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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

543210

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 over expressing 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

viii

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: jean [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] tokyo.ac.jp

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: Birgit.Stoffel [email protected] bonn.de 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, phos phatidylcholine; 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

1

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 phos pholipids 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 fragmenta tion (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 identifi cation 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 combina tion 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 informa tion. 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, compre hensively 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 phos pholipids. 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 phospho lipids, 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

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‐a k‐2‐acy ,18:0‐14:0) (1‐a k‐2‐acy ,16:0‐16:0) (1‐acy ‐2‐acy ,16:1‐16:1) (1‐acy ‐2‐acy ,14:0‐18:1) (1‐acy ‐2‐acy ,16:0‐16:1) (1‐acy ‐2‐acy ,16:0‐16:0) (1‐a k‐2‐acy ,16:0‐18:1) (1‐a k‐2‐acy ,18:1‐16:0) (1‐a k‐2‐acy ,18:0‐16:0) (1‐acy ‐2‐acy ,14:0‐20:4) (1‐acy ‐2‐acy ,16:1‐18:2) (1‐acy ‐2‐acy ,16:0‐18:3) (1‐acy ‐2‐acy ,16:0‐18:2) (1‐acy ‐2‐acy ,16:0‐18:1) (1‐acy ‐2‐acy ,16:0‐18:0) (1‐a k‐2‐acy ,16:1‐20:4) (1‐a k‐2‐acy ,16:0‐20:4) (1‐a k‐2‐acy ,18:0‐18:3) (1‐a k‐2‐acy ,18:1‐18:2) (1‐a k‐2‐acy ,18:0‐18:2) (1‐a k‐2‐acy ,18:1‐18:1) (1‐a k‐2‐acy ,18:0‐18:1) (1‐a k‐2‐acy ,18:0‐18:0) (1‐acy ‐2‐acy ,18:2‐18:2) (1‐acy ‐2‐acy ,16:0‐20:4) (1‐acy ‐2‐acy ,18:1‐18:2) (1‐acy ‐2‐acy ,16:0‐20:3) (1‐acy ‐2‐acy ,18:0‐18:3) 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‐acy ‐2‐acy ,18:0‐18:2) (1‐acy ‐2‐acy ,16:0‐20:2) (1‐acy ‐2‐acy ,18:1‐18:1) (1‐acy ‐2‐acy ,16:0‐20:1) (1‐acy ‐2‐acy ,18:0‐18:1) (1‐a k‐2‐acy ,18:0‐20:4) (1‐acy ‐2‐acy ,18:0‐20:7) (1‐acy ‐2‐acy ,16:0‐22:6) (1‐acy ‐2‐acy ,18:2‐20:4) (1‐acy ‐2‐acy ,18:1‐20:4) (1‐acy ‐2‐acy ,16:0‐22:5) (1‐acy ‐2‐acy ,18:0‐20:5) (1‐acy ‐2‐acy ,18:0‐20:4) (1‐acy ‐2‐acy ,16:0‐22:3) (1‐acy ‐2‐acy ,18:1‐20:2) (1‐acy ‐2‐acy ,18:0‐20:3) (1‐acy ‐2‐acy ,18:0‐20:2) (1‐acy ‐2‐acy ,18:0‐20:1) (1‐acy ‐2‐acy ,20:5‐20:5) (1‐acy ‐2‐acy ,18:1‐22:5) (1‐acy ‐2‐acy ,18:0‐22:6) (1‐acy ‐2‐acy ,18:0‐22:5) (1‐acy ‐2‐acy ,18:0‐22:4) (1‐acy ‐2‐acy ,18:0‐22:3)

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Source: Houjou et a . (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

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

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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 collabo ration 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 weight related 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 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

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

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 compensa tion 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 impor tant. (> 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.

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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 asymmetri cally 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 reac tions 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. Glycer ophospholipids 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, cytidinedi phospho; CTP, cytidine triphosphate; DAG, 1,2 sn diacylglycerols; DHA, docosahexaenoic acid; ER, endo plasmic 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 glyceropho spholipids 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 glyceropho spholipids 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 glyceropho spholipids 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 struc tural 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, well organized, and highly specialized structures involved in receiving, processing, transporting, and transmit ting information from one cell to another. Two unresolved fundamental questions related to choline and ethanolamine glycerophospholipid metab olism 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 mem branes 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 ethanol amine 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 cytidylyl transferase 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); PAF synthesizing 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 mole cule, 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 glyceropho spholipids contain palmitic acid or stearic acid at the sn 1 position with long chain polyunsaturated fatty

Choline and ethanolamine glycerophospholipids

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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 glyceropho spholipids 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 acyldihy droxyacetone 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 O alkyldihydroxyacetone 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 phosphocho line 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 synthe sized 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 subcellu lar 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 glyceropho spholipid 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 approxi mately 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.

