 5 of the 12 exon-containing Phgdh gene locus was flanked by lox P-sites (Yoshida et al., 2004). Subsequently, heterozygous mice were bred with Cre-deleter mice to introduce a null deletion into the gene. A further breeding step of heterozygous mice revealed the absence of homozygous litters within newborn offspring, thus indicating the vital role of Phgdh. Biochemical investigation of embryos at embryonic day E13.5 were evident for the Phgdh gene deletion by absence of enzyme and reduced phosphatidyl serine/phosphatidyl ethanolamine, ganglioside, and amino acid synthesis in embryonic whole head tissues.

. Figure 28‐12 Biosynthetic pathways of L-serine – Block of 3-phosphoglycerate dehydrogenase-mediated L-serine synthesis in Phgdh/ mice is shown

By histology Phgdh/ embryos demonstrated a retarded development — embryos were markedly smaller as their respective controls. Striking abnormalities were observed in the CNS at E13.5. Two types of brain phenotypes were described at this developmental stage. Type I embryos exhibited mild exencephalic phenotypes and failure of dorsal closure. Type II embryos displayed enlarged lateral ventricles, as well as hypoplasia of the telencephalon, diencephalon, and mesencephalon. As compared with control embryos, both types lacked clear structures of olfactory bulb, ganglionic eminence, and cerebellum development. The study emphasized that alternative pathways for the synthesis of L-serine obviously cannot substitute for the Pghdh-dependent phosphorylation pathway of L-serine synthesis. Therefore, a proper functioning Phgdh gene is essential for normal development and brain morphogenesis at least during embryogenesis. This mouse model bearing a floxed phosphoglycerate dehydrogenase gene enables further, more defined cell-specific deletions of Phgdh. Inducible cre/loxP-systems that may provide even more distinct insights into its function in adult mice might be useful to study.

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Mouse Models with Deletions of Sphingolipid Degrading Enzymes, Introduction

GSLs are degraded in the lysosomes by soluble exo-glycosidases. Consequently, downstream-acting hydrolases fail to degrade the primary substrates of upstream-acting glycosidases. Malfunction at any step therefore leads to the accumulation of the primary sphingolipid substrate and its insolubility to the formation of multivesicular bodies in lysosomes (> Figure 28‐13).

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. Figure 28‐13 Lysosomal degradative pathways of brain sphingolipids – The scheme aws modified according to (Kolter and Sandhoff, 2005). Glycosphingolipids are abbreviated according to the recommendations of the International Union of Pure and Applied Chemistry-International Union of Biochemistry Joint Commission on Biochemical Nomenclature of Glycolipids (IUPAC-IUB Joint Commission on Biochemical Nomenclature of Glycolipids (1999) J. Mol. Biol. 286, 963–970)

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Several neurodegenerative and visceral diseases are characterized by extensive storage of sphingolipids in lysosomal compartments in the cell caused by a defect or mutation in genes encoding sphingolipid digesting enzymes or proteins supporting sphingolipid degradation. Since only a few options existed for therapy of these inherited human lysosomal disorders with low success rate, mouse models were generated providing both, on the one hand insights into disease pathogenesis and on the other hand helpful tools to test treatment strategies.

2.1 Deficiency of b-Galactosidase 1 The human GLB1 gene encodes for the acidic b-galactosidase 1. In lysosomes, terminal b-linked galactosyl residues are cleaved from glycoconjugates by b-galactosidase 1. Mutations of the GLB1 gene accompanied with malfunction of b-galactosidase 1 cause GM1 gangliosidosis and Morquio B disease. GM1 gangliosidosis is primarily a neurological disorder with progressive brain dysfunction. Depending on the residual activity of the b-galactosidase 1, patients develop an infantile, juvenile, or late onset type gangliosidosis. Large amounts of GM1 and only minor amounts of GA1 accumulate in brains of patients. In addition to the neurological disorders, the infantile gangliosidosis can also be characterized by mucopolysaccharidosis-like features of visceral organs. Oligosaccharides derived from keratan sulfate and glycopeptides are enriched in these organs and abnormally excreted into the urine. Morquio B disease is characterized by severe bone deformities but almost no nervous system symptoms. Patients store keratan sulfate in visceral organs and excrete this compound excessively in the urine (Suzuki et al., 2001). The lack of extensive neuronal GM1 storage may be explained by imbalanced substrate specificity of mutant b-galactosidase in patients with Morquio B disease (Okumiya et al., 2003). Like in humans, the mouse Glb1 gene comprises 16 exons with the coding region from exon 1 to 16. Mouse models with b-galactosidase 1 deficiency have been established independently by two groups (Hahn et al., 1997; Matsuda et al., 1997b). Hahn and coworkers (1997) introduced a neomycin resistance gene into the middle of exon 6 resulting in null alleles. Mice were born with a Mendelian inheritance ratio. Residual b-galactosidase activity, as measured with 4-methylumbelliferyl b-galactoside, was 1% in spleen, 4% in brain, and 8% in kidney, probably due to normal galactosylceramidase activity. Using the natural substrate GM1, no activity was measured in Glb1/ mice. Starting from 5 months of age, mutant mice developed tremor, ataxia, and abnormal gait. With 3 weeks of age, storage material was conspicuous in the brain and with 5 weeks extensive storage was observed in neurons throughout the brain and spinal cord. GM1 and GA1 accumulated conspicuously and to similar extent in brain. Further investigations of Glb1/ mice revealed a progressive increase in microglia activation, infiltration of inflammatory cells, and altered blood– brain barrier permeability (Jeyakumar et al., 2003). Matsuda et al. (1997a, b) inserted a neomycin resistance gene into the Sal I site of exon 6. Mutant mice were left with 0–10% and heterozygote mice with 50% b-Galactosidase activity in tails, as measured with 4-methylumbelliferyl b-galactoside. Locomotor activities gradually decreased in Glb1/ mice after 4 months of age. They developed progressive spastic diplegia and died of emaciation at 7–10 months of age. Vacuolated neurons appeared in the spinal cord as early as 3 days after birth and vacuolation extended to neurons in the brainstem, cerebral cortex, hippocampus, and thalamus. Cerebral neurons accumulated extensively GM1 and GA1. In liver, GM1 storage was found mainly in hepatocytes, whereas GA1 accumulated in foamy Kupffer cells (Itoh et al., 2001b). In addition to GA1 and GM1, a minor but significant increase of ceramide dihexoside (probably Lactosylceramide) was observed in liver. Glb1/ mice secreted abnormal oligosaccharides in the urine. Only trace levels of keratan sulfate were detectable in the liver and undetectable in the urine of both mutant and normal mice (Matsuda et al., 1997b). Both mouse models showed no abnormalities in the skeletal system, which may be because mice do not synthesize skeletal keratan sulfate (Venn and Mason, 1985). The remarkable accumulation of GA1 in both mouse models differs from human pathology. This is due to different substrate specificities of human and mouse sialidase. Mouse sialidase, in contrast to human sialidase, converts not only GM3 but also GM1 and GM2 to a significant extent into the asialo-form (Riboni et al., 1995; Sango et al., 1995). As both GM1 and GA1 depend on b-galactosidase 1 degradation, GA1, in contrast to GA2, opens no

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additional degradative pathway and accumulates. Hence, Glb1/ mice are a useful mouse model for the neurological aspects of GM1 gangliosidosis.

2.2 Deficiencies of Isoforms of b-Hexosaminidase, Introduction Two genes, HEXA and HEXB, encode for the subunits a and b of the dimeric b-hexosaminidase leading to three isoforms: HexA (a/b), HexB (b/b), and HexS (a/a). Terminal b-linked N-acetylglucosaminyl and N-acetylgalactosaminyl residues are cleaved from glycoconjugates by b-hexosaminidases. Both subunits have enzyme activity but with different substrate affinities. The a-subunit prefers acidic and the b-subunit neutral substrates. Therefore, HexA (a/b) degrades acidic and neutral substrates, HexB neutral substrates, and HexS mainly acidic substrates (Kolter and Sandhoff, 1999). Nevertheless, at least in mice, complex neutral GSLs are to some extent also substrates for HexS (Sandhoff et al., 2002).

2.2.1 HexA Deficiency Mutations of the HEXA gene accompanied with malfunction of b-hexosaminidase isoforms HexA (a/b) and HexS (a/a) cause B variant (as HexB (b/b) activity is normal) of GM2 gangliosides. Depending on the residual activity of the a-subunit, patients develop an infantile (Tay–Sachs disease), juvenile, or late onset type gangliosidosis. All forms are characterized by neurological symptoms. Tay–Sachs patients develop motor weakness. Sudden noise causes abnormal startle reflexes. Macrocephaly and neurological disorders worsen until death occurs within the next few years. GM2 is the main neuronal storage compound (Gravel et al., 2001). In addition, another anionic derivative of GA2, SM2 (GA2 II3-sulfate), where the sialic acid of GM2 is replaced by a sulfate group, was found to accumulate in addition to GM2 in the liver of a Tay–Sachs patient, by revealing the presence of complex sulfatides, which are below detectional limits in normal human liver (Sandhoff et al., 2002). Like the human HEXA, the mouse Hexa gene comprises 14 exons with exons 1–14 encoding for the a-subunit of b-hexosaminidases (Yamanaka et al., 1994b). Mouse models of HexA deficiency were developed by three groups (Yamanaka et al., 1994a; Cohen-Tannoudji et al., 1995; Phaneuf et al., 1996), and Hexa/ mutant mice were born with expected Mendelian ratio in all three strains. Yamanaka et al. inserted a neomycin resistance gene into exon 8, resulting in a null allele. Less than 1% of normal HexA but 100% of normal HexB activity was found in Hexa/ mice. Mutant mice accumulated GM2 as membranous cytoplasmic bodies in neurons of the CNS in an age-dependent manner. Storage was mild compared with Hexb/ mice (see HexB deficiency) and no behavioral or motor abnormalities were detected in mice even older than 1 year. Neuronal lipid storage was accentuated in cerebral cortex, amygdala, piriform cortex, hippocampus, and hypothalamus, but not in the Purkinje and granular cells in the cerebellar folia and in neurons in the spinal cord. Large pyramidal neurons showed more storage than other cortical neurons in the cerebral cortex (Yamanaka et al., 1994a; Sango et al., 1995; Taniike et al., 1995). Later, it was shown that these mice accumulate large amounts of renal SM2 (GA2 II3-sulfate): more than 100-fold of normal levels at 19–20 weeks of age (Sandhoff et al., 2002). Cohen-Tannoudji and coworkers (1995) disrupted the Hexa gene by insertion of a neomycin resistance gene into exon 8 leading to complete enzyme deficiency of HexA and neuronal lysosomal storage. Unlike the mutant mice of Yamanaka et al. (1994a), Hexa/ mice generated by Cohen-Tannoudji and coworkers did not show hippocampal storage at 150 days. Phaneuf at al. (1996) interrupted exon 11 of the Hexa gene with a neomycin resistance gene. HexA activity was less than 5% in brain and less than 1% in liver extracts. HexB activity was normal. Findings of neuronal storage and lack of symptoms were identical to those found by Yamanaka and coworkers. Differences in the degree of GM2 storage in Hexa/ mice and in human Tay–Sachs patients are due to different catabolic pathways in mice and humans. Human lysosomal sialidase acts entirely on GM3 and not GM2. A deficiency of HexA thus results in strong GM2 storage. However, murine sialidase readily cleaves GM2 to GA2. Therefore, two catabolic pathways for GM2 exist in mice: (1) GM2 degradation by

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HexA to GM3 and (2) GM2 degradation by sialidase to GA2 (> Figure 28‐13). In Hexa/ mice, the latter pathway is not blocked and GM2 is degraded by sialidase to GA2 and further to lactosylceramide by HexB. However, this pathway appears to be overloaded in certain neuronal cell types of Hexa/ mice resulting in a region-specific mild and age-dependent storage of GM2 (Sango et al., 1995). In contrast to sialidase, murine arylsulfatase A appears to act mainly on SM3 (lactosylceramide II3-sulfate) but not very well on SM2, as the renal SM2-storage amount is similar in Hexa/ and Hexa//Hexb/ mice. (Sandhoff et al., 2002).

2.2.2 HexB Deficiency Mutations of the HEXB gene going hand in hand with malfunction of HexA (a/b) and HexB (b/b) cause 0 variant (as with the loss of HexA and HexB almost all Hex activity is lost) of GM2 gangliosides, also called Sandhoff ’s disease. In addition to GM2, which is stored in Tay–Sachs disease, neutral glycolipids (mainly neuronal GA2 and visceral globoside) accumulate, and increased amounts of oligosaccharides with terminal N-acetylgalactosaminyl residues are secreted with the urine. The clinical and pathological picture of Sandhoff ’s disease corresponds essentially to Tay–Sachs disease, although hypertrophy of internal organs and bone deformation also occur (Kolter and Sandhoff, 1999; Gravel et al., 2001). Like the human HEXB, the mouse Hexb gene comprises 14 exons, with exons 1–14 encoding for the b-subunit of b-hexosaminidases (Yamanaka et al., 1994b). Mouse models of Hexb deficiency were developed by two groups (Sango et al., 1995; Phaneuf et al., 1996) and Hexb/ mutant mice were born with expected Mendelian ratio. Sango et al. inserted a neomycin resistance gene into exon 13, resulting in a null allele. Only 2% residual b-hexosaminidase activity was found in liver extracts of mutant mice, most likely corresponding to intact Sandhoff isoenzyme, HexS (a/a). Normal at birth, mice started around 12 weeks with progressive worsening of motor coordination and balance. Severe motor dysfunction became apparent at about 3 months of age, including gait abnormalities with spastic movements starting from hindlimbs and progressing to forelimbs. At about 5 months of age, the mutant mice were unable to move and were sacrificed. Further investigations of Hexb/ mice revealed a progressive increase in microglia activation, infiltration of inflammatory cells, and altered blood–brain barrier permeability (Jeyakumar et al., 2003). Both, GM2 and GA2 accumulated in brain, and GM2 levels were fourfold as compared with Hexa/ mice at 41 days (Sango et al., 1995). Neuronal storage was present throughout the cerebrum, cerebellum, brainstem, spinal cord, trigeminal and dorsal root ganglia, retina, and myenteric plexus. Notably abundant storage was found in the Purkinje and granular cell neurons in contrast to Hexa/ mice. In addition to neurons, storage was noted in perivascular macrophage/microglia and included in addition very likely glycosaminoglycans. Cells of the hepatic sinusoids and epithelial cells of the renal proximal tubules stored similar compounds as brain microglia. Therefore, Hexb/ mice more closely resemble the clinical and pathological course of Sandhoff disease than Hexa/ mice do for Tay–Sachs disease (Sango et al., 1995). Subsequently, analysis of renal glycolipids confirmed remarkable storage of globoside and some GA2 accumulation. SM2 accumulation was only one tenth of that obtained in Hexa/ kidneys at similar age (Sandhoff et al., 2002). Therefore, HexS appears to play an important role in vivo in the degradation of renal SM2, which was confirmed later on in vitro (Hepbildikler et al., 2002). Paneuf and coworkers (1996) inserted a neomycin resistance gene into exon 2, thereby disrupting the Hexb gene. Clinical and pathologic phenotypes were similar to those generated by Sango and coworkers.

