Cytoskeleton and Human Disease [1 ed.] 1617797871, 9781617797873

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Cytoskeleton and Human Disease

Maria Kavallaris Editor

Cytoskeleton and Human Disease

Editor Dr. Maria Kavallaris Tumour Biology and Targeting Program Children’s Cancer Institute Australia for Medical Research Randwick, New South Wales Australia

ISBN 978-1-61779-787-3 ISBN 978-1-61779-788-0 (eBook) DOI 10.1007/978-1-61779-788-0 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2012934745 © Springer Science+Business Media, LLC 2012 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o 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. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

This book is dedicated to the memory of: Rhys Appelt Jarratt 13 June 1990—16 April, 2002. A passionate and aspiring young scientist that challenged me to find cures for cancer.

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Preface

The cytoskeleton is central to many critical cell processes including those that govern cell movement, migration, cell division, transport and cell signaling. It is well know that the cytoskeleton is often altered in disease and there is increasing interest in the role and mechanisms by which the cytoskeleton is involved in this process. Emerging evidence of the molecular and cellular events that drive cytoskeletal mediated disease including cancer, heart disease, myopathies and skin disorders, are also helping to shape targeted therapeutic approaches to treat these conditions. Diseases can also inform the normal function of genes and proteins and this has been the case for a number of cytoskeletal components. Advances in genetics, gene and protein identification, live cell and fixed imaging, and animal models of disease have accelerated knowledge in the cytoskeletal field. The Cytoskeleton and Human Disease book is intended to bring together key progress specifically related to the involvement of the cytoskeleton in different disease states from the basic science to clinical perspective. The complexity of the cytoskeleton from a mechanistic and structural perspective is enormous. The three major filamentous components—microtubule, microfilaments and intermediate filaments—interacting with each other and hundreds of accessory proteins impart plasticity to the cytoskeleton for many cellular functions. Mutations, deletions, alterations in components of the cytoskeleton can lead to defects in normal function and illness. The book is divided into three broad themes covering the basics of the cytoskeleton to associated disease states. Part I discusses the normal structure and function of actin and microtubules and also includes detailed descriptions of kinesin and myosin structure and function. The second part focuses on actin and disease, including alterations in actin interacting, cross-linking and regulating proteins that perturb the function of the actin cytoskeleton. These include, membrane organization and actin in T-lymphocytes, causes of thin filament disease in muscle, spectrum of filamin diseases, the actin regulating protein LIM kinase in metastasis, mutations in actin associated with deafness, and the therapeutic targeting of actin. Part III covers microtubules as cancer therapeutic targets and their role in drug resistance, as well as the emerging field of posttranslational modifications of tubulin. This part of the book also covers the microtubule interacting protein stathmin and its role in cancer, the biology and pathobiology of Tau and tubulinrelated malformations of cortical development. The final section, Part IV, centers on vii

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Preface

intermediate filaments and diseases that includes spectrin disorders, laminopathies, desmin and heart disease and neurodegenerative diseases and intermediate filaments. There have been great advances in our understanding of the cytoskeleton since the French scientist Paul Wintrebert first introduced the notion and coined the term cytoskeleton in 1931. This book links expert knowledge and brings to the fore unifying areas of overlapping interest that will stimulate further research and a better understanding of the cytoskeleton in health and disease. Maria Kavallaris PhD Editor, The Cytoskeleton and Human Disease

Acknowledgements

Enormous thank you to Melissa Vincent for her support, organizational skills and humor in the preparation of this book.

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Contents

Part I Basics of the Cytoskeleton 1 ACTIN . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vera Dugina, Richard Arnoldi, Paul A. Janmey and Christine Chaponnier

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Microtubules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pavel Dráber and Eduarda Dráberová

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The Kinesin Superfamily . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Linda Wordeman

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Basics of the Cytoskeleton: Myosins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Omar A. Quintero, Judy E. Moore and Christopher M. Yengo

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Part II Actin and Disease 5

The Actin Cytoskeleton and Membrane Organisation in T Lymphocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 Rhea Cornely, Thomas Grewal and Katharina Gaus

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Thin Filament Diseases of Striated Muscle . . . . . . . . . . . . . . . . . . . . . . . . 123 Anthony J. Kee and Edna C. Hardeman

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Filamins and Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 141 Stephen P. Robertson and Philip B. Daniel