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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 glycerophospholipid synthesizing 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 glyceropho spholipids 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.

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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 acyltransfer ase (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 glyceropho spholipids 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

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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; phospha tidic acid (PtdH); protein kinase C (PKC); arachidonic acid (AA); docosahexaenoic acid (DHA), and platelet activating 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 Glyceropho spholipids 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 docosahex aenoic 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). Iso forms of PLD are implicated in membrane trafficking, cell proliferation, mitogenesis, oncogenesis, inflam mation, 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

2

. 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

A2

D

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 physiologi cal 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, 17S resolvins, 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 acyl transferase, 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 glycer ophosphocholine is either metabolized to free choline by alkaline phosphatase or used by cytidylyltransfer ase 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) (Schutze 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 conden sation, 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 degenera tion 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 metallo proteinase 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 choli neglycerophospholipids 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 ex change, methylation, and decarboxylation). These reactions and activities of phospholipases A2, C, and D may be responsible for the turnover, compositional maintenance, and rearrangements of glyceropho spholipids 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.

References Atsumi G, Tajima M, Hadano A, Nakatani Y, Murakami M, et al. 1998. Fas-induced arachidonic acid release is mediated by Ca2þ -independent phospholipase A2 but not cytosolic phospholipase A2 which undergoes proteolytic inactivation. J Biol Chem 273: 13870-13877. Attucci S, Albani-Torregrossa S, Moroni F, PellegriniGiampietro DE. 2001. Metabotropic glutamate receptors stimulate phospholipase D via different pathways in the adult and neonate rat hippocampus. Neurochem Res 26: 1151-1155. Axelrad TW, Deo DD, Ottino P, Van Kirk J, Bazan NG, et al. 2004. Platelet-activating factor (PAF) induces activation of matrix metalloproteinase 2 activity and vascular endothelial cell invasion and migration. FASEB J 18: 470-492. Banchio C, Schang LM, Vance DE. 2004. Phosphorylation of Sp1 by cyclin-dependent kinase 2 modulates the role of Sp1 in CTP: Phosphocholine cytidylyltransferase alpha regulation during the S phase of the cell cycle. J Biol Chem 279: 40220-40226. Banno Y. 2002. Regulation and possible role of mammalian phospholipase D in cellular functions. J Biochem (Tokyo) 131: 301-306. Bayo´n Y, Herna´ndez M, Alonso A, Nunez L, Garcia-Sancho J, et al. 1997. Cytosolic phospholipase A2 is coupled to muscarinic receptors in the human astrocytoma cell line

1321N1: Characterization of the transducing mechanism. Biochem J 323: 281-287. Bazan NG. 2003. Synaptic lipid signaling: Significance of polyunsaturated fatty acids and platelet-activating factor. J Lipid Res 44: 2221-2233. Bazan NG. 2005. Lipid signaling in neural plasticity, brain repair, and neuroprotection. Mol Neurobiol 32: 89-103. Birner R, Daum G. 2003. Biogenesis and cellular dynamics of aminoglycerophospholipids. International Review of Cytology - A Survey of Cell Biology, Vol. 225. Jeon KW, editor. San Diego: Academic Press Inc; pp. 273-323. Bleijerveld OB, Klein W, Vaandrager AB, Helms JB, Houweling M. 2004. Control of the CDPethanolamine pathway in mammalian cells: Effect of CTP: Phosphoethanolamine cytidylyltransferase overexpression and the amount of intracellular diacylglycerol. Biochem J 379: 711-719. Carman GM, Kersting MC. 2004. Phospholipid synthesis in yeast: Regulation by phosphorylation. Biochem Cell Biol 82: 62-70. Carter JM, Waite KA, Campenot RB, Vance JE, Vance DE. 2003. Enhanced expression and activation of CTP: Phosphocholine cytidylyltransferase b2 during neurite outgrowth. J Biol Chem 278: 44988-44994. Chauhan V, Sheikh AM, Chauhan A, Spivack WD, Fenko MD, et al. 2005. Regulation of high molecular weight bovine

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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

3 3.1 3.2 3.3 3.4 3.5

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

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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, indepen dently 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 lyso somes, 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 methodol ogies 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 PtdSer synthesizing 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 co cultures 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 mecha nism 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 particu larly difficult for the nervous tissue because of the marked functional differences of brain regions, their cellular heterogeneity and developmental modifications.