2.2.3 HexA and HexB Deficiency Simultaneous homoallelic mutations of the HEXA and HEXB genes leading to total deficiencies of all three isoforms of b-hexosaminidase are not known in humans at this time. By crossing Hexa/ (Yamanaka et al., 1994a) into Hexb/ mice (Sango et al., 1995), Sango and coworkers (1996) obtained ‘‘double knockout’’ (DKO or Hexa//Hexb/) mice with a frequency expected from Mendelian principles, but DKO mice died 1–4 months after birth. DKO mice lacked all forms of b-hexosaminidase, including HexS. At 4–5 weeks of age, Hexa//Hexb/ mice could be distinguished

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from age-matched Hexa/, Hexb/, and control mice by their smaller size and physical dysmorphia. Heads were shorter and snouts broader; feet were thick and broad with flexion contracture of the digits. Corneal opacities, occasional seizure-like activities, and unresponsiveness to sharp noise suggesting deafness were observed. Kyphosis, an abnormally shaped rib cage, and short and thickened long bones were evident by radiographic study. Glycolipid storage of GM2 and GA2 in brain was similar to that of Hexb/ mice, which are still capable to produce HexS (a/a), indicating that HexS does not play a significant role in neuronal ganglioside degradation. Cerebroside and sulfatide levels were reduced, probably due to hypomyelination of the white matter, whereas at the terminal stage of disease, many axonal spheroids and scattered myelin figures suggestive of myelin degeneration were recognized. Storage was also obvious in trigeminal and dorsal root ganglia, neurons in the myenteric plexus, and visceral autonomic ganglia of the PNS. Although enzymes involved in classical glycosaminoglycan catabolism were found to be elevated, glycosaminoglycan fragments were 50-fold increased in the DKO urine as compared with Hexa/, Hexb/, and control mice (Sango et al., 1996). This implies a significant role of the minor Sandhoff isozyme, HexS, for the in vivo degradation of glycosaminoglycans, which was subsequently also verified in vitro (Hepbildikler et al., 2002). Further investigation of renal glycolipids revealed remarkable SM2 and mild GA2 storage, comparable with that of Hexa/ or Hexb/ mice. Globoside accumulated also, but not completely to the degree found in Hexb/ mice. This could be explained by the additional and remarkable storage of a catabolic globoside precursor, IV6-GlcNAcb-Gb4Cer, coming from the degradation of renal Gal-b1,4 (Fuc-a1,3-)-GlcNAcb1,6(Gal-b1,3-)-GalNAcb1,3-Gb3Cer, in DKO kidneys, a compound not detectable in Hexa/, Hexb/, or control kidneys. Hence, HexS contributes in vivo also to the degradation of complex neutral glycolipids with terminal b1,6-linked N-acetylhexosaminyl residues (Sandhoff et al., 2002). Biochemically, pathologically, and clinically, DKO mice showed features of severe mucopolysaccharidosis in various organs (Sango et al., 1996; Suzuki and Proia, 1998).

2.3 Lysosomal Activator Proteins, Introduction Lysosomal degradation of sphingolipids is performed by soluble hydrolases, whereas sphingolipid biosynthesis is performed by membrane proteins. Degradation of sphingolipids with short carbohydrate chains of four or fewer sugars in vitro is in the presence of lysosomal hydrolases solely and without additional factors a very slow process. In vivo several additional factors accelerate this process remarkably: sphingolipids are digested (1) from intra-endosomal/lysosomal membrane structures with a strong positive curvature, a low cholesterol, and a high bis(monoacylglycero)phosphate (BMP) content and (2) with the help of small nonenzymatic glycoproteins with lipid transfer activity, so-called sphingolipid activator proteins (SAPs). As they have a membrane-perturbing and lipid-binding property, they facilitate lipid access toward lysosomal-soluble hydrolases. Until now, five SAPs with different affinities for certain sphingolipids are known: SAP-A, -B (known before as sulfatide activator), -C (known before as glucosylceramide activator), -D, and GM2-activator protein (Kolter and Sandhoff, 2005; Locatelli-Hoops et al., 2006).

2.3.1 GM2-Activator Protein Deficiency Deficiency of the GM2-activator protein (GM2-AP) causes AB variant of GM2 gangliosidosis. The clinical course of the AB variant resembles mainly that of Tay–Sachs disease but with a slightly retarded onset. Loss of GM2-AP causes strong accumulation of GM2 and GA2 in brains of patients (Sandhoff, 1977; Sandhoff et al., 2001). Like the human GM2A gene (Klima et al., 1991), the mouse Gm2a gene has four exons with exons 1–4 encoding for GM2-AP. Deletion of GM2-AP was obtained by replacement of the entire exon 3 and part of exon 4 covering the coding region by the neomycin resistance gene. Gm2a/ mice were obtained in a Mendelian fashion, grew normally, and were fertile. Mutant mice showed subtle neurological dysfunction: overall rotorod performance was significantly impaired and passive-avoidance task suggested a possible memory deficit in the mutants. In the brain, GM2 accumulated to a similar extent in Gm2a/ as in

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HexA/ mice. In addition, a slight accumulation of GA2 was found in Gm2a/ brain that was not found in HexA/ brain (Liu et al., 1997). A further study revealed storage of small amounts of SM2 (GA2 II3sulfate) in Gm2a/ kidneys, a compound not detectable in control kidneys (Sandhoff et al., 2002). Hence GM2AP contributes in vivo not only to the degradation of GM2 and GA2, but also to that of SM2, which was later on confirmed in vitro (Hepbildikler et al., 2002; Hepbildikler et al., 2003). Neural storage was restricted to piriform, entorhinal cortex, amygdala, and hypothalamic nuclei similar to HexA/ mice (Liu et al., 1997).

2.3.2 Prosaposin Deficiency The four activator proteins SAP-A, -B, -C, and -D are encoded by a single gene (PSAP). Translation of this gene results in a single precursor protein, called prosaposin (pSAP), from which the four saposins/ sphingolipid activator proteins (SAP-A to -D) are produced by proteolytic processing (Furst et al., 1988; O’Brien et al., 1988; Nakano et al., 1989; Holtschmidt et al., 1991). Prosaposin deficiency is a very rare and fatal infantile storage disorder. Cases so far reported were either due to a homoallelic mutation in the initiation codon of the PSAP gene or due to a homoallelic 1 bp deletion within the SAP-B domain of the PSAP gene, which leads to a frame shift, a premature stop codon, and rapid degradation of mutant mRNA. The deficiency of pSAP causes hepatosplenomegaly and a neurological disease. Storage in form of intralysosomal oligolamellar vesicles includes ceramide, monoand dihexosylsceramides, sulfatide (SM4s), as well as Gb3Cer and Gb4Cer or GM3 and GM2, depending on the tissue investigated. No phospholipids including SM or cholesterol were increased (Bradova et al., 1993; Hulkova et al., 2001; Sandhoff et al., 2001). Supplementing the cell culture medium with the human protein pSAP nearly completely reversed the aberrant accumulation of multivesicular structures in cultured fibroblasts from human patients (Burkhardt et al., 1997). Like the human gene, mouse gene (Psap) of prosaposin consists of 15 exons. Exons 3–5, 6–9, 10–11, and 12–14 encode for the proteins SAP-A, -B, -C, and -D, respectively. In a mouse model, the Psap gene was disrupted by insertion of a neomycin-resistant cassette within exon 3. Generation of Psap/ mice from chimeric and heterozygote mice resulted in total loss of Psap mRNA. With less than 15% of all offspring, Psap/ mice were not generated in a Mendelian inheritance ratio, indicating death of Psap/ mice in utero or the perinatal period. Mutant mice surviving the perinatal period died around 35 days after birth. They developed neurological symptoms after 18–20 days and were by then slightly smaller than their littermates. Tremulousness of the head and mild weakness/ataxia of the hind legs were initial symptoms. Within the next 10 days gross shaking of the head and the trunk and severe weakness of all legs appeared. About 5 days before death intermittent seizures developed, which progressed to continual tonic status epilepticus. The pathology of Psap/ mice was that of combined neurovisceral storage and leukodystrophy. No organomegaly was observed. Neuronal storage, numbers of storage neurons as well as axonal spheroids increased with age, accompanied with hypomyelination, astrogliosis, and increase of macrophages. Lactosylceramide was most prominently increased in all tissues reported (brain, liver, and kidney). Further lipids accumulating in at least one of these organs were ceramide, glucosyl- and galactosylceramide, sulfatide, globotri- and tetraosylceramide as well as GM3, GM2, and GM1. Catabolism of these lipids was abnormally slow in cultured mutant fibroblasts and resembled closely that of cultured fibroblasts from the human patients. Among the lysosomal hydrolases necessary for the degradation of the accumulating GSLs, only galactosylceramidase and glucosylceramidase showed significant lower activities in mutant liver extracts. Therefore, the Psap/ mice mimic biochemical changes of human patients with pSAP deficiency, as well as the severe clinical phenotype (Fujita et al., 1996; Oya et al., 1998).

2.3.3 SAP-A Deficiency Human diseases caused by a specific defect or deficiency of the SAP-A protein are not known.

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A mouse model of SAP-A deficiency was established by introducing an amino acid substitution (C106F) into the SAP-A domain of Psap by the Cre/loxP system, which eliminated one of the three disulfide bonds (Matsuda et al., 2001). The maintenance of these three disulfide bonds is considered essential for the functional properties of all four SAPs (Vaccaro et al., 1995). PsapC106F mRNA was of normal length and stability, in contrast to its absence in Psap/ mice. PsapC106F/C106F mice were obtained with Mendelian frequency and survived up to 5 months. In about 2.5 months they developed slowly progressive hind leg paralysis. At the terminal stage, tremors and shaking were also obvious. The pathology and analytical biochemistry described a mild form of infantile globoid cell leukodystrophy (GLD) in man (Krabbe disease) with increased levels of GalCer especially in the kidney and 1-alkyl, 2-acyl, and galactosylglycerol (GalEAG) in the testis. Brain psychosine (galactosylsphingosine) levels were approximately twice the normal levels. Interestingly, similar to Psap/ mice, galactosylceramidase activity of PsapC106F/C106F mice was half of that of wild-type mice, suggesting a stabilizing function of SAP-A for galactosylceramidase (Matsuda et al., 2001).

2.3.4 SAP-B Deficiency Human diseases caused by a specific defect or deficiency of the SAP-B protein are known as variant forms of metachromatic leukodystrophy with juvenile or late infantile onset (Stevens et al., 1981; Kretz et al., 1990) (for review see also Sandhoff et al., 2001). Biochemical analyses of the patient’s urine reveals increased levels of sulfatide (SM4s), lactosylceramide, globotriaosylceramide, and digalactosylceramide (Li et al., 1985). A mouse model of SAP-B deficiency manifested slowly progressive sulfatide accumulation (Sun and Grabowski, unpublished data).

2.3.5 SAP-C Deficiency Deficiency of the SAP-C protein caused a juvenile variant of Gaucher disease (GD) (Christomanou et al., 1986) and (Sandhoff et al., 2001) and strong accumulation of glucosylceramide was demonstrated from the liver of those patients (Christomanou et al., 1989). In a Mouse model of SAP-C deficiency slowly progressive glucosylceramide accumulation has been observed (Sun and Grabowski, unpublished data).

2.3.6 SAP-D Deficiency Human diseases caused by a specific defect or deficiency of the SAP-D protein are not known. A mouse model of SAP-D deficiency was established by introducing an amino acid substitution (C509S) into the SAP-D domain of Psap, eliminating an essential disulfide bond. The mutation resulted in stable mRNA of expected length. PsapC509S/C509S mice were obtained with Mendelian frequency. Maximum life span was estimated to be 1.4 years. Within 2 months, mice developed progressive polyuria because of renal tubular degeneration. Ataxia was evident at the age of 4 months. Cerebellar Purkinje cells selectively disappeared progressively and almost completely by the age of 12 months. Ceramides, particularly those containing hydroxy fatty acid moieties, most prominently increased in the cerebellum, suggesting cytotoxicity of ceramide (Matsuda et al., 2004).

2.3.7 Combined SAP-C and SAP-D Deficiency Human diseases caused by the combined deficiency of SAP-C and -D proteins are not known. A mouse model of combined SAP-C and -D deficiency was established by simultaneous substitution of the fifth cysteine of the SAP-C (C384P) and SAP-D (C509S) domains in Psap to prolin and serine,

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respectively1. PsapC384P&C509S/C384P&C509S-offspring were obtained with Mendelian ratio from the heterozygote crosses and survived not more than 58 days ( 8 weeks) after birth. In 6-week-old animals, partial loss of Purkinje cells was evident and storage bodies were present in neurons of the spinal cord, brain, and dorsal root ganglia. Especially prominent was the accumulation of glucosylceramide and hydroxy ceramides in brain and kidney. Lactosylceramide, which was the main storage compound in Psap/ mice, increased only to a minor extent. Mutant prosaposin increased in the endoplasmic reticulum, failed to be secreted but reached the endosomal/lysosomal compartment. Slowed processing of the mutant precursor protein, pSAPC384P&C509S, resulted in decreased levels of SAP-A and -B. In addition, b-glucosidase activity and protein decreased because of the lack of SAP-C, as compared with SAP-D-deficient (PsapC509S/C509S) mice (Sun et al., 2007).

2.4 a-Galactosidase A Deficiency A deficiency in the lysosomal degrading enzyme a-galactosidase A (EC 3.2.1.22) causes Fabry disease, a human X-linked inherited disorder that is accompanied by accumulation of the neutral GSL globotriaosylceramide (Gb3Cer) (Desnick et al., 1995). Gb3Cer expression is predominantly restricted to organs derived from mesodermal origin such as heart, spleen, kidney, and vascular endothelium. One major part of the burden of the disease results from vascular involvement associated with progressive damage of the heart, kidney, as well as cerebrovascular strokes. Additional disease manifestations include paresthesias in extremities, corneal dystrophy, and angiokeratomata. A mouse model of Fabry disease was created in 1997 by insertion of a neomycin selection cassette into a part of exon/intron 3 of the enzyme (Ohshima et al., 1997). Gene deletion was confirmed by southern blot analysis and absence of enzyme activity. The mouse model showed similarities to the human disorder indicated by concentric lamellar inclusions in the lysosomes of kidney tubular cells, as well as in cultured fibroblasts detected by electron microscopy. The glycolipid Gb3Cer content of liver and kidney was increased. Peripheral nerve involvement, neuropathic pain, and chronic acroparesthesiae have been reported to be the most frequently observed neurological signs in the human disease (Clavelou and Besson, 2007), obviously caused by an affected vascular system. However, similar distinctive features, so far, were not described in the mouse model.

2.5 Galactosylceramidase Deficiency Low or absent galactocerebrosidase activity causes GLD, also known as Krabbe disease, an autosomal recessively inherited disorder (Wenger et al., 2001). The disease with an incidence of 1 case in 100,000 births is characterized by the presence of globoid cells in affected areas of the brain, impairment of the protective myelin sheaths, and oligodendrocyte degradation in particular in the white matter of the brain. Symptoms appear before 6 months of age and progress until patients die within the next 1.5 years. With lower frequency, adult patients have also been diagnosed having GLD but with high variability in their clinical course. Phenomena of the disease are weakness, tremor, vision loss, and impairment of the nervous system. In 1976, the twitcher (twi) mouse was discovered in the Animal Resources Division of the Jackson Laboratory in a CE/J/C57Bl6 mouse strain. Homozygous mice were generated by crossbreeding of heterozygous couples after backcross to C57Bl6. Initially it was not known which gene defect might have caused the obvious symptoms, for example, ataxia, appearing in these mice. During maturation, starting from day 20 post partum, twitcher mice displayed a marked retardation in weight gain as compared with control mice. Homozygous twi/twi-mice showed symptoms similar to those observed in human patients suffering

1 The position of the exchanged cystein residue within the prosaposine amino acid sequence refers to the sequence published with the NCBI accession number: NP_035309.2.