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LIM Kinase and Cancer Metastasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159 Alice Schofield and Ora Bernard

9 Actin Mutations and Deafness . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 Matías Morín, Fernando Mayo, Felipe Moreno and Miguel A. Moreno-Pelayo

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Contents

10 Therapeutic Targeting of the Actin Cytoskeleton in Cancer . . . . . . . . . 181 Teresa Bonello, Jason Coombes, Galina Schevzov, Peter Gunning and Justine Stehn Part III Microtubules and Disease 11 Microtubules as a Target in Cancer Therapy . . . . . . . . . . . . . . . . . . . . . . 203 April L. Risinger and Susan L. Mooberry 12 Microtubules, Drug Resistance, and Tumorigenesis . . . . . . . . . . . . . . . . 223 Joshua A. McCarroll and Maria Kavallaris 13 Posttranslational Modifications of Tubulin . . . . . . . . . . . . . . . . . . . . . . . . 241 Suzan K. Chao, Chia-Ping H. Yang and Susan Band Horwitz 14 Stathmin and Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 259 Dominic Chi Hiung Ng and Frances Byrne 15 The Biology and Pathobiology of Tau Protein . . . . . . . . . . . . . . . . . . . . . . 285 Garth F. Hall 16 Tubulin-Related Malformations of Cortical Development . . . . . . . . . . . 315 Xavier H. Jaglin, Jamel Chelly and Nadia Bahi-Buisson Part IV Intermediate Filaments and Disease 17 Spectrins in Human Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 345 Marie-Christine Lecomte 18 Laminopathies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 375 Nadir M. Maraldi and Giovanna Lattanzi 19 Desmin and Heart Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 411 J. Scott Pattison and Jeffrey Robbins 20 Neurodegenerative Diseases and Intermediate Filaments . . . . . . . . . . . 425 Rodolphe Perrot and Jean-Pierre Julien Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 449

About the Editor

Maria Kavallaris, Ph.D. is Head of the Tumour Biology and Targeting Program and a National Health and Medical Research Council Senior Research Fellow at the Children’s Cancer Institute Australia. She also holds appointments as Director of the Australian Centre for Nanomedicine and Professor at the University of New South Wales. Maria has made fundamental contributions in identifying mechanisms of action and resistance to anticancer drugs that target tubulin and discovering new cytoskeleton interactions in cell division and tumor formation. She obtained her Ph.D. from the University of New South Wales and undertook her postdoctoral studies as an International Agency for Cancer Research Fellow at the Albert Einstein College of Medicine, New York. She has served on numerous grant review panels, scientific advisory committees, pharmaceutical advisory boards and is a past President of the Australian Society for Medical Research. Maria is a Director of the Australian Institute for Policy and Science.

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Contributors

Richard Arnoli Department of Pathology and Immunology, Faculty of Medicine, University of Geneva, Geneva, Switzerland Nadia Bahi-Buisson Université Paris Descartes, CNRS UMR 8104, Institut Cochin, Laboratoire de Génétique des Maladies Neurodéveloppementales, Paris, France and Institut National de la Santé et de la Recherche Médicale (Inserm), U1016, Paris, France Ora Bernard St. Vincent’s Institute of Medical Research and The University Melbourne Department of Medicine, St. Vincent’s Hospital, Fitzroy, VIC, Australia Teresa Bonello School of Medical Sciences, University of New South Wales, Sydney, NSW, Australia Frances L. Byrne Children’s Cancer Institute Australia, Lowy Cancer Research Centre, University of New South Wales, Sydney, NSW, Australia Australian Centre for Nanomedicine, Faculty of Engineering, University of New South Wales, Sydney, NSW, Australia Suzan K. Chao Department of Molecular Pharmacology, Albert Einstein College of Medicine, Bronx, NY, USA Christine Chaponnier Department of Pathology and Immunology, Faculty of Medicine, University of Geneva, Geneva, Switzerland Jamel Chelly Université Paris Descartes, CNRS UMR 8104, Institut Cochin, Laboratoire de Génétique des Maladies Neurodéveloppementales, Paris, France and Institut National de la Santé et de la Recherche Médicale (Inserm), U1016, Paris, France Jason Coombes School of Medical Sciences, University of New South Wales, Sydney, NSW, Australia Rhea Cornely Centre for Vascular Research, University of New South Wales, Sydney, Kensington, NSW, Australia