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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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Brain phosphatidylserine: metabolism and functions

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, 20:4n 6 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 well known 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 incor poration 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  105, 4.33  105 and 6.75  104, 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 ethanol amine incorporation and vice versa suggested that the enzyme catalyzes the incorporation of both ethanol amine 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 expres sion 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).

Brain phosphatidylserine: metabolism and functions

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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 ethanol amine 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 incorpora tion 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, acetyl choline, 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 oxotremor ine 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

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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 3 phosphoethanolamine 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. Further more, 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 phos phoglycerides 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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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 2 acyl 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.

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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 inhibi tion 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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Putney JW Jr. 1993. Excitement about calcium signaling in inexcitable cells. Science 262: 676-678. Raghavan S, Rhoads D, Kanfer J. 1972. In vitro incorporation of (14C)serine, (14C)ethanolamine, and (14C)choline into phospholipids of neuronal and glial-enriched fractions from rat brain by base exchange. J Biol Chem 247: 7153-7156. Rhodes PG, Hu ZY, Sun GY. 1993. Effects of chlorpromazine on phosphatidylserine biosynthesis in rat pup brain exposed to ethanol in utero. Neurochem Int 22: 75-80. Rodriguez FD, Alling C, Gustavsson L. 1996. Ethanol potentiates the uptake of [14C]serine into phosphatidylserine by base-exchange reaction in NG 108-15 cells. Neurochem Res 21: 305-311. Ross BM, Mamalias N, Moszczynska A, Rajput AH, Kish SJ. 2001. Elevated activity of phospholipid biosynthetic enzymes in substantia nigra of patients with Parkinson’s disease. Neuroscience 102: 899-904. Rossi M, Corazzi L, Fratto G, Arienti G. 1990. The effect of membrane lipid molecular species on rat brain baseexchange reactions: An appraisal of phosphatidylserine and of polyunsaturated phosphatidylcholine. Farmaco 45: 1067-1073. Rouser G, Yamamoto A, Kritchevsky G. 1971. Cellular membranes. Structure and regulation of lipid class composition species differences, changes with age, and variations in some pathological states. Arch Intern Med 127: 1105-1121. Ryu SB, Palta JP. 2000. Specific inhibition of rat brain phospholipase D by lysophospholipids. J Lipid Res 41: 940-944. Sahu SK, Gummadi SN, Manoj N, Aradhyam GK. 2007. Phospholipid scramblases: An overview. Arch Biochem Biophys 462: 103-114. Saito M, Bourque E, Kanfer J. 1975. Studies on base-exchange reactions of phospholipids in rat brain particles and a ‘‘solubilized’’ system. Arch Biochem Biophys 169: 304-317. Sakane F, Yamada K, Imai S, Kanoh H. 1991. Porcine 80-kDa diacylglycerol kinase is a calcium-binding and calcium/phospholipid-dependent enzyme and undergoes calcium-dependent translocation. J Biol Chem 266: 7096-7100. Salem N Jr, Litman B, Kim HY, Gawrisch K. 2001. Mechanism of action of docosahexaenoic acid in the nervous system. Lipids 36: 945-959. Sato T, Aoki J, Nagai Y, Dohmae N, Takio K, et al. 1997. Serine phospholipid-specific phospholipase A that is secreted from activated platelets – a new member of the lipase family. J Biol Chem 272: 2192-2198. Scorrano L, Oakes SA, Opferman JT, Cheng EH, Sorcinelli MD, et al. 2003. BAX and BAK regulation of endoplasmic reticulum Ca2+: A control point for apoptosis. Science 300: 135-139.