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from Krabbe disease. Twitcher mice were investigated immunohistochemically and by electron microscopy to disclose their secret (Suzuki and Taniike, 1995). Electron microscopy of the funiculus cuneatus in the spinal cord did not offer appreciable differences between twi/twi-mice and controls at postnatal day P10. At day P20, the myelin sheaths in the Twitcher mice were slightly thinner than those from wild type. Numerous demyelinated axons were seen in the spinal white matter at P40. In addition, single axons were surrounded by macrophages. In oligodendrocytes in culture, major spreading processes could be seen; however, secondary and tertiary processes were much less developed in twi/twi-mice than in controls. In Schwann cells of peripheral nerves, rare cytoplasmic inclusions were observed but did not appear to affect early myelination (Suzuki and Taniike, 1995). In search for genetic alterations associated with the symptoms observed in twitcher mice, a mutation in the gene encoding galactocerebrosidase (human:GALC; mouse Galc) was detected (Sakai et al., 1996). Investigations of twitcher mice paired with observations in humans revealed that GLD indeed might mainly be restricted to the white matter of the brain (Itoh et al., 2002). Caused by the absence or lowered galactocerebrosidase activity, psychosine (galactosylsphingosine) accumulated in brains of Krabbe disease patients as well as in the twitcher mouse animal model (Whitfield et al., 2001). Excessive storage of psychosine, which may be toxic to cells in high amounts, was proposed to be causative of the extreme demyelinating processes and loss of oligodendrocytes observed in both Krabbe disease and in the animal model for GLD. In two recent publications, indirect effects of psychosine were suggested. Both activation of phospholipase A2 accompanied with an increase of lysophosphatidylcholine (Giri et al., 2006) and loss of peroxisomal protein and function (Haq et al., 2006) were assumed to be triggered by the excess of psychosine and to be secondarily associated with oligodendrocyte cell death in GLD. Since the use of twitcher mice as models for therapy studies on GLD is limited due to their low life span, a new mouse model with a defined point mutation in exon 5 of galactocerebrosidase has been created (Luzi et al., 2001). These mice produced less psychosine as twi/twi-mice. Mutant mice gained weight similar to their wild-type controls. Although they developed symptoms in the CNS and PNS, their disease was less pronounced. Mutant mice lived about 15 days longer as compared with twitcher mice. These advantages may make them more suitable to be used as an animal model in search for therapeutic approaches for treatment of Krabbe disease in humans. Initial studies with bone marrow transplantations of normal donors enhanced the life expectancy of twi/twi-mice from 35 to 100 days (Suzuki, 1995). Furthermore, it could be demonstrated that galactocerebrosidase-lacking oligodendrocytes transplanted into acceptor mice, in principle, are able to internalize externally applied enzyme to maintain stable myelin (Kondo et al., 2005). Single-dose galactocerebrosidase administration directly into the CNS of twi/twi-mice at postnatal day P20 led to an increase in the life span of approximately 10 days as compared with untreated littermates that died after 40–42 days (Lee et al., 2007). These experiments highlight the possible relevance of exogenously applied enzyme as an appropriate tool for the therapy of human GLD patients.

2.6 b-Glucosidase Deficiency 2.6.1 Acid b-Glucosidase, Lysosomal Deficiency Glucocerebrosidase deficiency, also known as GD, is an autosomal recessive GSL storage disorder. Patients lacking the lysosomal hydrolase responsible for degradation of glucosylceramide store massive amounts of this compound in cells of the reticuloendothelial system. In humans, GD is classified into three clinical subtypes (Beutler and Grabowski, 2001). GD-1, a non-neuronopathic form, is characterized by visceral symptoms. However, recent studies in GD patients of this type revealed neurologic alterations at a subclinical level, which were not clinically noticeable (Mercimek-Mahmutoglu et al., 2007). GD-1 was highly prevalent in the Ashkenazi Jewish population. GD-2 is characterized by an acute early onset of a CNS involvement, and GD-3 reflects a variable degree of neurologic involvement in the disease. Symptoms observed in the nervous system range from

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supranuclear gaze palsy, mycoclonic seizures, and spasticity in GD-2-patients to additional slowly progressive dementia and ataxia in GD-3 patients. A subset of patients with severe type 2 GD display an ichthyosislike skin (Sidransky et al., 1996). More than 200 point mutations in the gene locus of GBA1 in humans have been discovered. This fact proved to be the most difficult part for the development of an appropriate mouse model for the study of GD. Nevertheless, in the past years, several models have been generated, which promise to be very useful tools for drug development and better understanding of the disease and its prognosis. The mouse glucocerebrosidase gene (Gba1) contains 11 exons from which exon 1, as well as parts of exon 2 and 11 remain untranslated. The first deletion of glucocerebrosidase was performed in 1992 by Tybulewicz et al. To disrupt the gene, a targeting vector was constructed in which parts of exon 9 and 10 were replaced by a neomycin selection cassette. The gene deletion could be confirmed by the absence of Gba1 enzyme activity and an increased storage of glucosylceramide (GlcCer) in brain, lung, and liver of Gba1/ mice. Homozygous litters were born but appeared severely compromised and gained less weight as controls. Gba1/ mice had abnormal respiration and developed rapidly progressing cyanosis; feeding and movement were decreased. All homozygous mutants died within 24 h after birth. Although correlations to the human GD type 2 clearly could be drawn, symptoms in the homozygous mutants like liposomal lipid storage in macrophages appeared to be milder. Lysosomal storage in liver was restricted to Kupffer cells in the liver sinusoids while hepatocytes remained unaffected (Tybulewicz et al., 1992). Further detailed investigation of Gba1/ mice revealed storage of GlcCer in microglia cells and in neurons of brainstem and spinal cord, but not in neurons of the cerebellum and cerebral cortex (Willemsen et al., 1995). The patchy pattern of neuronal storage in Gba1/ mice replicates the pattern seen in human GD-2 patients, thus demonstrating a pronounced variation of GSL catabolism within different brain regions. Severe alterations in skin of Gba1/ mice have been observed, which match similar findings in infantile human type 2 GD patients. Since it is known that the main GSL in the epidermis consists of GlcCer, investigations whether an increased storage of GlcCer might have any influence on skin permeability and barrier function were initiated (Holleran et al., 1994). In this study, a dramatic elevated GlcCer content in the epidermis and stratum corneum of Gba1/ mice could be demonstrated. Impaired skin permeability, indicated by an extreme transepidermal water loss and alterations in the formation of the essential intercellular bilayer between the stratum granulosum/stratum corneum interstices, was observed. These results suggest that GlcCer degradation by Gba1 plays a vital role for delivery and correct arrangement of free and protein-bound ceramides for the development of the stratum corneum and may be a model for the ichthyosis-like skin observed in some patients with severe type 2 GD. Clinical course and severity of the disease in humans strongly depends on the site of the GBA1 gene mutation. To obtain more detailed insights into clinical GD subtypes, two mouse models mimicking human GD type 2 and 3 were generated. Insertion of point mutations in the Gba1 gene locus in mice was performed using the single insertion mutagenesis procedure (SIMP) (Liu et al., 1998). The models carried either a double mutation (RacNci I) mimicking severe GD-2 or a single mutation (L444P) associated with less severe GD-3 disease (Liu et al., 1998). RacNci mice contained a mutation in the Gba1 gene locus in which leucine-444 and alanine-456 in exon 10 were substituted by proline. In L444P mutant mice solely leucine-444 was substituted by proline. Studies with mice bearing RacNci I/RacNci I mutations revealed only little residual Gba1 enzyme activity and concomitantly accumulation of GlcCer in brain and liver. Mice homozygous in L444P mutation exhibited higher activity of Gba1 and therefore, not surprisingly, no detectable accumulation of GlcCer was seen in brain and liver. However, homozygous offspring derived from both mutant strains died within 48 h of age (Liu et al., 1998) because of an epidermal permeability barrier defect similar to that described in Gba1/ mice (Holleran et al., 1994). An appropriate balance in GlcCer degradation and synthesis (Jennemann et al., 2007) is necessary for an intact barrier function of the skin. In a study published in 2002 by Mizukami and coworkers, homozygous L444P adult mice were generated by crossbreeding of Gba1 mutants bearing the L444P mutation with heterozygous glucosylceramide synthase Ugcg+/ animals (Yamashita et al., 1999). After several breeding steps, viable mice with homozygous L444P mutation were created, which exhibited partial Gba1 enzyme deficiency (Mizukami

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et al., 2002). Multisystem inflammation and B-cell hyperproliferation, an aspect also observed in human GD patients, could be demonstrated. Since in these mice no obvious lysosomal storage of GlcCer occurred, it may be speculated that even a minimal increase of GlcCer, without causing direct symptoms, still may trigger inflammatory events in Gaucher patients. Several more animal models with different point mutations in the b-glucocerebrosidase gene locus have been generated by Xu et al. in 2003. The resulting mouse models mimicked multiple phenotypes of GD, including CNS and visceral, as well as immunological alterations. Combination of these mutant mice with animals displaying a low level of prosaposin C, an activator protein required for optimal in vitro hydrolysis activity of Gba1, resulted in further disease variants in mice (Sun et al., 2005). Recently, a conditional deletion of the glucocerebrosidase gene was reported (Sinclair et al., 2007). Exons 9–11 were flanked with loxP sites. Floxed Gba1 animals were bred with cre-recombinase-expressing mice to generate cell-specific deletions of glucocerebrosidase. For this purpose, mice expressing cre either under control of the Tie2 promoter limited to endothelial and hematopoietic cells or LysMCre mice with cre-activity in the myeloid lineage were used. In both models, storage of GlcCer in organs exhibiting cre-expression could be shown, leading to progressive splenomegaly with Gaucher cell infiltrations and modest storage of GlcCer in the liver. All these various mouse models as described before appear to be obligatory tools to study GD at a certain status in analogy to the human disease. They might prove to be useful in search for improved treatment methods such as enzyme replacement (Butters et al., 2003; Sly, 2004), enzyme inhibition (Priestman et al., 2000), and gene therapeutical approaches using retroviral transfection vectors driving the glucocerebrosidase gene (Enquist et al., 2006; Sun et al., 2006).

2.6.2 b-Glucosidase, Endoplasmic Reticulum Deficiency In contrast to Gba1 that is active in lysosomes, Gba2 is a membrane-bound protein of the ER. Gba2 is highly expressed in brain, liver, testis, and epididymis. Deletion of exons 5–10 of 17 exons of the gene led to a constitutive deletion of Gba2 activity in mice (Yildiz et al., 2006). Deletion of the Gba2 gene did not cause obvious neurologic symptoms, organomegaly, or reduction in life span. Mice exhibited impaired fertility indicated by abnormal sperm heads and acrosomes, paired with defective mobility. The observed symptoms were associated with progressive GlcCer accumulation in Sertoli cells.

2.7 Arylsulfatase A Deficiency Arylsulfatase A (Arsa, EC 3.1.6.1) deficiency, also known as metachromatic leukodystrophy, is a human autosomal recessively inherited lysosomal storage disease. Due to the absence of the degrading enzyme, storage of its substrate sulfatide (cerebroside-3-sulfate, SM4s) occurs in several organs such as kidney, liver, and gall bladder (Sugita et al., 1974; von Figura et al., 2001). In addition, extensive accumulation can also be seen in granules from oligodendrocytes, in astrocytes, and in distinct types of neurons in the nervous system, for example, sensory neurons of dorsal root ganglia. The disease is accompanied with progressive demyelination, ataxia, gait disturbance, and later on by loss of speech, epileptic seizures, and spastic quadriplegia. The symptoms are progressive with age. To gain a more detailed insight into the human disease, in 1996, a mouse model was generated by Hess et al. The mouse Arsa gene contains nine exons from which exon 1 is noncoding. Exons 2 and 9 contain additional noncoding sequences. Gene deficiency was achieved by placing a neomycin selection cassette into coding exon 4. Offsprings were born according to Mendelian inheritance and were fertile. Gene deletion was confirmed by absence of Arsa-mRNA and enzymatic activity. Enzyme activity was measured by the SM4s degradation products: GalCer or ceramide in fibroblast or oligodendrocytes in culture. Phenotypic investigations of Arsa/ mice revealed neuromotor abnormalities indicated by reduced ability to stay on a rotating rod and significantly lowered swimming velocity. Impaired learning and memory as well as hyperactivity was observed. Some of the distinctive features like impaired motor coordination

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and equilibrium, reduced learning, and loss of memory were obvious only in older mice at the age of 1 year (D’Hooge et al., 2001). Sulfatase-deficient mice showed an absence of brainstem auditoryevoked potentials. The decline to acoustic stimulation might be explained by a dramatic loss of spiral cells in Arsa/ mice (D’Hooge et al., 1999). In addition, tremor occurred late at the age of 1 year in Arsa-deficient mice. Histochemical investigations showed pronounced sulfatide storage in granules in the white matter and prismatic inclusions in astrocytes of the brains of Arsa/ mice. Sulfatide storage without obvious cellular damage was apparent in several classes of neurons from brainstem, diencephalons, spinal cord, and cerebellum. Although Purkinje cells did not store sulfatide, surprisingly, their morphology was altered. In addition, progressively increased microglia activation was seen. Astrogliosis was observed in white matter tracts such as corpus callosum or in optical nerves (Hess et al., 1996). A retarded expression of myelin proteins such as myelin basic protein (MBP), PLP, and myelin and lymphocyte protein (MAL) may be indicative for a delay of myelin formation in Arsa/ mice (Yaghootfam et al., 2005). Although characteristics for the disease in humans such as extreme demyelination or absence of metachromatic material in hepatocytes were not found in the murine model, Arsa/ mice demonstrated phenomena reminiscent in particular to an early stage of the human disease. Recently, a mouse model was described exhibiting multiple sulfatase deficiency by deletion of the Sumf1 gene. The phenotype in mice was even stronger as in Arsa-deficient mice, since all known sulfatases were affected (Settembre et al., 2007). Sumf1 is involved in posttranslational modification of sulfatases in prokaryotes and eukaryotes. A mutation in the gene leads to multiple sulfatase silencing and to an autosomal recessive disorder. A mouse model with gene deletion of exon 3 of the Sumf1 gene locus was generated. Drastic reduction of activity of all sulfatases was achieved. Mice had retarded growth, skeletal abnormalities, neurological defects, and early mortality. In all tissues generalized inflammatory events, a strong increase of cytokines and apoptotic markers was observed. Moreover, progressive cell vacuolization and lysosomal storage of glycosaminoglycans was seen. Additional storage of sulfatides was not described but may also occur in these mice. Activation of microglia paired with remarkable astrogliosis was observed in cerebellum and cortex.

2.8 Sphingomyelinase Deficiency, Introduction Two groups of sphingomyelinases, acid and neutral sphingomyelinases (EC 3.1.4.12), are known to cleave the phosphodiester group of SM into ceramide and phosphocholine. SM is ubiquitously distributed in membranes of the endoplasmic reticulum, Golgi apparatus, in lysosomes, and in the plasma membranes, whereas sphingomyelinases differ in their tissue and subcellular distribution. Acidic sphingomyelinase (Smpd1) is expressed in the lysosomes. Gene mutations in humans cause Niemann-Pick disease (NPD), an autosomal recessive neurodegenerative disorder (Schuchman and Desnick, 1995; Vanier and Suzuki, 1996; Kolodny, 2000). Two clinical types of NPD are known, type A and type B, bearing mutations in the SMase gene locus. NPD type A causes severe neurovisceral storage of SM in particular in brain and patients usually die within 3 years of age. Type B is milder; patients show only little or aberrant neurological symptoms and often live until adulthood. The mechanism of such a phenotypic polymorphism of NPD is not yet clear but might be explained by residual Smpd1 activity in NPD type B patients. Neutral sphingomyelinases (Smpd2 and Smpd3) are, in contrast to Smpd1, found in the endoplasmic reticulum and are expressed predominantly in the brain but also in lower amounts in several mammalian tissues. Neutral sphingomyelinases are believed to play critical roles ranging from stress-induced ceramide generation to anti-apoptosis, cell survival, proliferation, and cell senescence. In order to elucidate the functions of these SM digesting enzymes, genetic mouse models were generated.