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Contributors

Philip B. Daniel Department of Paediatrics and Child Health, Dunedin School of Medicine, University of Otago, Dunedin, New Zealand P. Dráber Laboratory of Biology of Cytoskeleton, Institute of Molecular Genetics, Academy of Sciences of the Czech Republic, Prague, Czech Republic E. Dráberová Laboratory of Biology of Cytoskeleton, Institute of Molecular Genetics, Academy of Sciences of the Czech Republic, Prague, Czech Republic Vera Dugina Department of Pathology and Immunology, Faculty of Medicine, University of Geneva, Geneva, Switzerland and A. N. Belozersky Institute of Physical and Chemical Biology, Moscow State University, Moscow, Russia Katharina Gaus Centre for Vascular Research, University of New South Wales, Sydney, Kensington, NSW, Australia Thomas Grewal Faculty of Pharmacy, University of Sydney, Sydney, NSW, Australia Peter Gunning School of Medical Sciences, University of New South Wales, Sydney, NSW, Australia Garth Hall Center for Cellular Neuroscience and Neurodegeneration Research, Department of Biological Sciences, University of Massachusetts Lowell, Lowell, MA, USA Edna C. Hardeman Neuromuscular and Regenerative Medicine Unit, School of Medical Sciences, University of New South Wales, Sydney, NSW, Australia Susan B. Horwitz Department of Molecular Pharmacology, Albert Einstein College of Medicine, Bronx, NY, USA Xavier H. Jaglin Université Paris Descartes, CNRS UMR 8104, Institut Cochin, Laboratoire de Génétique des Maladies Neurodéveloppementales, Paris, France and Institut National de la Santé et de la Recherche Médicale (Inserm), U1016, Paris, France Paul A. Janmey Institute for Medicine and Engineering, University of Pennsylvania, Philadelphia, PA, USA Jean-Pierre Julien Department of Psychiatry and Neurosciences, Laval University, Research Centre of CHUQ, QC, Canada Maria Kavallaris Children’s Cancer Institute Australia, Lowy Cancer Research Centre, University of New South Wales, Sydney, NSW, Australia Australian Centre for Nanomedicine, Faculty of Engineering, University of New South Wales, Sydney, NSW, Australia Anthony J. Kee Neuromuscular and Regenerative Medicine Unit, School of Medical Sciences, University of New South Wales, Sydney, NSW, Australia Giovanna Lattanzi National Research Council of Italy CNR-IOR, Institute of Molecular Genetics, Unit of Bologna, Bologna, Italy

Contributors

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Marie-Christine Lecomte INSERM, U665, Paris, France; Institut National de la Transfusion Sanguine, Paris, France and Université Paris, Paris, France Nadir M. Maraldi Laboratory of Musculoskelatal Cell Biology, Rizzoli Orthopaedic Institute, Bologna, Italy and Department of Anatomical Sciences, University of Bologna, Bologna, Italy Fernando Mayo Unidad de Genética Molecular, Ramón y Cajal Institute of Health Research (IRYCIS) and Biomedical Network Research Centre on Rare Diseases (CIBERER), Madrid, Spain Joshua McCarroll Children’s Cancer Institute Australia, Lowy Cancer Research Centre, University of New South Wales, Sydney, NSW, Australia and Australian Centre for Nanomedicine, Faculty of Engineering, University of New South Wales, Sydney, NSW, Australia Felipe Moreno Unidad de Genética Molecular, Ramón y Cajal Institute of Health Research (IRYCIS) and Biomedical Network Research Centre on Rare Diseases (CIBERER), Madrid, Spain Miguel A. Moreno-Pelayo Unidad de Genética Molecular, Ramón y Cajal Institute of Health Research (IRYCIS) and Biomedical Network Research Centre on Rare Diseases (CIBERER), Madrid, Spain Matias Morin Unidad de Genética Molecular, Ramón y Cajal Institute of Health Research (IRYCIS) and Biomedical Network Research Centre on Rare Diseases (CIBERER), Madrid, Spain Susan L. Mooberry Department of Pharmacology, University of Texas Health Science Center at San Antonio, San Antonio, TX, USA Judy E. Moore Department of Biology, University of North Carolina Charlotte, Charlotte, NC, USA Dominic Ng Department of Biochemistry and Molecular Biology, Bio21 Molecular Science and Biotechnology Institute, University of Melbourne, VIC Australia J. Scott Pattison Department of Pediatrics, Division of Molecular Cardiovascular Biology, Cincinnati Children’s Hospital Medical Center, Cincinnati, OH, USA Rodolphe Perrot Department of Psychiatry and Neurosciences, Laval University, Research Centre of CHUQ, QC, Canada Omar A. Quintero Department of Cellular and Molecular Physiology, The Penn State College of Medicine, Hershey, PA, USA April L. Risinger Department of Pharmacology, University of Texas Health Science Center at San Antonio, San Antonio, TX, USA Jeffrey Robbins Department of Pediatrics, Division of Molecular Cardiovascular Biology, Cincinnati Children’s Hospital Medical Center, Cincinnati, OH, USA