Singh IN, McCartney DG, Sorrentino G, Massarelli R, Kanfer JN. 1992a. Phorbol esters modulate phospholipid metabolism in a human cholinergic cell line, LA-N-2: A possible role for the base exchange enzymes. J Neurosci Res 32: 583-592. Singh IN, Sorrentino G, Massarelli R, Kanfer JN. 1992b. Oleoylamine and sphingosine stimulation of phosphatidylserine synthesis by LA-N-2 cells is protein kinase C independent. FEBS Lett 296: 166-168. Smrz D, Lebduska P, Draberova L, Korb J, Draber P. 2008. Engagement of phospholipid scramblase 1 in activated cells; implication for phosphatidylserine externalization and exocytosis. J Biol Chem 283(16): 10904-10918. Stone SJ, Cui Z, Vance JE. 1998. Cloning and expression of mouse liver phosphatidylserine synthase-1 cDNA. Overexpression in rat hepatoma cells inhibits the CDPethanolamine pathway for phosphatidylethanolamine biosynthesis. J Biol Chem 273: 7293-7302. Stone SJ, Vance JE. 1999. Cloning and expression of murine liver phosphatidylserine synthase (PSS)-2: Differential regulation of phospholipid metabolism by PSS1 and PSS2. Biochem J 342(Pt 1): 57-64. Stone SJ, Vance JE. 2000. Phosphatidylserine synthase-1 and 2 are localized to mitochondria-associated membranes. J Biol Chem 275: 34534-34540. Sun GY, Foudin LL. 1985. Phospholipid composition and metabolism in the developing and aging nervous system. Eichberg J, editor. New York: Wiley. Sun GY, Huang HM, Lee DZ, Sun AY. 1984. Increased acidic phospholipids in rat brain membranes after chronic ethanol administration. Life Sci 35: 2127-2133. Sun GY, Samorajski T. 1972. Age changes in the lipid composition of whole homogenates and isolated myelin fractions of mouse brain. J Gerontol 27: 10-17. Sun GY, Sun AY. 1983. Chronic ethanol administration induced an increase in phosphatidylserine in guinea pig synaptic plasma membranes. Biochem Biophys Res Commun 113: 262-268. Sun GY, Sun AY. 1985. Ethanol and membrane lipids. Alcohol Clin Exp Res 9: 164-180. Suzuki TT, Kanfer JN. 1985. Purification and properties of an ethanolamine-serine base exchange enzyme of rat brain microsomes. J Biol Chem 260: 1394-1399. Szalai G, Krishnamurthy R, Hajnoczky G. 1999. Apoptosis driven by IP(3)-linked mitochondrial calcium signals. EMBO J 18: 6349-6361. Tait JF. 2008. Imaging of apoptosis. J Nucl Med 49: 1573-1576. Tijburg LB, Geelen MJ, Van Golde LM. 1989. Biosynthesis of phosphatidylethanolamine via the CDP-ethanolamine route is an important pathway in isolated rat hepatocytes. Biochem Biophys Res Commun 160: 1275-1280.

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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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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, 3a hydroxysteroid 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), nico tinic, 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 sulfotransfer ase; 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 transcrip tionally 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.

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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 medi cally 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 metabo lized 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 (Thig pen 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 (Stoffel Wagner 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 17b HSD 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 immunocyto chemistry (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 syn thetase (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 mineralo corticoids (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 expres sion 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).

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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, dihydrotestoster one; 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

Metabolism and enzymology of cholesterol and steroids

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Steroid hormone effects on the brain have typically been associated with gene regulation via intracellu lar 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 ligand gated 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 GABA ergic 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 metabo lism 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 concentra tions 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 concentra tions 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

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

2 2.1 2.2 2.3 2.4

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 bind ing 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 peroxisome proliferator 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 docosa hexaenoyl 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 ethano lamide, 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 degra dative 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

Anandamide and other acylethanolamides

5

. 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 glycerol phospho 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, arachidonoylethanola mide (anandamide); FAAH, fatty acid amide hydrolase; FAAH 2, fatty acid amide hydrolase 2; NAAA, N acyletha nolamine 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

Anandamide and other acylethanolamides

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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 immunor eactive, 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 preferen tially 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 pharmaco logically 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 dioleyl glycerol 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 eicosatrienoyl ethanolamide

7Z,10Z,13Z,16Z docosatetraenoyl ethanolamide

Di homo g linolenoyl ethanolamide

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 acylethano lamides 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 phospholi pase 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 mechan isms 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 eicosatrienoyl ethanolamide, 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 CB1 mediated 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 endo cannabinoids 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 endocanna binoids 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 hypothal amus, 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 partici pates 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 neuroin flammatory 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 immedi ately 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 neuro logical 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 strengthen ing 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 antinocicep tive 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 neuropathologi cal 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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Chemistry, Tissue and Cellular Distribution, and Developmental Profiles of Neural Sphingolipids*

G. Tettamanti . L. Anastasia

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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 aminoal cohol (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 glyco sphingolipids, 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” detec tion, 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 develop mental 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 sphin gosine 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 1,3 dihydroxy 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 (Cer 1 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 (sphingosylphospho choline, 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 glucuro nic 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). Glyco sphingolipids 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 amino 3,5 dideoxy D glycero D galacto non 2 ulopyranosonic acid, 5 N acetylneuraminic acid, Neu 5Ac), fol lowed 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], ganglio triose[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 Chol 1) 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 galactosyl ceramide (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 Sa sman (1965) Kuhn and Wiegandt (1963)