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2.8.1 Acidic Sphingomyelinase The gene encoding acidic sphingomyelinase (Smpd1) contains six exons with untranslated regions in exons 1 and 6. Smpd1-deficient mice were generated by two groups in parallel either by targeting exon 3 (Otterbach and Stoffel, 1995) or exon 2 (Horinouchi et al., 1995) with a neomycin selection cassette. Gene deletion was confirmed by northern/southern blot. Absence of Smpd1 activity was determined in several organs. In both studies, a pronounced neutral sphingomyelinase activity still remained in all tissues. Lipid analysis revealed a strong accumulation of SM in several organs of Smpd1-deficient mice. Mutant mice generated by Otterbach et al. developed normally until 8–10 weeks. From this point, Smpd1/ mice showed signs of tremor and their gait became progressively ataxic; an initial hint for possible degenerations in the cerebellum. The latter observations were substantiated by light microscopy. A degeneration of the Purkinje cell layer could be detected, obviously responsible for the severe impairment of motor coordination in Smpd1 mutant mice. In addition, ganglion cells in gray matter of Smpd1/ mice were frequently swollen and occasional vacuolization in the cytoplasma occurred. Liver and spleen of Smpd1/ mice were drastically enlarged. Upon histologic examination, Kupffer cells and hepatocytes of the liver were shown to be swollen, displayed a foam cell-like appearance with extended vacuolization. Similar observations were made in spleen and lungs. Life expectancy of Smpd1-deficient mice was about 4 months (Otterbach and Stoffel, 1995). In principle, the mouse model showed phenotypically close similarities to NPD type A in humans. Smpd1/ mice generated by Horinouchi et al. (1995), however, survived longer and reached an age up to 8 months. Although tremor and a dramatic loss of Purkinje cells in cerebellum also occurred, this murine model demonstrated some major differences to the one previously described. In comparison with Smpd1/ mice, generated by Otterbach et al., a diffuse atrophy of cerebellum and midbrain was observed resulting in a reduction of brain weight to less than half of control mice. In addition, hepatosplenomegaly could not be detected in the mice from Horinouchi et al., one of the most striking observations in Smpd1/ mice from Otterbach et al. Blood cholesterol levels were found to be elevated. The reasons for the phenotypic differences in both mouse models are not yet clear.

2.8.2 Neutral Sphingomyelinases Two non-lysosomal neutral sphingomyelinases, Smpd2 and Smpd3, have been discovered and cloned. Smpd2 is ubiquitously expressed throughout all organs. The Smpd2 gene contains ten exons and genetic deletion of the enzyme was initiated by insertion of a neomycin cassette from the end of exon 2 to start of exon 8 (Zumbansen and Stoffel, 2002). Gene deletion was shown on the molecular and protein levels. In addition, a striking reduction of enzyme activity was seen in kidney and heart, whereas liver, lung, thymus, and testes still displayed significant residual activities. High SMase activities were still present in brain and intestine. Smpd2/ mice developed inconspicuously. To get more insight into the functions of the neutral sphingomyelinases, the Smpd3 gene was also deleted. A neomycin cassette was placed into exon 3, the first coding exon of the gene that contains 9 exons in total (Stoffel et al., 2005). Mice lacking Smpd3 enzyme activity were obtained, and Smpd3 was found to be the most prominent neutral sphingomyelinase abundant in neurons of the CNS. Phenotypic analysis of Smpd3/ mice revealed a novel form of dwarfism indicated by growth retardation and deformation of the skeleton paired with delayed puberty and generalized hypoplasia. Hypothalamus-induced pituitary hormone secretion was impaired in Smpd3-deficient mice. Results suggested that the functional role of Smpd3 may be based on maintenance of domain structures of the Golgi secretory pathway perhaps triggered by a close cooperation of SM-digesting and -synthesizing enzymes. Both sphingomyelinase and SM synthase colocalize in the Golgi compartment. Complete loss of neutral sphingomyelinase enzyme activity could be seen in Smpd2/Smpd3 double null mice indicative for the existence of only these two enzymes exerting sphingomyelinase activity and for their ability to substitute partly the deficiency of the other enzyme, respectively. Lipid storage, similar to

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that reported for Smpd1/ mice (NPD type A mice) in brain and visceral organs, was not observed, neither in Smpd3- nor Smpd2/Smpd3 double-deficient mice. Complete rescue of Smpd3/ mice from severe short-limbed skeletal dysplasia was recently shown by introduction of a full-length Smpd3-cDNA (Stoffel et al., 2007).

2.9 Acid and Neutral Ceramidase Deficiency Acid ceramidase or N-acylsphingosine amidohydrolase (Asah1), EC 3.5.1.23, catalyzes the degradation of ceramide to free fatty acid and sphingosine. Acid ceramidase deficiency, also known as Farber Disease (FD) or Farber lipogranulomatosis, constitutes an inherited recessive gene disorder in humans. FD is a rare lysosomal disorder characterized by symptoms such as deformed joints, subcutaneous nodules, progressive hoarseness, lipid accumulation, neurological features, and early death. Based on the clinical severity and sites of involved tissues, FD was categorized into seven subtypes (Park and Schuchman, 2006). Different prognosis of the disease might be explained by the respective site of the acid ceramidase gene point mutation. FD is diagnosed by high levels of ceramide in urine, biopsy samples, and cell cultures, or by the presence of commashaped curvilinear tubular structures, ‘‘Farber bodies,’’ by electron microscopy, for example, in reticuloendothelial cells. The mouse Asah1 gene encoding acidic ceramidase contains 14 exons. To disrupt the gene, a targeting vector was constructed in which exons 3–5 were replaced by a neomycin selection cassette (Li et al., 2002). None of the investigated embryos or newborn mice contained the homozygous mutated Asah1 alleles indicating that homozygous gene disruption was lethal already during embryogenesis. Therefore, phenotypical investigation concentrated on heterozygous mice. Asah1+/ mice were viable for longer than 1.5 years. After birth, they developed normally and exhibited no obvious phenotype. However, at the age of 6 months, significant pathologic abnormalities were observed in the heterozygous mutants. Light microscopic investigation of liver revealed numerous lipid-laden inclusions throughout the parenchyma, most prominent in Kupffer cells. Other organs investigated were also affected but rather modestly. Lipid analysis of several investigated organs resulted into up to twofold elevated ceramide levels in liver and more modest elevations in the other organs of Asah1+/ mice. Results from the Asah1 gene deletion model suggested that normal ceramide metabolism is required to sustain mammalian development. Recently, a report described the presence and deletion of a neutral ceramidase (Asah2) in mice (Kono et al., 2006). Asah2/ mice exerted a normal life span and, different to Asah1/ or Asah1+/ mice, do not show obvious abnormalities or major alterations in total ceramide levels in tissues. However, since the neutral ceramidase was shown to be highly expressed along the brush border of the small intestine, the study of Asah2/ mice emphasized that neutral ceramidase is indeed required for intestinal ceramide degradation of dietary sphingolipids.

2.10 Neuraminidase 1 Deficiency Lysosomal sialidases (neuraminidases, EC 3.2.1.18) cleave sialic acid residues from glycoconjugates including gangliosides. Four genes have been identified to encode for neuraminidases (Neu1-4) in mice (Carrillo et al., 1997; Igdoura et al., 1998; Hasegawa et al., 2000; Kotani et al., 2001; Comelli et al., 2003 ); and Neu1 and human Neu4 were found to localize to lysosoms (Carrillo et al., 1997; Igdoura et al., 1998; Seyrantepe et al., 2004). Until now, only a mouse model for Neuraminidase 1 deficiency has been published (de Geest et al., 2002). The mouse Neu1 gene consists of six exons with the coding region on exons 1–6. De Geest and coworkers inserted a b-geo cassette with the LacZ reporter gene and the neo-selectable marker driven by the PGK promoter (LacZ/PGK/neo) in frame into exon 1. Neu1/ mice transcribe the LacZ gene from the Neu1 endogenous promoter and the truncated signal peptide of neuraminidase 1 protein. No endogenous Neu1 transcript was found in Neu1/ mice. They developed severe nephropathy,

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progressive edema, splenomegaly, and kyphosis, and excrete sialylated oligosaccharides with the urine. Furthermore, progressive deformity of the spine, a high incidence of premature death, and age-related extramedullary hematopoiesis is found in Neu1/ mice, but they lack early degeneration of cerebellar Purkinje cells. Preliminary investigations resulted in no evidence of a secondary accumulation of gangliosides, even not when Neu1/ mice were crossed into Glb1/ mice (A. d’Azzo, personal communication). Hence, either Neu1 contributes to the lysosomal degradation of gangliosides and other sialidases like Neu4, and maybe even the plasma membrane-bound Neu3 (Miyagi et al., 1999; Monti et al., 2000) compensates for Neu1 deficiency; or Neu1 is not involved in ganglioside degradation. The answer to this question will come from the development of mice with deletions for the other neuraminidases and probably their combinations.

3

Conclusions

Great efforts have been undertaken to develop animal models for sphingolipid synthesizing and digesting enzymes in the past 20 years. These models provide new insights into the cellular processes sphingolipids might be involved in. Although different animal models generated by independent researchers in some instances slightly varied in their phenotypes — possibly by lack of defined genetic backgrounds (Tominaga et al., 2004) — in vitro data from the last decades could be substantiated in vivo. By deletion of sphingolipid synthesizing enzymes, it could be convincingly demonstrated that sphingolipids play critical roles in many cellular events in the CNS and PNS such as differentiation of neurons and oligodendrocytes. They significantly modulate axonal branching of neurons, and the stability of axonal/ glial interactions. Furthermore, their importance as signaling molecules in some immunological processes and modulation of cell adhesion could be shown in these models. Mouse models with defects of sphingolipid degrading enzymes were generated to study human storage diseases in vivo. Although not all models match human disease completely, they appear to be tools to study human inherited disorders. With few exceptions, these models provide important mechanistic insights into the development and progression of the disease. Furthermore, these mouse models might also be very valuable for the development of improved treatment strategies such as enzyme replacement, enzyme inhibition, and in particular gene therapeutic approaches (Jeyakumar et al., 2005). However, the molecular mechanisms by which sphingolipids act intracellularly remain largely undefined.

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Index

A Abluminal, 463 Acetylation, 466, 467 Acetylcholine (ACh), 446, 458–462, 465–467, 470, 474–481 Acetylcholine receptor, 465 Acetylcholine release, 458, 460, 465, 467, 474–476, 478, 480, 481 Acetylcholinesterase (AChE), 361, 364, 460, 476 Acetylcholine synthesis, 458, 459, 461, 466, 467, 476, 478, 480, 481 Acetyl-CoA, 461, 462, 466, 467, 470, 476 Acetyltransferase, 25 Acid ceramidase, 391, 678, 680, 691 Acid ceramidase deficiency – ceramide storage, 733 – Farber disease (FD), 733 – Farber lipogranulomatosis, 733 Acid exohydrolases, 672, 676 Acid sphingomyelinase (ASM), 673, 678 Activator proteins – GM2-activator protein, 724, 725 – prosaposin, 725 – saposins, 725 – storage diseases, 725 Acylation, 354, 357, 358, 361, 366 Acyl-CoA, 25, 27, 31 Acyl-CoA: lysophosphatidylcholine acyltransferase, 31 Acyl-CoA oxidase deficiency (ACOX1), 636 1-Acyldihydroxyacetone phosphate, 25 N-Acylethanolamine (NAEs), 386, 391 N-Acylethanolamine-hydrolyzing acid amidase (NAAA), 391 N-Acyl-phosphatidylethanolamine (NAPE), 388, 389, 393, 398, 400 N-Acyltransferase (NAT), 388, 389, 396, 400 ADAM, 361–363, 365 Adenosine, 459, 473

Adenosine triphosphate (ATP), 456, 462, 466, 467, 473, 478 Adenylyl cyclase (AC), 394 Adequate (daily) choline intake, 447, 449 Adrenalectomy, 475 Adrenaline, 480 a-Adrenoceptors, 475, 476 Adrenocorticotropic hormone (ACTH), 458, 475, 480 Adrenoleukodystrophy (ALD) – with Addison’s Disease, 635 – biochemical defect, 635 – childhood cerebral form, 634 – clinical and pathological features, 634 – symptomatic heterozygotes, 635 – therapies, 635, 636 Adrenomedullary catecholamine release, 476 Adrenomedullary tyrosine hydroxylase, 467 Adrenomyeloneuropathy (AMN) – clinical and pathological features, 634 Adult refsum disease (ARD), 636, 650, 655 Aging, 476, 479 – biochemistry, 426 – cognition, 426, 427 Agonist, 465, 470, 479 Akt, 538, 539, 542, 548, 549 Alcoholic liver injury, 457 Alkaline phosphatase, 31, 32 1-Alkyl-2-acyl-sn-glycerol, 25 1-Alkyldihydroxyacetone phosphate, 25 1-Alkylglycerol-sn-3-phosphate, 25 1-Alkyl-2-lyso-sn-glycero-3phosphocholine, 25 Alpha-lipoic acid, 613 Alzheimer – beta-peptide, 358, 362, 363

Alzheimer’s disease (AD), 63, 68, 471, 474, 480 – b-amyloid peptide (Ab), 509 – beta-amyloid precursor protein (bAPP), 571 – Ca2+-independent phospholipase A2 (iPLA2), 570 – central nervous system (CNS), 569 – ceramide, 569, 570 – cholesterol, 570 – ethanol and, 509 – HNE and acrolein, 571–573 – isoprostanes (IsoPs), 573–575 – lipid peroxidation and, 509 – neuroprostanes (NPs), 573–575 – TBA reactive substances, 573 g-Aminobutyric acid, 397, 477 DL-Amino-3-hydroxy-5-methylisoxazole-4-propionic acid (AMPA), 477 Aminophospholipid translocase, 22 Amphiphilic, 461 Amyloid precursor protein (APP), 460 Analgesic, 475 Anandamide (AEA), 386, 389, 391, 396 Anandamide membrane transporter (AMT), 388, 390, 392–396, 398, 400 Anchor – lipid, 354–366 – transmembrane, 361–366 Angiogenesis – VEGF in, 538, 540, 547, 548 Anionic phospholipids, 680 Annexin V, 32 Antagonist, 467, 471, 475, 478, 479 Anterograde transport, 26 Antinociception, 475, 480 Antioxidants, 32 – free radicals, 427 – vitamin E – tocopherols, 428

744

Index Apo-cytochrome c, 201 Apoptosis, 29, 31–34, 200, 204, 213, 476, 479, 537–542, 546, 547, 550 – ceramide role, 187 – dysregulation of nuclear calcium, 190 Apoptosis signal-regulating kinase 1 (ASK1), 400 Arachidonic acid (AA), 24, 28, 29, 33, 451 Arachidonic acid (AA)-omega-6, 228, 230 N-Arachidonoyldopamine (NADA), 395 N-Arachidonoylethanolamine (anandamide), 230, 231 N-Arachidonoylethanolamine. See Anandamide (AEA) O-Arachidonoyl-ethanolamine, 386, 387 2-Arachidonoylglycerol (2-AG), 29, 386–397, 399, 401 2-Arachidonoyl-glyceryl-ether, 386, 387 N-Arachidonoylphosphatidylethanolamine (NArPE), 388, 389, 396 Arylsulfatase A (ASA), 678, 679, 682, 688, 689 Arylsulfatase A deficiency – metachromatic leukodystrophy, 730 – sulfatide storage, 731 Ascorbic acid (AA), 611 Aspirin, 475 Astrocytes, 23, 464 ATPases – Ca2+-ATPase, 33 – Ca2+,Mg2+-ATPase, 33 – Na+,K+-ATPase, 33 Atropine, 475 Autocannibalism, 460, 466 Autonomic ganglia, 465 Autosomal recessive, 681, 684, 687 Autotaxin, 548, 549 – during brain development, 294, 295 – cannabinoids, 296 – ceramide 1-phosphate, cell signaling, 296 – c-terminal region oligodendrocyte adhesion, 294 – diacylglycerol, cell signaling, 298, 299 – epilepsy with mental retardation (EPMR), 302

– epileptic seizures, 301, 302 – expression in brain, 294 – expression in neurological disease, 294, 295 – extracellular signaling, 293, 294 – intracellular signaling, 296 – knockout mice, 295 – lipid phosphate phosphatases – expression in central nervous system, 298, 299 – extracellular functions, 295, 296 – intracellular functions, 296–298 – lipin – cell signaling, 299 – glycerolipid synthesis, 299, 300 – lysophosphatidate, brain tumors, 301 – myelination, 294 – neurite retraction, 300, 303 – receptor knockout mice, 295 – receptors, 294, 295 – synthesis of extracellular lysophosphatidate, 293–295 Axonal elongation, 26