Ghidoni et a . (1976)

Ariga et a . (1987a)

Kuhn and Wiegandt (1963)

Ghidoni et a . (1976)

Ando and Yu (1977)

GM1

Fuc-GM1

Fuc-Ga -GM1

GD1a

GD1a(Neu5Gc)

GT1a

Ledeen et a . (1973)

Yu and qba (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 a . (1995)

Sonnino et a . (1978)

GD1b

9-O-Ac-GD1b

Fuc-GD1b

Ariga et a . (1987a)

Svennerho m et a . (1973)

Kuhn and Wiegandt (1963)

Fuc-Ga -GD1b

Ga NAc-GD1b

GT1b

Riboni et a . (1986)

Kuhn and Wiegandt (1964)

GD2

Lactone-GD1b

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

b-series GD3 9-O-Ac-GD3

continued

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

6 105

Ghidoni et a . (1980)

Kuhn and Wiegandt (1963)

Chigorno et a . (1982b)

Yu and Ando (1980)

Waki et a . (1993b)

Yu and Ando (1980)

Waki et a . (1993a)

shizuka 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 a . (1990)

Ando et a . (1992)

Ando et a . (1992)

Mo in et a . (1987)

Mo in et a . (1987)

GD1a

GT1a

GQ1a

Others 3’isoLM1

6’LM1

continued

Ro¨sner et a . (1985)

GP1c

a-series

shizuka and Wiegandt (1972)

GQ1c

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

6 107

Chou et a . (1996)

Fukushima et a . (1984)

9-O-Ac-3’LD1

SLex-gang iosidea

a

SLex: sia y Lewisx This compound was found to occur in endothe ia ce s, inc uding brain microvesse s b The presence of this compound was observed in PC 2 pheochromucytoma ce s (neurotumora ce s)

 O3S-6’,6-bis-a SLexgang iosideb

O3S-6-SLexgang iosidea



O3S-6’-SLexgang iosidea

Ga ustian et a . (1997)

Ga ustian et a . (1997)

Ga ustian et a . (1997)

Chou et a . (1996)

3’LD1

6



Chou et a . (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

6

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 acetal linkage 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 (endocerami dases) (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 glycosphin golipids, 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 dihydrosphingo sine (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 com pounds 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 5,6 dicyanobenzoquinone (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 sphingo sine, 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 lyso sphingomyelin is treated with a bacterial phospholipase D, with the formation of labeled sphinganine 1 phosphate (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 phosphorylcho line 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 glyco sphingolipids) 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 dipyrro methene 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, BOD 12 O3S Gal Cer, DMB 5 O3S Gal Cer, BOD 12 Lac Cer, DMB 5 Lac Cer, BOD 12 GalNAc Gal Glc Cer, 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 suc cessfully used as a fluorescent probe for gangliosides GM1, GM2, GM3, asialo GM1, asialo GM2, lactosyl ceramide, 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 laser induced 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. Accord ing 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 sialoglycoconju gates 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 N acetylation 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 bioti nyl 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 sphingo lipids, 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 photo labeling 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 (glycosphingoli pids). 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 toluene sulfonyloxyethyl) 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 compart ment 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 con solidated 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 develop ments 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 hexosa minidase 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 (hydropho bic interactions) (Hakomori, 1981; Hakomori and Igarashi, 1993). Other important features of sphingo lipids 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 oligosaccha ride 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 3 phospho 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 neo glycolipids 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 adaman tyl 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 glycosphingo lipids 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 cycloexane dial, 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 sulpha tides 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 proce dures 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 accom plished colorimetrically by spraying appropriate reagents [resorcinol HCl (Svennerholm, 1957) or p dimethylaminobenzaldehyde (Chigorno et al., 1982a) reagent for gangliosides; sulfuric acid for phos pholipids 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 immuno staining 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; Muthing, 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 compo nents 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 sensitivi ty 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 saccha ride 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

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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 ESI QTOF) 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 dissoci ation mass spectrometry (SORI CID MS MS), by which sialylated, sulfated and glucuronylated glyco sphingolipids 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 (tridimension al) structure of the molecules and the dynamics around a flexible linkage (presence of different con formers). 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 (Kohler and Milstein, 1975), the immunization principle to adsorb glycolipids (or bacterial lipopolysac charides) 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 ganglio sides of the b series (carrying a di sialosyl residue on the inner galactose moiety), namely GD3, 9 O Ac GD3, 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 concentra tion 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 glycosphin golipids 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 glyco sphingolipid 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 glyco sphingolipid by chemical methods. A final matter of caution, when using anti glycosphingolipid antibody for direct detection of glyco sphingolipids 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 power ful 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 glyco sphingolipids 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 pre cipitates 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 characteriza tion 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 g 1 fresh tissue (/g fresh myelin in the case of myelin). The molecular weight of ceramide, Gal Cer, sulphatide and sphingomyelin was established as sug gested 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), Ogawa Goto 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 g 1 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 g 1 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 bo d character: individua gang iosides covering more than 10% of the tota gang ioside bound Neu5Ac. Figures in bo d ita ic character: individua gang iosides covering from 7 to 10% of the tota gang ioside bound Neu5Ac a With six or more sia osy residues

Animal: Human 2.0 Bovine Pig Rat Mouse Chicken Pigeon Lizard Tortoise Frog Sa amander Be one Sa mon 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)

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(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 demon strated 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 detectab e

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

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 detectab e

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

GD3

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

136

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

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 tetra sialogangliosides 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 ganglio sides (Chou et al., 1990). Notably, all these ganglioside species are not present in adult mouse cerebrum

137

138

6

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 1 phosphate and ceramide 1 phosphate, are typical membrane components, particularly of the plasma mem brane. 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 character ized 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 synapto somes (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 fraction ation 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 gang iosides in each anima are given in bo d character. n.d. = not detectab e *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 Mo ecu ar ayer Parkinje ce ayer Granu ar ayer White matter Cerebral cortex Mo ecu ar ayer ( ) Externa granu ar ayer ( ) Externa pyramida ayer ( ) nterna granu ar ayer ( V) nterna pyramida ayer (Va/Vb) Po ymorphic ce ayer (V ) White matter Hippocampal formation Hippocampus: A veus Stratum oriens Stratum pyramida e

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 –

– – – – –

3+, strong reactivity; 2+, moderate reactivity; 1+, weak reactivity; -, negative reactivity; nd, not determined a O-AcDG, O-acety ated disia ogang iosides

Stratum radiatum Stratum acunosum–mo ecu are Dentate gyrus: Po ymorphic ayer Granu ar ayer Mo ecu ar ayer Spina cord Gray matter: Dorsa horn Anterior horn Ventra horn Posterior horn ntermediate zone White matter: Anterior funicu us Ventra funicu us Latera funicu us Dorsa funicu us Posterior funicu us 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

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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 (Rosner and Rahmann, 1987; Rosner 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 depen dency, selective apoptosis; (5) further extension of neuronal connections, second period of glial prolifera tion, 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 pre natal 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). Folch Pi (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 concentra tion 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 arboriza tion, 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

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. Figure 6 5 Effect of post natal age on the distribution of total phospholipids, cholesterol, Gal Cer, sphingomyelin, sulpha tide and gangliosides in the whole brain of rat. All substances are expressed as mmol g 1 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, sphin gomyelin, sulphatide and gangliosides in the human cerebral cortex (gray matter and white matter). All substances are expressed as mmol g 1 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), Svenner holm 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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. Figure 6 7 Distribution of individual gangliosides in whole chicken brain during pre and post hatching life. Each ganglio side 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, matura tion 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

3 – 38 80 140 290 63 90

5 – 80 190 130 195 120 28

7 – 170 375 140 70 90 35

1/2 110 230 750 190 130 45 42

1 – 270 845 245 305 40 43

2 102 365 1,020 290 380 38 45

5 98 440 1,030 430 420 36 44

8 96 463 1,053 527 423 77 –

Frontal cortex Post-natal age (years)

The most abundant gang iosides in each tissue are given in bo d character

Ganglioside GM2 GM1 GD1a GD1b GT1b GQ1b GD3

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 ganglio side 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 develop mental behavior of GD3 in rat cerebellum. Notably, GM1 and O Ac GT1b undergo the highest accumula tion (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 differentia tion. 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 Stallberg 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 unfractio nated 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

C25:1 C26:1

0.4 0.6

0.1 0.2

0.1 0.2

0.2/1.5 0.2/3.4

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 /3.4 /6.0

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

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

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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. Glyco sphingolipids 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 associ ation 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

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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