B BACE, 362 Basal forebrain, 476 Base-exchange reactions, 25 Basic fibroblast growth factor, 478 Behavior, 470, 476, 478, 479 Benzodiazepine, 477 Betaine, 447, 448, 454, 456, 457 Betaine aldehyde, 456, 457 Betaine dehydrogenase, 454 Betaine homocysteine methyltransferase, 457 Bilayers, 22, 25, 26, 33, 34 Bioactive lipids, 386, 400 Bioactive sphingolipids – ceramide (Cer), 378, 379 – ceramide 1-phosphate (C1P), 379 – dihydroceramide (dhCer), 378 – sphingosine (Sph), 379 – sphingosine 1-phosphate (S1P), 379 Bis-(monoacylglycero)-phosphate (BMP), 676, 681 Blockade, 474, 475 Blood, 446–459, 470, 473, 475 Blood–brain barrier (BBB), 393, 394, 400, 462, 463 Blood pressure, 474, 475, 478, 480

Body temperature, 475 Bone marrow transplantation (BMT), 686, 688, 690–693 Boron dipyrro-methene difluoride, 115 Bovine brain microvessels (BBM), 393 Brain, 446, 449, 451, 452, 454, 456, 458–468, 470–474, 476–480 – animal – intra-uterine growth retardation, 420 – learning performance, 415 – monkey, 413–415, 417, 419 – mouse, 425 – rat, 411, 413–416, 419, 420, 425, 426 – astrocytes, 413, 428 – baby formula, 411, 415, 417, 418 – blood–brain barrier, 427–429 – brain development, 411, 413–420, 428, 429 – breastfeeding, 417–420 – circulation, 424 – cortex, 413–418, 420, 426 – endoplasmic reticulum, 414 – enzymes, 414, 427–429 – hippocampus, 414, 426 – human – cognition, 426, 427 – foetus, 426 – infant, 415–420, 427, 428 – newborn, 415–419, 428 – membrane, 411, 414, 415, 417–420, 422–429 – microsomes, 426 – mitochondria, 414, 426 – morphology alterations, 504 – neuron, nerve endings, 425, 426 – neurotransmission, 415, 419, 425, 426 – oligodendrocyte, myelin, 412–414, 426, 427, 429 – peroxysomes, 427 – receptor, 419, 420, 425, 426 – uptake, 414, 416, 417, 419–423, 427–429 Brain mitochondria – and aging, 211 – biosynthetic origin of lipids, 207–209 – in ischemia, 211, 212 – lipid composition, 202–205 – in neurodegeneration, 213, 214 – purification and biochemical characterization, 202 Brain oxidative stress. See Oxidative stress (OS)

Index Breast-fed infant, 457 Breast feeding, 457 a-Bungarotoxin, 475, 476 Butyrylcholinesterase (BuChE), 460

C Ca2+ – hexagonal phase of lipids, 201 – and lipid translocation, 207 – in membrane damage, 212 – and PLA2, 212 Ca2+ binding-C2 domain, 230 Ca2+/Calmodulin-dependent protein kinase II (CaM-kinase II), 461, 462, 465 Ca2+-independent phospholipase A2 (iPLA2), 570 Calcineurin, 356, 357 Calcium – homeostasis in the nucleus, 189 – Sodium-calcium exchanger/GM1 complex, 179 Calf pulmonary artery endothelium (CPAE), 393 cAMP response element (CRE), 386 Campylobacter jejuni, 127 Cannabinoid receptors (CB1), 231, 386, 394–396, 400 Cannabinoids – anandamide, 537, 538 – N-arachidonoyl-ethanolamide, 537 – 2-arachidonoyl-glycerol, 537, 538 – D9-tetrahydrocannabinol, 537 Capsaicin, 395 Cardiolipin (CL), 479 – in apoptosis, 204 – in Barth syndrome, 201 – biosynthesis, 209 – in brain and hearth, 202, 205 – in cerebral ischemia and oxidative stress, 14, 15 – in contact sites, 203 – and cytidine-5’diphosphocholine, 212 – distribution in the inner mitochondrial membrane, 204 – enrichment in brain mitochondria, 205 – fatty acid composition, 204, 205 – liposomes of, 205 – and mitochondrial membrane potential, 203 – in mitochondrial protein import, 203

– – – –

and nonyl acridine orange, 203 and peroxidation, 212 and proton pump, 205 release from brain mitochondria, 212 – remodeling by transacylation, 202, 209 Cardiovascular, 474 Carnitine, 459, 463 Caspases, 33 Catecholamine, 465, 475, 476 Cationic glycosphingolipids, 111 Caveolae, 365 Caveolin, 365 CD1 molecules, 674, 680 CDP-choline, 23–25, 32, 33 – CDP-choline: 1,2-diacylglycerol cholinephosphotransferase, 472, 473 CDP-ethanolamine, 23, 25, 451, 470, 478 Cell-mediated ‘‘cross correction,’’ 672, 692 Cell proliferation, 30 Cell-type specific expression of glycosphingolipids, 682 Central nervous system (CNS), 388, 393–395, 397–400, 464, 672–693 Ceramidase, 233 Ceramide kinase (CERK), 246, 247 Ceramide 1-phosphate (C1P), 379 Ceramide, 228, 233, 234, 673, 674, 676, 678–680, 682, 688, 691–693 – ceramide/diacylglycerol ratio, 185 – gangliosides, 705, 714 – inducer of apoptosis, 187 – myriocin, 505 – neuraminic acid, 705 – neutral glycosphingolipids, 705 – product of ceramidase, 187 – sulfatides, 705 Ceramide transfer protein (CERT), 249 Cerebellar granule neurons, 479 Cerebral edema, 478 Cerebral ischemia, 477–480 Cerebroside sulfotransferase deficiency – infertility, 716 – oligodendrocyte differentiation, 715–717 Chemical chaperones, 692, 693 Chemical sympathectomy, 474 Chemical synthesis, neural sphingolipids – biological investigation, 113–118 – a-Gal-Cer, 119 – Golgi sialyl transferase(s), 121 – hexosaminidase hydrolysis, 120

– lyso-GM3 dimer, 120 – myelin-associated glycoprotein (MAG), 120 – Neu5Ac residue, 122 – sialyl(a2!1)sphingosine, 120 – soluble ceramides, 122 – thio-Lac-Cer, 118 – un-natural sphingolipids, 119, 121 Cholera toxin B subunit – cytochemical agent for GM1 ganglioside in endonucleus, 179 – cytochemical agent for GM1 ganglioside in nuclear envelope, 188 Cholera toxin holo-enzyme, 128 Cholesterol, 22, 224, 225, 228, 232–234, 354, 356–358, 362, 395, 536, 549, 550, 673, 681, 682, 687 – ABCA1, 506 – association with sphingomyelin, 179 – in brain mitochondria, 210 – component of nuclear membranes, 177 – conversion of, 61, 64 – metabolism in brain, 210 – in neurodegeneration, 213 Cholesterol biosynthesis, 245, 246 Cholesterol synthesis, 645 Cholesteryl esters, 549 Choline acetyltransferase (ChAT), 458, 461, 462, 466, 467, 476 Choline-deficient diet, 447, 449 Choline dehydrogenase, 454, 456 Choline glycerophospholipids, 22, 24–28, 32, 33 Choline homeostasis, 232 Choline kinase (CK), 451, 461, 462, 466, 467 Choline oxidase, 454, 456, 457 Cholinephosphotransferase, 472 Choline reuptake, 458 Cholinergic nerve terminals, 459, 461 Choline-rich diet, 476 Choline-transporter-like protein (CTP), 451, 466–468, 470–474 Choloylglycine hydrolase, 391 Choroid plexus (CP), 459 Circulating precursor, 451, 466, 468, 470, 471, 473 Citicoline, 480 Coenzyme Q (CoQ), 614, 615 Coincidence detection, 230 Collision-induced dissociation (CID), 10

745

746

Index Complex sphingolipids – cationic glycosphingolipids, 111 – gangliosides, 101, 103–109 – neutral glycosphingolipids, 110 – sphingoid bases and ceramide, 100, 101 – sulfatides and sulfoglycosphingolipids, 104–108, 110 Concentrative nucleoside transporter (CNT), 470, 473 Consumption, 447, 457, 470, 471, 476 Contraction, 30 Cortisol, 458 COS-1 cells, 451 CREB, 476 Cross-talk, 28, 29 Cross talks between sPLA2 and other PLA2s – COX-1 and COX-2, 526 – prostaglandin synthesis, 526 CTP: phosphocholine cytidylyltransferase, 471, 472 CTP synthase, 474 Cyclases – adenylate cyclase, 31–33 – guanylate cyclase, 31 Cyclic 3,5-nucleotide phosphohydrolase, 31 Cyclooxygenase, 537, 538 Cytidine, 470, 473, 474, 477 Cytidine deaminase, 477 Cytidine diphosphate (CDP), 473 Cytidine-5’-diphosphocholine (CDPcholine), 477–479 Cytidine monophosphate, 477 Cytidine triphosphate (CTP), 471–474 Cytidylyltransferases – choline, 23, 24 – ethanolamine, 23, 24 Cytochrome C, 200–202, 205, 212 Cytokines, 27, 31, 33 Cytoplasm, 456, 457 Cytoprotective, 477 Cytosol, 23 Cytosolic form, 471

D D-Bifunctional enzyme deficiency (DBP), 636, 637 Decarboxylation, 24, 34 Demyelination, 689, 691 Dendritic spine, 470, 476 Dephosphorylation, 23 Depolarization, 460, 464, 465 Depolarization-induced suppression of excitation (DSE), 397, 398

Depolarization-induced suppression of inhibition (DSI), 397, 398 D-erythro-sphingopine, 673 Diacylglycerol (DAG), 23, 24, 27–29, 31–33, 391, 392, 396 – activation of protein kinase C, 187 – conversion to phosphatidic acid, 184 – kinase, 180, 184, 228 – migration to the nucleus, 184 – product of phospholipase C, 183 – ratio relative to ceramide, 187 Diacylglycerol lipase (DAGL), 391–393, 396 Diet – alcohol, 425 – animal feeding, 413, 415 – bioavailability, 429 – capsule, 417, 421, 424, 429 – fish, 416, 427 – food, 421–425, 428, 429 – linseed, 413, 415, 416 – milk, 411, 417–420, 423, 428 – oils – canola, 411, 416, 429 – rapeseed, 411, 413, 415, 416, 429 – seafood, 422–425, 429 – shellfish, 416, 427 – soybean – walnut, 413, 429 Dietary reference intake (DRI), 447, 449 Dihydroceramide (dhCer), 378 Dihydroxyacetone phosphate, 25 Directional transport, 393, 394 Disease – cherry-red spots, 683 – Fabry’s, 686, 687 – Farber, 691 – Gaucher, 689, 690 – Krabbe, 690, 691 – myelin, 690 – Niemann-Pick disease (type A and B), 687, 688 – Niemann-Pick type C, 673, 687 – Sandhoff, 682, 684–686, 691, 693 – Schindler, 684 – Tay–Sachs, 684–686, 693 Dizocilpine, 477 DNA interaction with phospholipids, 180 Docosahexaenoic acid (DHA), 25, 28, 30, 31, 451, 452, 454, 467, 468, 470, 471, 616, 617 Docosahexaenoic acid (DHA)-omega3, 228, 230

Docosahexaenoic acid synthesis, 639–641 Docosanoids, 29, 31 Docosatrienes, 31 Dopamine, 460, 470, 479 – oxidation of, 586, 587

E Eicosanoids, 28, 29, 31, 180, 183, 184, 189 – leukotrienes, 537, 538, 540, 541 – prostaglandins, 537–541 Eicosapentaenoic acid (EPA), 451, 452, 471 Electrogenic, 463 Electrostatic interactions, 230 Endocannabinoids – anandamide, 83, 86–88 – 2-arachidonoylglycerol, 83 Endocytosis, 395 Endonuclear domains, 175–177, 179, 180, 184, 185 Endoplasmic reticulum, 23–25, 34 – continuity of outer nuclear membrane, 176 b-Endorphin, 458, 475 Endotoxin, 474, 477, 480 Endotoxin-induced multiple organ injury, 477 End-stage renal disease, 457 Energy-dependent, 459, 463 Energy homeostasis – food intake, 76, 86, 87 – gastric secretion, 87, 88 – intestinal motility, 87, 88 – nutrient assimilation, 87 Enzyme, 447, 449, 451, 454, 456–462, 466–468, 470–474, 477, 478, 481 Ependymoma, 536, 539, 545, 546 Epidermal growth factor (EGF), 538, 546 Epilepsy, 357 – influence of neuroactive steroids on, 61, 65–68 – temporal lobe epilepsy, 61, 62 – treatment of, 68 Epileptic seizures, 476 Equilibrative nucleoside transporter (ENT), 473 17b-Estradiol, 398 Estrogen. See 17b-Estradiol Estrogen receptor (ER), 398 Ethanol, 457 Ethanolamine, 451, 460, 462, 471 Ethanolamine glycerophospholipids, 21–34 Ethanolamine kinase, 462

Index Ethanolaminephosphotransferase (EPT), 472 Experimental autoimmune encephalomyelitis (EAE) – lipid microarray in, 595, 597 – sulfatide-reactive T cells, 597 – sulfatide-specific antibodies, 595, 597 Extracellular fluid (ECF), 459, 461, 463 Extracellular signal-regulated kinase (ERK), 399, 400

F Fabry disease – a-galactosidase A deficiency, 727 – globotriaosylceramide storage, 727 Facilitated diffusion, 459, 461–463 Facilitated transport, 390 Farnesyl, 358, 359 Fatty acid amide hydrolase (FAAH), 388–390, 396 Fatty acids, 451, 452, 454, 459, 462, 466–468, 470–472, 478 – deacylation and reacylation cycle, 209 – desaturases, 411, 428 – import into mitochondria, 208 – in ischemia, 212 – monounsaturated fatty acids – nervonic acid, 412 – oleic acid, 412 – in neurodegeneration, 212 – in phospholipids of brain mitochondria, 203, 206 – polyunsaturated fatty acids – omega-3 fatty acids –alpha-linolenic acid (ALA), 411–422, 424–426, 428, 429 –docosahexaenoic acid (Cervonic acid) (DHA), 411–429 –eicosapentaenoic acid (EPA), 411, 412, 416–419, 421–427, 429 – omega-6 fatty acids –arachidonic acid (ARA), 411, 414–420, 422, 423, 426, 428, 429 –linoleic acid (LA), 411, 412, 420, 427 – saturated fatty acids – lignoceric acid, 412, 429 – palmitic acid, 412 – stearic acid, 412

– trans fatty acids, 415, 420, 424, 427 Fetal alcohol syndrome – apoptotic neurodegeneration, 508, 509 – cell replication, 508 – EGF receptor, 508 – ganglioside GM1, 508 – 4-hydroxynonenal, 508 – membrane fluidity, 508 – Na+-K+ pump, 508 – phospholipids, 508 Fetus, 457, 476 Fingolimod (FTY-720) – S1P receptor, 598 Fishy odor, 456 Focal adhesion kinase (FAK), 394 Folic acid, 452, 457 Food and Nutrition Board (FNB), 447

G a-Galactosidase A, 676 a-Galactosidase A deficiency – Fabry disease, 272 – globotriaosylceramide (Gb3Cer) storage, 727 b-Galactosidase 1 deficiency – GM1-gangliosidosis, 721 – GM1 storage, 721 – morquio B disease, 721 Galactosylceramidase deficiency – globoid cell leukodystrophy (GLD), 727–728 – Krabbe disease, 727, 728 – psychosine storage, 728 – twitcher mouse, 728 Galactosylceramide-b-galactosidase, 678, 679, 683, 690, 691, 693 Galactosylceramide synthase deficiency – astrogliosis, 714 – myelination defects, 714 Galactosylsphingosine, 682, 691 b1,4-GalNAc transferase deficiency – axon/myelin interaction, 709 – GM3/GD3 accumulation, 708 – infertility, 708 – myelin associated glycoprotein, 709 Ganglionic, 475, 480, 481 Gangliosides, 233 – gangliosides of endonuclear domains, 180 – gangliosides of the nuclear envelope, 179 – GM1 complex with sodiumcalcium exchanger, 179

– GM1 detection in endonucleus, 177 Gaucher cell, 690 Gaucher disease – b-glucosidase deficiency, 728–730 – glucosylceramide storage, 728 Gavage, 468, 470 GD3 synthase deficiency – b-series gangliosides, lack of, 709 Gene expression, 22, 31, 33 Geranyl, 358 Gerbil, 468, 470–473, 477, 478 Glioblastoma, 536, 548 Glioma, 364 Globoid cell leukodystrophy (GCL), 690 Glucose, 459, 463, 467, 475, 476, 478 b-Glucosidase deficiency – Gaucher disease (GD), 728–730 – glucosylceramide storage, 728 Glucosylceramide-b-glucosidase, 676, 680–682, 689, 690 Glucosylceramide synthase deficiency – embryonic lethality, 713 Glucosylceramide synthase deficiency, conditional in brain – brain maturation defect, 714 – neural dysfunction, 713 – neuron differentiation defect, 714 Glucosylceramide synthase deficiency, conditional in epidermis – skin barrier defect, 714 Glutamate decarboxylase (GAD65), 357, 360 Glutamate uptake, 31 Glutathione (GSH), 613, 614 Glycerolipids – diacylglycerol, 183, 184 – inositol phosphoglycerides, 181–183 – phosphatidic acid, 183, 184 – phosphatidylcholine, 181, 183, 184 – phosphatidylethanolamine, 180, 181 – phosphatidylinositol, 181 – phosphatidylinositol 4,5 bisphosphate, 179 – phosphatidylserine, 178 Glycerophosphocholine (GPC), 32, 446, 451, 452, 458, 471 Glycerophosphoethanolamine, 471 Glycerophospholipid biosynthesis, 247, 248 Glycerophospholipids, 207, 209, 213 Glycocalix, 674, 675 Glycolipids, 22 Glycophospholipid, 366

747

748

Index Glycoprotein, 357, 361, 363, 364, 366 Glycosphingolipids (GSLs), 175, 178, 187–190, 395, 673–676, 678, 681, 682, 684, 705, 707, 720 – galactosyltransferase, 507 – gangliosides (GM1, GD1b, GT1b, GD1a), 507 – lipid rafts, 506 – sialidases, 507 – sphingosine, 506 Glycosyl-phosphatidylinositol (GPI), 354, 361–366 Glycyl-glutamine, 474 GM2-activator, 676–682, 684–686 GM1-b-galactosidase, 676, 681, 683, 684 GM2-gangliosidosis – 0-variant of, 685 – AB-variant of, 684 – B-variant of, 684, 685 GM3 synthase deficiency – human infantile-onset epilepsy syndrome, 710 – insulin receptor phosphorylation, 711 – insulin sensitivity, 711 Golgi sialyl transferases, 121 G-protein-coupled receptors, 394, 399, 470 – acetylcholine receptors, 82, 86 – cannabinoid receptors, 82, 83 – glycine receptors, 82 – orphan G-protein-coupled receptor 55 (GPR55), 82, 83, 90 – orphan G-protein-coupled receptor 119 (GPR119), 82, 83, 87 G-proteins, 32, 33 Group I metabotropic glutamate receptor, 391 Growth associated protein 43 (GAP-43), 354, 355, 357, 366 GTPase, 360 Guinea pig, 465, 481

H Halothane, 233 Hamster, 472 Heart, 464, 474, 481 Heart rate, 474 HeLa cells, 472 Hemicholinium-3 (HC3), 458, 459, 462–464, 467 Hemodialysis, 457 Hemorrhage, 474, 478 HexA deficiency – GM2-storage, 724 – Tay–Sachs disease, 723

Hexagonal phase, 32 HexB deficiency – globoside storage, 723 – GM2-storage, 723 – Sandhoff disease, 723 b-Hexosaminidase A, 676, 678, 684, 685, 693 b-Hexosaminidase B, 676, 684 Hexosaminidase hydrolysis, 120 b-Hexosaminidase S, 676, 684 High-affinity choline transporter (CHT), 460, 463 Hippocampal CA1 pyramidal cells, 476 Hippocampus, 476, 479 HNK-1 antibody, 127 Homeostasis, 22, 32, 33 Homocysteine, 447, 457 Hormone release, 32 Hot-plate test, 475 Human umbilical vein endothelial cells (HUVEC), 393, 398 Hydrophilic, 461 b-Hydroxybutyrate dehydrogenase, 33 6-Hydroxydopamine, 479, 481 Hyperglycemic response, 475 Hypotension, 447, 474, 480 Hypotensive, 474, 475, 478 Hypothalamic-pituitary-adrenal (HPA) axis, 398 Hypothermia, 475

I Ikaros, 390 Immortalized mouse brain endothelial microvessels, 463 Immortalized rat brain endothelial microvessels, 459, 463 Immunoglobulin, 364 Immunohistochemistry, 64 Impoverished environmental conditions, 479 Infantile Refsum disease (IRD), 633 Infarct volume, 478 Inflammation, 29–31, 33 – neurogenic inflammation, 89, 90 – neuroinflammatory disorders, 88, 89 Inflammatory signaling pathways, 517–528 Inner nuclear membrane – locus of nuclear sodium-calcium exchanger/GM1 complex – unique lipid composition, 176 Inositol glycerophospholipids, 22, 28 Inositol phosphoglycerides – canonical phosphoglycerides, 183 – D-3 phosphoglycerides, 183

Inositol (1,4,5)-tri-phosphate [Ins (1,4,5)P3], metabolite of PIs, 228 Inositol trisphosphate (IP3), 458, 465 – product of phospholipase C, 183 – receptors for on inner nuclear membrane, 183 In situ brain perfusion technique, 459, 463 Insulin, 475, 481 Interleukin-4, 32 Intracerebroventricular, 474, 475, 480 Intracisternal, 474 Intramuscular, 474 Intraperitoneal, 474–476, 478, 479, 481 Intrathecally, 475 Intravenous, 474, 475, 477, 480 Invasiveness – cell adhesion in, 544, 547 – cell motility, 541, 543, 548 – metalloproteinases in, 547 Ion channels, 22, 26, 31, 34 – potassium channels, 90 – transient receptor potential (TRP) channels melastatin type-8 (TRPM8), 79, 82, 84 – transient receptor potential (TRP) channels vanilloid type-1 (TRPV1), 79, 82 – voltage activated calcium channels, 85 Isolated perfused liver, 456, 457 Isolation procedures – nuclear envelope, 177 – nuclear membranes, inner and outer, 178 – nuclei, 177 Isoprene, 358, 359 Isoprostanes (IsoPs), 573–575, 608, 609

J c-Jun N-terminal kinase (JNK), 400

K Kennedy cycle, 23, 451, 460, 462, 467, 472, 477 3-Ketosphinganine, 242 Kidney, 451, 454, 456, 463, 464, 473 Kinases – cAMP-dependent, 23 – choline, 23, 24 – diacylglycerol, 23, 24, 28 – ethanolamine, 23, 24 – protein kinase C, 24, 28, 29, 31, 32 – Ras-Raf-MAP, 32 – tyrosine, 32

Index Krabbe disease – galactosylceramidase deficiency, 727 – globoid cell leukodystrophy (GLD), 726 – psychosine storage, 726

L Lactating rats, 476 Lactation, 457, 471 Lamellar phase, 32 Lamin, 358, 363 L-1,2-dehexadecyl-sm-glycero-3phospho ethanolamine (DHPE), 121 L-1,2-dipalmitoyl-sm-glycerophosphoethanolamine (DPPE), 121 Learning, 31 Learning and memory, 476 Leptin, 390 Leukotriene C4, 479 Leukotrienes, 31 LIGA-20 in neuroprotection, 190 Ligand-gated ion channel, 465 Linoleic acid, 24 Linolenic acid, 24 Lipid composition – endonuclear domains, 179, 180 – nuclear envelope, 178, 179 – whole nuclei, 178 Lipidomics, 23 – database and analytical tools, 18 – electrospray ionization (ESI), 5 – lipid molecule identification, 4 – mass spectrometry – collision-induced dissociation (CID), 10 – FTICRMS, untargeted and global analyses, 7 – ion spectrum, phospholipids, 10, 12, 15, 16 – Lipid Search, 12 – [M þ HCO2]-ions, 9, 10 – multiple reaction monitoring (MRM), 14 – m/z values, 9, 10 – phosphatidylethanolamine, 13, 14 – practical methods, 6, 7 – precursor ion/neutral loss scanning, 8 – sensitivity, 5 – shotgun LC-MS/MS, untargeted and global analyses, 7, 8 – targeted methods, 8 – UPLC-MS, untargeted and global analyses, 7

Lipid peroxidation – 4-hydroxynonenal (HNE) in, 587 Lipid peroxidation and antioxidants – cyclooxygenase (COX-2), 508 – cytochrome P4502E1, 508 – glutathione peroxidise, 507, 508 – grape polyphenols, 508 – green tea antioxidants, 508 – lipid hydroperoxides (malondialdehyde and 4-hydroxynonenal), 507 – metal ions, 507 – N-methyl-D-aspartate (NMDA), 508 – microsomal ethanol-oxidizing system (MEOS), 508 – nitric oxide synthase, 508 – vitamin E, 508 Lipid rafts, 395, 397, 400 Lipids, 452, 471 – Alzheimer’s disease (AD) – beta-amyloid precursor protein (bAPP), 571 – Ca2+-independent phospholipase A2 (iPLA2), 570 – central nervous system (CNS), 569 – ceramide, 569, 570 – cholesterol, 570 – HNE and acrolein, 571–573 – isoprostanes (IsoPs), 573–575 – neuroprostanes (NPs), 573–575 – TBA reactive substances, 573 – cholesterol, 415, 423, 426, 427 – lysophosphatidic acid (LPA), 594 – lysophospholipids, 594 – microdomain, 362, 363, 365 – phosphatidylcholine, 426, 427, 429 – phosphatidylserine, 414, 419, 426 – phospholipid asymmetry, 575 – phospholipids, 411, 413, 414, 416, 418–420, 423, 425, 426, 429 – plasmalogens, 426 – plasma membranes – phospholipid transport and synthesis, 566 – phospholipid transporter identification, 566–569 – types, 565, 566 – raft, 357, 358, 362, 363, 365 – sphingolipids, 597, 598 – triglycerides, 416 – in vitro and in vivo, 575, 576

Lipid Search, 7–10, 12, 17, 18 Lipoxygenase, 537, 538 15-Lipoxygenase, 31 Lithium, 459 Liver, 446, 449, 451, 452, 454, 456, 457, 460, 462, 464, 470, 472, 473 Long-chain fatty acyl-CoA synthetase, 470 Long term depression (LTP), 397 Long-term potentiation, 33, 397 Low-affinity choline transport, 464 Low-density lipoproteins, 32 Luminal, 463 Lung, 464 Lyso-GM3 dimer, 120 Lysophosphatidic acid (LPA), 24, 32, 33, 548, 549 Lysophosphatidylcholine (LPC), 452 – precursor of lysophosphatidate, 293, 294, 296 – receptors, 290–293 – source of, 301 Lysophospholipids, 519 Lysosomal storage diseases, 672, 673, 692, 693

M Madin-Darby Canine Kidney (MDCK) cells, 249 MAG. See Myelin-associated glycoprotein Malondialdehyde (MDA), 609 MAPK, 476 Marathon running, 457 Mass spectrometry, lipidomics – collision-induced dissociation (CID), 10 – FTICRMS, untargeted and global analyses, 7 – ion spectrum, phospholipids, 10, 12, 15, 16 – lipid search, 12 – [M þ HCO2]-ions, 9, 10 – multiple reaction monitoring (MRM), 14 – m/z values, 9, 10 – phosphatidylethanolamine, 13, 14 – practical methods, 6, 7 – precursor ion/neutral loss scanning, 8 – sensitivity, 5 – shotgun LC-MS/MS, untargeted and global analyses, 7, 8 – targeted methods, 8 – UPLC-MS, untargeted and global analyses, 7 Medial prefrontal cortex, 464

749

750

Index Medulloblastoma, 536, 542, 545–547, 550 Membrane-bound form, 461, 471 Membrane curvature – breakdown by, 230 – generation of lysophosphatidylcholine (LPC), 231 – phospholipase A (PLA) – phospholipase D (PLD) Membrane fluidity – cholesterol, 505 – linoleic:arachidonic acid (18:2/20:4) ratio, 505 – phospholipids saturated fatty acids, 505 Membrane fusion, 228, 231, 233 Membrane glycerophospholipids, deacylation-reacylation, 519 Membranes, 445, 452, 454, 456, 458, 460–468, 470–473, 477, 478, 481 – membrane fusion, 32, 34 – membrane trafficking, 30, 32 Memory, 31, 33 Meningioma, 537, 540, 545 Metabolism – acylethanolamide biosynthesis, 76–80 – acylethanolamide inactivation, 76–80 Metachromatic leukodystrophy – arylsulfatase A deficiency, 730 – sulfatide storage, 731 Metachromatic lukodystrophy (MLD), 688, 689 Methionine, 447, 448, 452, 457 3-Methoxy-4-hydroxyphenylglycol, 479 Methyl-arachidonoyl fluorophosphonate (MAFP), 386 Methylation, methyltransferases, 24 Methyl-b-cyclodextrin (MCD), 395 N-Methyl-D-aspartate (NMDA), 397, 476, 477 1-Methyl-4-phenyl1,2,3,6-tetrahydropyridine (MPTP) – inhibition of mitochondrial function by, 585 – MPP+ reactive metabolite of, 585 – toxic effects of, 585 Metillycaconitine, 475 Microdomains, 22 Microglia, 23 Microsome, 472 Middle cerebral artery, 478 Mitochondria, 456, 457, 467

Mitochondria-associated membranes (MAM), 200, 202, 207, 208, 214 Mitogen-activated protein kinase (MAPK), 394, 400 Mitogenesis, 30 Modification, post-translational, 354, 355, 358 Molecular species, 23, 24, 26, 28, 29, 34 Monoacylglycerol acyltransferase, 32 Monoacylglycerol lipase (MAGL), 392, 393, 396 Monounsaturated fatty acid, 452 Morphine, 475, 477, 480 Morquio B disease – b-Galactosidase deficiency, 721 – GM1-gangliosidosis, 721 Morquio type B disease, 683, 684 Morris water maze, 476 Mortality rate, 478 Mouse, 463, 464, 472–474 MPTP-like toxicity – b-carbinoles, 586 – n-hexane, 586 – hydrocarbons, 586 – isoquinolines, 586 mtDNA, 201, 211 Multiple sclerosis – autoimmunity, 595 – Borrelia burgdorferi in, 595 – CD1 molecule, 597 – chlamydi pneumoniae in, 595 – glycolipid autoantigens in, 595 – glycolipid reactive lymphocytes, 595, 597 – GM1, 594, 597 – molecular mimicry in, 595 – mycoplasma pneumoniae in, 595 – sulfatide-specific antibodies, 595 Muscarinic acetylcholine receptor (mAChR), 459, 465, 479 Myelin, 22, 26, 674, 682, 689, 691 – abnormalities in, 655–659 – cholesterol, 594 – galactocerebroside (GalC), 594 – galactosylceramide, 594 – gangliosides, 594 Myelin-associated glycoprotein (MAG), 120, 127 Myelin lipids – biogenesis and maintenance, 251–253 – cholesterol biosynthesis, 245, 246 – oligodendrocyte physiology – fatty acids (FAs), 253 – gangliosides, 255, 256 – sphingolipids, 253–255

– phospholipid biosynthesis – glycerophospholipid, 247, 248 – phosphosphingolipid, 243, 246 – sorting and transport – ceramide transfer protein (CERT), 249 – fatty acids, 250 – GalCer, 249 – polarized cells, 249, 250 – sphingolipids/cholesterol, intracellular transport, 250, 251 – sphingolipid biosynthesis – ganglioside, 242–245 – 3-ketosphinganine, 242 Myelin protein – myelin basic protein (MBP), 595 – myelin oligodendrocyte proein (MOG), 595 – proteolipid protein (PLP), 595 Myristate, 354–358, 361 Myristoylation, 355–357