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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 diacylgly cerol 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 compo nent 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 neuro protective 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 ganglio side 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 phos pholipase C; PL, phospholipid; PLA2, phospholipase A2; PLase, phospholipase; PLC, phospholipase C; PLD, phospholipase D; PtdCho, phosphatidylcholine; PtdEtn, phosphatidylethanolamine; PtdIns, phosphatidy linositol; PtdIns(4)P, phosphatidylinositol 4 phosphate; PtdIns(4,5)P2, phosphatidylinositol 4,5 bisphosphate; 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 phosphatidylethanol amine (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 endo nuclear 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 compo sition 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 methodolo gy 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 transcrip tional 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 middle sized 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 differen tial 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 centrifuga tion 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 phosphati dylethanolamine (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 peroxida tion 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). Phospha tidylinositol 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 phosphoi nositide, 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 com plexes 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 phosphoi nositide 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 phosphoinosi tides 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 consen sus, 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 cholesterol sphingomyelin/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 SM synthase, 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 oligosac charide 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 ganglio sides 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 ganglio side; 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, devel oped 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 lympho cytes 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.

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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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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 perme ability 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 Sjostrand (Palade, 1952; Sjostrand, 1953). Structural and func tional 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 mem branes (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. Interest ingly, 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 manifesta tion 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. Charac terization 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 mito chondria 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 character istics 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 mitochon drial 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 mito chondrial 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 18:2 (n 20:3 (n 20:4 (n 22:6 (n

9) 6) 6) 6) 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) a Ruggiero et al., 1992

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 remain ing 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 demon strated 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 (Rytomaa 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

Note: Data are mean  SD and are expressed as percent (wt/wt) a Nakahara et a ., 1991; b Sun and Gi boe, 1994b; c Unpub ished data from our aboratory

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 (Rusinol 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 radioac tive 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 promot ing 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 mitochon dria 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 mitochon dria (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. Experi ments 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 distin guished 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 palmitoyl transferase 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 phos pholipase 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 laborato ry). 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 over expression 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 intra mitochondrial 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, submitochon drial 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 phosphati date 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 conver sion 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, sphingo myelin, 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 forma tion 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 hyper glycemic 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 reoxygena tion‐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 neurolog ical disorders have been reported in an excellent review (Farooqui et al., 2000b). Alterations in glyceropho spholipid 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). Plas malogens 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 choles terol 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 mitochon drial 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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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 glycerophospha tidylinositol, 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, lysophosphati dylcholine; 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 clathrin mediated 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.

225

ns(1,4,5) P3 DAG

PS

PE

PC

nosito (1,4,5) triphosphate Diacy g ycero

Phosphatidy serine

Phosphatidy ethano amine

Phosphatidy cho ines

Lipid name Phosphatidy inosito -4,5bisphosphate

Membrane structure, cho ine homostatsis, ACh precursor

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

SV priming

SV priming, Ca2+ eve s

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

P asma membrane (hydrophobic region), P asma membrane (inner eaflet), SV membrane P asma membrane (inner eaflet), SV membrane P asma membrane (outer eaflet), SV membrane

Location (mainly) P asma membrane (inner eaflet), SV membrane, Cytoso

Schizophrenia, progressive epi epsy with menta retardation Binding to snake venom phospho ipase A2, A zheimer’s disease

A zheimer’s disease

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

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

Michae son et a . (1983), Kim et a . (2004), G omset (2006), Yeung et a . (2008) Hansen et a . (1998), G omset (2006), Okamoto et a . (2007)

Reference Cremona et a . (1999), Di Pao o et a . (2004), Lee et a . (2004), Wenk and De Cami i (2004), Mi osevic et a . (2005), Nakano-Kobayashi et a . (2007)

Neuronal membrane lipids

Common Abbrev. P (4,5)P2

9

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

226 their role in the synaptic vesicle cycle

Cer

SM

Ceramide

Sphingomye in

Su fatides

Cho

Cho estero

Unc ear

Membrane structure via ceramide backbone

SV formation, fusion, structure, and membrane fluidity, signa ing

Membrane fluidity

Mye in membrane

Cytoso , p asma membrane (outer eaflet), SV membrane Ce p asma membrane (outer eaflet), SV membrane

SV and ce p asma membrane (inner and outer eaflet)

A zheimer’s disease, metachromatic eukodystrophy

Metachromatic eukodystrophy

Farber’s, Sandhoff, Gaucher diseases

Niemann–Pick disease, metachromatic eukodystrophy

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

Pe kofer and Sandhoff (1980), Deutsch and Ke y (1981), Goritz et a . (2002), Vance et a . (2006), Gy ys et a . (2007), Wasser et a . (2007) Carrer et a . (2003), Rohrbough et a . (2004), Snook et a . (2006)