N a4b4 nAChRs, 466 a7 nAChRs, 475 N-acylethanolamine (NAE), 231 N-acylphosphatidylethanolamine (NAPE), 231 NADPH oxidase, 31 Naloxone, 474, 475 Natural Killer T (NKT) cells – alpha-galactosylceramide (a-GalCer), 596 – Borrelia burgdorferi’s glycolipid-2 (BbGL-2), 595 – CD1d, 596, 597 – glycolipid antigens, 596 – isoglobotrihexosylceramide (iGb3), 596 – non-polymorphic MHC-I-like molecules, 596 – sphingomonas antigen GalAGSL, 596 nDNA, 201 Neonatal, 476, 477 Neonatal adrenoleukodystrophy (NALD), 633 Neonatal alcohol exposure, 476 Nerve growth factor (NGF), 386, 464, 468, 470, 472, 476 Nerve terminals, 451, 458–461, 467 Neural cell adhesion molecule (NCAM), 354, 361, 364

Index Neural sphingolipids – analytical biochemistry – dimethylsulfoxide (DMSO), 126 – gas–liquid chromatography (GLC), 124 – high performance thin layer chromatography (HPTLC), 123 – matrix-assisted laser desorption ionization timeof-flight mass spectrometry, 125 – proton nuclear magnetic resonance, 125, 126 – sensitivity, 123 – cellular and subcellular localization, vertebrates – alkali-labile gangliosides, 132, 133 – Gal-Cer, 129 – gray matter, 130, 136 – N-glycolylneuraminic acid, 130 – sphingomyelin, 133 – chemical synthesis – biological investigation, 113–118 – a-Gal-Cer, 119 – golgi sialyl transferase(s), 121 – hexosaminidase hydrolysis, 120 – lyso-GM3 dimer, 120 – myelin-associated glycoprotein (MAG), 120 – Neu5Ac residue, 122 – sialyl(a2!1)sphingosine, 120 – soluble ceramides, 122 – thio-Lac-Cer, 118 – un-natural sphingolipids, 119, 121 – complex sphingolipids – cationic glycosphingolipids, 111 – gangliosides, 101, 103–109 – neutral glycosphingolipids, 110 – sphingoid bases and ceramide, 100, 101 – sulfatides and sulfoglycosphingolipids, 104–108, 110 – developmental profiles, brain – age effect, 142–144 – Gal-Cer, 142

– gangliosides and glycosphingolipids, 144–152 – fatty acid and long chain base composition, 152–154 – immunochemical methods, in situ detection, 126 – Campylobacter jejuni, 127 – Cholera toxin holo-enzyme, 128 – HNK-1, 127 – immune response, 126 – myelin-associated glycoprotein (MAG), 127 – Salmonella minnesota mutant R595, 126 Neurite outgrowth, 470, 472 Neuritogenesis, 29 Neuroactive steroids – allopregnanolone, 66, 68 – alteration of the excitability of central nervous system, 60, 67 – anaesthetic effects, 67 – anticonvulsive effects, 67 – anxiolytic effects, 67 – circulating concentrations, 68 – contribution to development of brain, 62, 64, 67, 68 – contribution to functions of brain, 64 – contribution to vitality of brain, 64 – dehydroepiandrosterone (DHEA), 64, 65, 67, 68 – dehydroepiandrosterone sulfate (DHEAS), 64, 65, 67, 68 – effects on cognitive function, 68 – genomic actions, 60 – influence on transmitter-gated ion channels, 60, 67 – inhibition of N-methyl-Daspartate (NMDA) receptor function, 60, 67 – inhibition of the inhibitory effects of GABA, 67 – interaction with neuropeptide receptors, 67 – modulation of GABAA receptor function, 67 – neuroprotective functions, 65 – non genomic actions, 65, 67 – pathophysiological relevance, 65 – physiological relevance, 65 – potentiation of inhibitory effects of GABA, 67 – pregnenolone, 61, 64, 65 – pregnenolone sulfate, 64

– responsiveness to specific inhibitors, 61 – synthetic derivates of, 68 – 3a,5a-tetrahydroprogesterone, 61, 66, 68 Neuroanatomic, 476, 479 Neuroblastoma, 536, 539, 542, 545–547, 550 Neurochemical, 476 Neurodegeneration, 29, 31 – Alzheimer’s disease, 518, 519, 521–523, 526 – cerebral ischemia, 523, 526 Neurodegenerative inherited diseases – Gaucher disease (GD), 726, 728–730 – GM1-gangliosidosis, 721, 722 – GM2-gangliosidosis, 724 – Krabbe disease, 726–728 – metachromatic leukodystrophy, 726, 730 – Morquio B disease, 721 – Niemann-Pick disease (NPD), 731, 732 – Sandhoff disease, 723 – Tay–Sachs disease, 723, 724 Neuroendocrine, 475 Neurological disorders – adrenoleucodistrophy, 426 – epilepsy, 89 – glutamate-mediated excitotoxicity, 89 – handicap, 425, 427 – Huntington’s disease, 89 – multiple sclerosis, 427 – neuroinflammation, 88, 89 – neuronal ceroid lipofuscinosis, 427 – Parkinson’s disease, 89, 427 – stroke, 89 – traumatic brain injury, 88, 89 Neuromodulin, 354 Neuronal migration, abnormalities in, 651–654 Neuronal plasticity, 30 Neurons, 23, 26, 29–33 Neuroprostanes (NPs), 573–575, 609 Neuroprotectins, 31 Neuroprotection – D-b Hydroxybutyrate, 586 – influence of neuroactive steroids on, 68 Neuroprotective, 477–479 Neuropsychiatric disorders – depressive disorders, 60 – effects of neuroactive steroids, 67

751

752

Index – ethanol withdrawal, 68 – fatigue during pregnancy, 68 – multi-infarct dementia, 68 – post partum depression, 68 – premenstrual syndrome, 68 Neurotoxicology, 415, 417–420, 424, 427 Neurotransmitter release, 397 Neutral glycosphingolipids, 110 Neutral protease, 33 Neutral sphingomyelinase, 400 Newborn, 457 N-glycolylneuraminic acid, 130 Nicotine, 465, 476, 477 Nicotinic acetylcholine receptor (nAChR), 465, 466, 474–477 Niemann-Pick cell, 688 Niemann-Pick disease (NPD) – sphingomyelinase deficiency, 731–733 – sphingomyelin storage, 731–733 Nimodipine, 478 Nitric oxide (NO), 393, 465 Nitric oxide synthase (NOS), 393, 394 Noladin ether. See 2-Arachidonoylglyceryl-ether Non-degradable compounds, storage of, 682 Noradrenaline, 479, 480 Normotensive, 474 Nuclear envelope – Calcium regulatory mechanisms, 179 – inner nuclear membrane, 179 – outer nuclear membrane, 179 Nuclear membrane, 452 Nuclear pore complex, 176–178, 189 Nuclear structure, 175–177 Nuclear transcription factor kB (NfkB), 32 Nucleoside diphosphate kinase (NDPK), 473, 474 Nucleus as ‘‘cell within a cell,’’ 175, 191 Nutritional, 457

O Oleic acid, 24 N-Oleoylethanolamine (OEA), 386 Oligodendrocyte physiology, lipids – fatty acids (FAs), 253 – gangliosides, 255, 256 – sphingolipids, 253–255 Oligodendrocytes, 23, 682, 689 – oligodendrocyte progenitor cell (OPC), 598 Oligodendroglioma, 539, 542 Omega–3 fatty acid, 452, 466, 468, 471

Oncogenesis, 30 Organic cation/carnitine transporter (OCTN2), 463 Organic cation transporter (OCT), 459, 461, 463, 464 Osmotic stress, 475 Outer nuclear membrane, 176, 177, 179, 189, 191 Ovary, 464 Oxidative damage – free radicals, role of, 586 – glutathione, role of, 586 – monoamine oxidase (MAO), role of, 586 – nitric oxide (NO), role of, 586 – protection against, 587 – reactive oxygen species (ROS), role in, 586 – superoxide dismutase (SOD), role of, 585, 586 Oxidative signaling pathways – Ca2+-dependent enzymes, 518, 520 – MAPK pathway, 521 – NADPH oxidase, 521 – reactive oxygen species (ROS), 518 Oxidative stress (OS) – aging brain, 623, 624 – definition, 604 – enzymatic lipid peroxidation – cyclooxygenases (COX), 607 – lipoxygenases (LOX), 607, 608 – lipid-and water-soluble antioxidants – alpha-lipoic acid, 613 – ascorbic acid, 611 – carotenoids, 612, 613 – coenzyme Q (CoQ), 614, 615 – docosahexaenoic acid (DHA), 616, 617 – glutathione (GSH), 613, 614 – low molecular weight antioxidants (LMWA), 610, 611 – vinyl ether bond, 615, 616 – vitamin E, 612 – lipids, 605–607 – nonenzymatic lipid peroxidation – 4-hydroxy-2-trans-nonenal (4-HNE), 610 – isoprostanes, 608, 609 – malondialdehyde (MDA), 609 – perinatal brain

– adverse environments, 620, 621 – docosahexaenoic acid (DHA), 621–623 – oxygen deprivation, 621 – phospholipid asymmetry, 617–620 – traumatic brain injuries, 605 Oxysterol, 228, 232

P Pain, 475, 480 – inflammatory, 90 – neuropathic pain, 89, 90 Palmitate, 354, 356–358, 366 Palmitic acid, 24, 452, 472 Palmitoylation, 355, 357, 358, 360, 362, 366 N-Palmitoylethanolamine (PEA), 386, 391, 401 Pancreas, 464 Paracetamol, 390 Paraoxon, 477 Parkinson’s disease (PD), 477, 480 – cannabinoids receptors in, 589 – Lewy bodies (LB) in, 584, 588 – Lewy neuritis (LN) in, 584, 588 – mitochondrial function damage in, 587 – oxidation in pathogenesis of, 586 – symptoms, 586, 589 Parturition, 457 Parvocellular neurons, 399 Passive avoidance behavior, 476 Patch-clamp, 465 PC12 cells, 468, 470, 472, 477 PEMT-/-mice phenotype, 232 Pentylenetetrazol, 477 Perikaryon, 466 Perinatal, 476 Perinatal period, 476 Peritoneal dialysates, 457 Peritumoral edema, 540 Peroxisomal a-methylacyl-CoA racemase, 636 Peroxisomal biogenesis, defective peroxisomal biogenesis, 632 Peroxisomal disorders – classification of, 632 – elevated VLCFA in, 637 – lipid metabolism, 637 – molecular aspects, 645, 646 – pathology, 651–654 Peroxisome-proliferator activating receptors (PPARs) – PPAR-a, 76, 82, 85, 88–90

Index Peroxydienyl radicals (LOO), 605, 606 Phenylalanine hydroxylase, 31 Phopshatidylcholine (PC), 231, 232 Phopshatidylethanolamine, 536 Phosphatase and tensin homolog deleted on chromosome 10 (PTEN), 542, 549 Phosphatidate, cell signaling, 290, 296, 298, 299 Phosphatidate phosphatases-1 – cell signaling, 299 – glycerolipid synthesis, 299, 300 Phosphatide, 446, 447, 451, 452, 454, 457, 460, 466–471, 473, 477 Phosphatidic acid (PA), 208, 228, 231, 451, 458, 459, 470 Phosphatidylcholine (PC), 247, 248, 445–482, 536, 548 – in brain mitochondria, 202–205 – disaturated forms, 181 – import in mitochondria, 208 – in ischemia, 212 – in neurodegeneration, 213 – presence in endonucleus, 181 – synthesis, 213 – synthesis and metabolism, 187 Phosphatidylethanol – carbohydrate-deficient transferrin, 509 – gamma-glutamyl transpeptidase, 509 – mean corpuscular volume, 509 – phosphatidic acid, 509 – phospholipase D, 509 Phosphatidylethanolamine (PE), 13, 14, 175, 178, 180, 181, 191, 230, 231, 247, 248, 449–452, 459, 460, 468, 470, 474, 478 – asymmetric distribution, 204 – in brain mitochondria, 202 – importation and synthesis in mitochondria, 207, 208 – in ischemia, 212 Phosphatidylethanolamine-Nmethyltransferase (PEMT1), 449, 451, 460 Phosphatidylglycerol (PG) – in brain mitochondria, 202, 208 – synthesis, 208 Phosphatidylinositol (PI), 203, 204, 208, 212, 228–230, 248, 270, 272–282, 354, 361–366, 452, 459, 468 Phosphatidylinositol-4,5-bisphosphate [PI(4,5)P2], 228–230 – binding to nuclear proteins, 183 – kinase, 183, 184

– precursor of phosphatidylinositol 3,4,5 trisphosphate, 183 – presence in endonucleus, 180 – regulatory function in RNA splicing, 183 Phosphatidylinositol 3-kinase (PI3K), 400 Phosphatidylinositol 4-kinase, 32 Phosphatidylinositol 3,4,5 trisphosphate, 179, 183 Phosphatidyl-N-methylethanolamineN-methyltransferase (PEMT1), 449–451, 460 Phosphatidylserine (PS), 247, 248, 445, 451, 452, 460, 468 – asymmetric distribution, 204 – in brain damage, 50, 51 – in cell signaling and apoptosis, 40, 51 – decarboxylation to phosphatidylethanolamine, 51 – degradation by phospholipases, 42, 51 – import and decarboxylation, 207 – PS synthase 1/2 (PSS1/2), 230 – synthesis by base exchange and its modulation, 51 Phosphatidylserine synthase, 24, 25 Phosphocholine, 23–27, 32, 446, 447, 451, 452, 458, 459, 461, 462, 466, 467, 471, 472, 477 Phosphoethanolamine, 23, 24 3-Phosphoglycerate dehydrogenasedeficiency, L-serine synthesis defect, 719 Phosphoinositide kinases (PKIs), 270, 271, 273 Phosphoinositide phosphatases, 271, 278–282 Phosphoinositides, 228, 230 Phosphoinositide signaling – endonucleus, 181 – nuclear envelope, 181 Phospholipase, 361, 363, 365, 366 – lysophospholipase, 27, 31 – lysoplasmalogenase, 28 – phosphoinositide specific phospholipase C, 181, 183, 184, 191 – phospholipase A2, 180, 181, 184 – phospholipase A (PLA), 25, 27–32 – phospholipase C, 181, 183, 184 – phospholipase C (PLC), 24, 27–30, 32 – phospholipase D, 181, 183, 184

– phospholipase D (PLD), 24, 27–30, 32 – plasmalogen-selective PLA, 28 Phospholipase A1 (PLA1), 451 Phospholipase A2 (PLA2), 209, 212, 451, 452, 478 – generation of lysophosphatidate, 296 – group II secretory PLA2 – in Alzheimer’s disease, 518 – in cerebral ischemia, 523 – in inflammatory responses, 524–527 – neurotoxic action of exogenous sPLA2-IIA, 525 – presynaptic PLA2 neurotoxin, 525 – properties of, 525 – sPLA2 receptors, 525 – group IV cytosolic PLA2 – in Alzheimer’s disease, 518, 519 – in cerebral ischemia, 523 – cPLA2 knockout mice, 518 – link to NADPH oxidase, 521 – oxidative signaling pathways in astrocytes, 521 – oxidative signaling pathways in neurons, 521 – properties of, 520, 521, 523 Phospholipase A1, C, D (PL A1/C/D), 388, 391, 392, 394 Phospholipase C (PLC), 228, 451, 452, 459, 465 Phospholipase D (PLD), 451, 452, 459, 476 Phospholipid association with RNAcontaining structures, 185 Phospholipid asymmetry – cardiolipin (CL), 620 – diacylglycerol kinase (DGK), 619 – flippase, 618 – phosphatidylethanolaminebinding protein (PEBP), 619 – phospholipid scramblase 1 (PLSCR1), 619 – polar head groups (PHG), 617 – polyunsaturated fatty acids (PUFA), 618, 619 Phospholipids, 356, 361, 362, 365, 366, 446, 452–454, 458, 464, 466–468, 470, 474, 478, 481 – N-acylphosphatidylethanolamines, 78 – diacylglycerol, 506 – phosphatidyl-choline (PC), 506, 508