Neuronal membrane lipids their role in the synaptic vesicle cycle

9 227

228

9 2

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 phosphory lation 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 4,5 bisphosphate (PI(4,5)P2, > Figure 9 3a). Many neuronal proteins bind to the inositol phosphate head group 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 pre synaptic 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) Phospha tidylserine 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 glyco sphingolipid that has a sulfate group on the 3 OH position of the galactose

Neuronal membrane lipids . Figure 9 3 (continued)

their role in the synaptic vesicle cycle

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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 di valent 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

9

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 Arachidonoyletha nolamine, 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 ac ylation of PE to generate N acylphosphatidylethanolamine (NAPE); (2) cleavage of NAPE by a phosphodi esterase 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).

Neuronal membrane lipids

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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 Echten Deckert 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 ganglio side (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 hyperex citability, 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 glycerophospholi pids, 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 endocanna binoids 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 schizophre nia, 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, Galb3 GalNAcb4(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, Neu5Aca8Neu5Aca3 Galb3GalNAcb4(Neu5Aca8Neu5Aca3)Galb4GlcCer; Gro 3P, glycerol 3 phosphate; GT1a, Neu5Aca8Neu5 Aca3Galb3GalNAcb4(Neu5Aca3)Galb4GlcCer; GT1b, Neu5Aca3 Galb3GalNAcb4(Neu5Aca8Neu5Aca3) Galb4GlcCer; GT1c, Galb3GalNAcb4(Neu5Aca8Neu5Aca8Neu5Aca3)Galb4GlcCer; HMG CoA, 3 hy droxy 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 biosyn thesis 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) physiol ogy. 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 phospho sphingolipids 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 dihydro ceramide, 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 longevity assurance 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. Gang liosides 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) dihydrocer amide 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 sphingo sine 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 phosphatidylinositol 4 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 decarboxyl ase, (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 acetoacetyl CoA 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 biosyn thesis 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 phos phohydrolase, (8) phosphatidate cytidyltransferase, (9) phosphatidylinositol synthase, (10) phosphatidylcholine: ceramide choline phosphotransferase, (11) diacylglycerol cholinephosphotransferase, (12) diacylglycerol ethanola minephosphotransferase, (13) phosphatidylethanolamine N methyl transferase and phosphatidyl N methyletha nolamine 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 3 phosphate 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 glycer ophospholipids of the mammalian tissue are phosphatidylcholine (PtdCho), phosphatidylethanolamine (PtdEtn), phosphatidylserine (PtdSer), and phosphatidylinositol (PtdIns). Besides the above phospho lipids, 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 succes sively 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 CDP choline 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 phosphatidylgly cerol phosphate (PtdGroP) by PtdGroP synthase. PtdGroP is subsequently dephosphorylated to phospha tidylglycerol (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 catalyz ing 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 antero grade 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; Muniz 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 acyla tion. 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 myelin like 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 compart ment, 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 membrane associated 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 con firmed 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 CHAPS insoluble membrane fraction after leaving the ER, but before exiting the Golgi, suggesting that myelin lipids

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and proteins assemble in the Golgi complex before transport to the myelin sheath (Kramer 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 compari son 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 phospho lipids. 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 prepara tions 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

10

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, long chain fatty acids (LFAs), and very long chain FAs (VLCFAs). The richest source of saturated and monoun saturated 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 supplemen tation 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 OL specific 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 pro oligodendroblast (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 char acterized 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)

Functional dynamics of myelin lipids

10

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 differen tiation 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 anti GalCer 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 ganglio sides 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 sialosylparaglobo side. 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

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The The The The

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

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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 myo inositol 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 endo plasmic 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).

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. 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 consider able 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

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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 PDGF stimulated 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 pleckstrin homology (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 mod ulates 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 phos phorylate 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 over expression 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 morphol ogy, 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). Knock down 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 Golgi localized, 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 establish ment 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 over expressed 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 presum ably 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 wild type 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 to plasma 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 endocy tosis, 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 phos phoinositides. 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. Pulse labeling 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)P2 binding PH domains, such as those of the b spectrins, inhibit ER to Golgi transport of VSVG in permea bilized 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 phospho lipid 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 zebra fish 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 Sacchar myces 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 phosphoi nositides (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 Charcot Marie 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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The phosphoinositides

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 OCR