753

754

Index – phosphatidyl-ethanolamine (PE), 506, 508 – phosphatidyl-inositol (PI), 505 – phosphatidyl-serine (PS), 505 – phospholipase C, 78 Phosphorylation, 23, 24, 28, 31, 33, 461, 462, 464, 466, 467, 471–474 Phosphosphingolipid biosynthesis, 243, 246 Phosphotransferases – choline, 23–25 – ethanolamine, 23–25 Phototransduction, 30 PI3K-dependent process, 464 PI3-kinases – regulation, function and enzymatic activity of type I, II and III PI-3 kinases, 273, 274 PI4-kinases – regulation, function, subcellular localization and enzymatic activity of type I, II and III PI-4 kinases, 274–276 PI3-phosphatases – regulation, function and enzymatic activity of PTEN, MTM and MTM related proteins, 278, 279 PI4-phosphatases – regulation, function and enzymatic activity of PI4phosphatases type I, II and of Sac1, 279, 280 PI5-phosphatases – systematic description of type I, II, III, IV PI5-phosphatases, 277 PIP5KI-gamma, generation of PI (4,5)P2 – Synaptojanin-1, 230 – Synaptotagmin I, 230 PIP-kinases – regulation and function of PI4P5Ks, PI5P4Ks and PI3P5K, 273–278 Placenta, 457, 464, 480 PLA2 inhibitors, list of, 527 Plasma, 452, 464, 470 Plasmalogen biosynthesis, 644, 645 Plasmalogens, 25, 27, 32, 224, 228, 231, 506 Plasma membrane, 452, 464, 470, 673–675, 681, 686 Plasticity-related genes (PRG) – during brain damage, 303 – during brain development, 303 – cell signaling, 302, 303

– expression in central nervous system, 302, 303 – response to lysophosphatidate, 302, 303 – sphingosine 1-phosphate (S1P), 294, 295, 297, 298 Platelet-activating factor (PAF), 24, 25, 28, 29, 31, 33, 446, 452 – biosynthetic pathways of, 317, 321, 324, 326, 331, 332, 338 – in brain ischemia, 334, 335 – degradation pathways of, 313, 317, 320–323, 326, 332, 338 – in gene expression, 330, 331 – in HIV infection, 337–338 – in long-term potentiation (LTP), 327–329, 335, 338 – in Miller–Dieker lissencephaly, 337 – in neurodegenerative diseases, 331, 335–337 – in oxidative stress, 323, 335, 336 – PAF receptor agonists, 326, 327 – PAF receptor antagonists, 325–328, 331, 332, 335, 336, 338 – in proliferation and differentiation of neural cells, 329, 330 – receptors of, 325, 326 – in synaptic transmission, 327 Platelet closure time, 477, 480 Platelet count, 477, 480 Platelet-derived growth factor (PDGF), 546 Polyamine, 462 Polyunsaturated fatty acids (PUFA), 451, 467 – arachidonic acid (AA), 519, 536, 537 – cis-paranaric acid, 537 – docosahexaenoic acid (DHA), 519, 536, 537 – eicosapentaenoic acid, 536 – g-linolenic acid, 536 – linoleic acid, 537, 540 Postnatal, 457, 476 Postsynaptic, 460, 465, 467 Potassium cyanide, 478 Potency, 465 Precursor, 446, 451, 457, 460, 466, 468, 470, 471, 473, 476 Pregnancy, 457, 471 Pregnant rats, 476 Prenatal, 476, 477 Prenatal choline supplementation, 476, 477

Prenyl, 354, 357–360 Presynaptic, 461, 464, 467, 475 Pre-synaptic membrane protein – Clathrin/AP2, 230 – M/UNC-13, binding to DAG, 228 Prion, 356, 361, 363 Progesterone (P), 390, 398 Prolactin (PRL), 386, 458, 475 Proliferator activated receptor (PPARg), 540 Promoter, 390, 398 Prosaposin, 680, 691 Prostaglandins, 31, 228, 230, 231 Protein kinase A/B, 386 Protein kinase-C, 486, 487, 489, 496, 497 Proteome, 358, 361 PSD-95, 468 Psychiatric diseases – autism, 422, 425, 426, 429 – bipolar disorder, 423 – dementia – major depression, 424 – schizophrenia, 425, 429 – depression – post-natal depression, 423 – seasonal affective disorder (SAD), 423 – drug addiction, 422 – dyslexia, 422 – hyperactive children, 422 – mood – aggression, 421 – suicide, 423 – pain, 421 – stress, 421, 424, 427 Psychosine, 682, 691 PtdCho. See Phosphatidylcholine PtdEtn. See Phosphatidylethanolamine PtdIns. See Phosphatidylinositol PtdSer. See Phosphatidylserine Purkinje neurons, 477 P2Y receptors, 470, 471 Pyridoxine, 457 Pyrimidine, 467, 471, 473, 474

Q Quantitative structure–activityrelationship (QSAR), 390 Quaternary nitrogen atom, 458 [3H]-Quinuclidinyl benzilate binding, 465

R Rabbit, 462 Radial-arm maze, 476 Radiation resistance, 540

Index Rafts, 673 Ras, 356, 358–360 Rat, 451, 454, 456, 457, 459, 462–465, 467, 471–476, 478–481 Rat brain endothelium (RBE), 394 Rate-limiting, 461, 462, 467, 468 Reactive oxygen species (ROS), 211, 537, 604, 606 Reacylation/deacylation cycle, 22, 27, 28 Receptors, 459–461, 465, 466, 470, 471, 474–478 – a-amino-3-hydroxyl-5-methyl-4isoxazole-propionate (AMPA), 357 – cannabinoid, 29 – cytokine, 27 – dopamine, 27 – GABA, 357 – GABAA receptor, 63, 67 – glutamate, 27 – glycine receptor, 67 – growth factor, 27 – influence of neuroactive steroids on, 67 – intracellular steroid receptor, 67 – kainate receptor, 67 – N-methyl-D-aspartate (NMDA) receptor, 60, 67 – muscarinic receptor, 67 – nicotinic receptor, 67 – noradrenergic, 31 – platelet-activating factor (PAF), 25, 33 – serotonergic, 31 – serotone (5-HT3) receptor, 67 – sigma receptor, 67 – steroid receptor, 67 Recombinant tPA, 478 Regeneration, 28, 29 Renal transplantation, 457 Residual activity, 673, 683, 686, 687 Resolvins, 31 Retrograde messenger, 33 Retrograde signaling, 397 Retrograde transport, 26 Rhizomelic chondrodysplasia punctata (RCDP) – types 1, 2 and 3, 633, 634, 636, 637 RNA – interaction with phospholipids, 179 – spliceosomes and lipid interaction, 180 Ro09-0198, 32, 33 Rodent, 468, 472, 473, 477

S S-adenosylmethionine, 24, 445, 447, 448, 450 Salmonella minnesota mutant R595, 126 Sandhoff disease – globoside storage, 723 – GM2-storage, 723 – HexB deficiency, 723, 724 Sap-A, 678–680, 691 Sap-B, 676, 678–680, 689 Sap-C, 676, 678, 680, 681, 690, 691 Sap-D, 678, 680, 691 Saposins, 678 Sap-precursor, 678–680, 682, 691 Saturated fatty acid, 454, 470 Scopolamine, 467, 479 SCPx deficiency, 650 Seafood, 471 Second messengers, 23, 28–31, 34, 458 Secretase, 356, 362, 363 Secretion, 30 Seizure-promoting drug, 477 Sensory organs – electroretinogram (ERG), 415, 419 – hearing, 420 – olfaction, 420 – photoreceptor, 419 – retina, 415, 419 – taste, 420 – vision, 417, 419, 420 – visual acuity, 417, 419, 420 Serine glycerophospholipids, 22, 24–26, 32, 33 Serine palmitoyl-CoA transferase deficiency – sphingosine synthesis defect, 718 Serine-palmitoyltransferase (SPT), 242, 400 Serotonin, 460, 466 Serum, 455, 457, 458, 477 Short-term memory, 476 Sialyl(a2!1)sphingosine, 120 Signal transducer and activator of transcription (STAT), 390 Signal transduction, 22, 28, 30, 31 Single peroxisomal b-oxidation proteins, 634–637 Small intestine, 464 Sodium-calcium exchanger – complex with GM1 ganglioside, 177 – expression on inner nuclear membrane, 177 – role in nuclear calcium homeostasis, 179 Solid-state NMR, 396

Spatial memory, 476 Spermidine, 459, 462 Spermine, 462 Sphingolipid degradation, 676–681 Sphingolipid metabolism – ceramide, 374, 375 – de novo pathway, 374, 376, 377 – salvage/recycling pathway, 374, 376, 378 – sphingomyelinase pathways, 374, 376, 377 – sphingosine kinase (SK) pathway, 374 Sphingolipidoses, 681–693 Sphingolipids, 225, 228, 233, 234, 362 – ceramide, 187, 541, 542, 544, 545, 548 – galactosylcerebroside, 545 – gangliosides, 187, 190, 544–548 – glycosphingolipids, 187–190 – paragloboside, 545 – sphingomyelin, 184–186 – sphingosine-1-phosphate – receptors for, 542, 543, 546, 547 Sphingomyelin (SM), 224, 228, 233, 234, 445–447, 452, 456, 458, 466–468, 471, 479 – bridge in double stranded RNA, 180, 186 – endonucleus, 185 – nuclear envelope, 185 – sphingosine 1-phopsphate (S1P), 597 Sphingomyelinase (SMase), 542 – Acid SMases, 233 – involvement in apoptosis, 185 – Neutral SMase2, 233 Sphingomyelinase deficiency – acidic/neutral sphingomyelinases, 731–733 – Niemann-Pick disease (NPD), 731 – sphingomyelin storage, 731 Sphingomyelinase pathways, 374, 376, 377 Sphingomyelins, 22, 26 Sphingomyelin synthase, forward and reverse reactions, 185 Sphingosine (Sph), 379 Sphingosine kinase (SphK), 542–544 Sphingosine kinase deficiency – signal transduction, 717 – sphingosine-1-phosphate receptor, 717 Sphingosine kinase (SK) pathway, 374 Sphingosine-1-phosphate (S1P), 186, 187, 379

755

756

Index Spleen, 464, 473 Spliceosomes, 178, 180, 191 Statin, 358 Stearate, 354 Stearic acid, 24, 452 N-Stearoylethanolamine (SEA), 386 Steroid hormones – aldosterone, 65, 66 – androstenedione, 63, 64 – biosynthesis of, 60, 63, 64 – corticosterone, 65, 66 – cortisol, 65 – deactivation of, 63, 64 – dehydroepiandrosterone (DHEA), 64, 65, 67, 68 – dehydroepiandrosterone sulfate (DHEAS), 64, 65, 67, 68 – estradiol, 63, 64 – estrone, 63, 64 – glucocorticoids, 60, 65 – mineralocorticoids, 60 – progesterone, 61, 62, 65, 66, 68 – testosterone, 62–64 Steroidogenic enzymes – biochemical characteristics, 61 – brain, presence in, 61, 63–66 – catalytic reactions catalyzed by, 61, 64 – cofactor requirement patterns, 64 – cytochrome P450 aldosterone synthetase (P-450aldo), 65, 66 – cytochrome P450 aromatase, 63 – cytochrome P450 cholesterol side-chain cleavage (P450SCC), 61 – discovery, 60 – enzyme activity in the brain (cerebral cortex and subcortical white matter), 62, 65 – 7a-hydroxylase (CYP7B1), 64, 65 – 11b-hydroxylase (cytochrome P45011b), 65 – 21-hydroxylase (cytochrome P450c21), 65 – 3b-hydroxysteroid dehydrogenase (3b-HSD), 61–63, 65 – 17b-hydroxysteroid dehydrogenase (17b-HSD), 63–65 – hydroxysteroid sulfotransferase (SULT2), 64 – isoforms, 62, 63 – kinetic properties, 64 – mRNA expression in the brain (cerebral cortex and subcortical white matter), 61–65

– organic anion transporter polypeptides (OATP), 64 – P450c17 (17a-Hydroxylase/C1720-lyase), 64, 65 – 5a-reductase, 61, 62, 66 – sex differences, 61, 64 – steroid sulfatase (STS), 64–66 – substrate specificity, 63–65 – tissue distribution, 61 Steroids, 398–400 Sterol, 362 Storage diseases – activator protein defects, 725 – sphingolipid degradation defects, 721 Striatum, 462, 463, 479 Stroke, 477, 478, 480 Strychnine, 477 Study – DATATOP, 587 – ELLDOPA, 586 Subcutaneous, 475, 481 Substantia nigra, 584, 585, 587 Substrate-saturation, 461, 462, 470, 471 Sulfated ceramide (Sulfatides), arylsulfatase A, 233 Supplementation, 468, 476, 477, 479 Surface charge, 26 Surgical stress, 458 Survival rate, 474 Sympathoadrenal system, 476, 481 Synapse, 458, 460, 465, 467, 480 Synapsin-1, 468 Synaptic transmission, 32 Synaptosome, 451, 463, 464 Syntaxin-3, 468, 470, 471 a-Synuclein – effect of lipid vesicles on, 588 – human aS gene of (Snca), 588 – interaction of lipids with, 588 – mutations of human gene of, 588 – role in neurodegeneration of, 588

T Tau, hyperphosphorylation by LPA, 300 Tay–Sachs disease – GM2-storage, 724, 725 – HexA deficiency, 722, 723 Testis, 464 Tetanus toxin, 118, 120 Tetramethylrhodamine (TMR), 115 Therapeutic exploitation, 395 Therapy – cell-mediated therapy (CMT), 692 – enzyme-enhancement, 692, 693

– enzyme replacement therapy (ERT), 687, 692 – gene, 692, 693 – substrate deprivation, 692, 693 Thermal pain, 475 Thromboxane, 538–541 Thromboxanes, 31 Thy-1, 361, 363, 364, 366 Trancription factor, 390 Transamidase, 361 Transferases – farnesyl, 359 – geranylgeranyl, 359, 360 – glycosyl transferase, 31 – palmitoyl, 357 – prenyl, 359 – sialyl transferase, 31 Transfer proteins, 25, 34 Transient receptor potential channel vanilloid receptor subunit 1 (TRPV1), 394–396, 400 Transport – anterograde, 26 – retrograde, 26 Transport protein, 459–464, 470, 473 Traumatic brain injury, 477, 480 Triglyceride (TG), 505 Trimethylamine, 456 Tryptophan, 445, 466 Tumor necrosis factor-a (TNF-a), 32, 477

U United States Department of Agriculture (USDA), 449 Upper (daily) limit (UL), 447 Uptake, 446, 458–461, 463, 464, 467, 473, 481 Uridine, 451, 466–468, 470, 471, 473, 474, 477 Uridine-cytidine kinase (UCK), 445, 473, 474 Uridine diphosphate, 445 Uridine monophosphate, 468 Uridine triphosphate, 445, 470 Urinary catecholamine output, 475 Urine, 456 Urokinase, 478

V Vasopressin, 475, 480 Verbal memory, 479 Very long chain fatty acids (VLCFA) – b-oxidation, 637–640 Vesicles – intraendosomal, 675, 676 – intralysosomal, 675, 676, 681

Index Virodhamine. See O-Arachidonoylethanolamine Vitamin B12, 448, 452, 457 Voluntary muscle, 465

X

W

Z

Weight loss, 477 Withdrawal, 477, 480

Zellweger spectrum disorders (ZSD)

X-chromosomal-linked inherited disease, 686

– clinical features, 632, 633 – therapy of, 633, 634 Zinc metallohydrolases, 388

757