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Contributors

Stephen P. Robertson Department of Paediatrics and Child Health, Dunedin School of Medicine, University of Otago, Dunedin, New Zealand Galina Schevzov School of Medical Sciences, University of New South Wales, Sydney, NSW, Australia Alice Schofield St. Vincent’s Institute of Medical Research and The University Melbourne Department of Medicine, St. Vincent’s Hospital, Fitzroy, VIC, Australia Justine Stehn School of Medical Sciences, University of New South Wales, Sydney, NSW, Australia Linda Wordeman Department of Physiology and Biophysics, University of Washington, Seattle, WA, USA Chia-Ping H. Yang Department of Molecular Pharmacology, Albert Einstein College of Medicine, Bronx, NY, USA Christopher M. Yengo Department of Cellular and Molecular Physiology, The Penn State College of Medicine, Hershey, PA, USA

Abbreviations

+TIPS 53BP1 AAA AAP ABP AD AD-CMT2 ADF ADLD ADNSHL ADP AIS ALS AO APC APS Arp2/3 Arp2/3 ARX ASPM ATP a-WA AXD BAF BCL BOS BTB CAP-Gly CBD CC CCM CCP1

Plus end binding proteins p53 binding protein 1 ATPases Associated with various cellular Activities Actin associated proteins Actin binding protein Alzheimer’s Disease Autosomal dominant for of CMT2 Actin-depolymerizing factor Autosomal dominant leukodystrophy Autosomal dominant nonsyndromic hearing loss Adenine Diphosphate Axin initial binding Amyotrophic lateral sclerosis Atelosteogenesis Adenomatous Polyposis Coli Atypical progeroid syndrome Actin related protein Actin related protein Aristaless-related homeobox gene Abnormal Spindle-like microcephaly-associated Adenine Triphosphate Atypical Werner syndrome Alexander Disease Barrier to autointegration factor B-cell lymphoma Buschke-Ollendorff syndrome Broad-complex, Tramtrack, and Bric a brac Cytoskeleton-Associated Protein Glycine-rich Cardio binding domains Critical Concentration Central core myopathy Cytosolic carboxypeptidase 1 xix

xx

CCP5 CD CFTD CH CLASP CLIP CMD1A CMs CMT CMT 2B1 CNS CP CSF cSMAC CSP CTT DA DBSs DCM DCX DES DLG DRM dSMAC DYT1 EB EBH ECM EDMD EGF ERK ERM EZH2 F-actin FAK FCM FERM FGF FH2 FHC FLN FLPD FMD FRET G-actin

Abbreviations

Cytosolic carboxypeptidase-like protein 5 Cap disease Congenital fiber type disproportion Calponin homology CLIP-Associated Protein Cytoplasmic Linker Protein Cardiomyopathy and conduction-system disease Cardiomyopathy Charcot-Marie Tooth Charcot-Marie Tooth disorder type 2 Central nervous system Cortical plate Cerebral spinal fluid Central supramolecular activation cluster Cystein string protein C Terminal Tail Distal arthrogryposis Double-strand breaks Dilated cardiomyopathy Doublecortin Desmin gene Disc-large Desmin related myopathy Distal supramolecular activation cluster Torsion dystonia End-binding EB homology Extracellular matrix Emery-Dreifuss muscular dystrophy Epidermal growth factor Extracellular signal-regulated kinase Proteins proteins of the ezrin, radixin and moesin family Enhancer of zeste homologue 2 Filamentous Actin Focal Adhesion Kinase Familial cardiomyopathies Founding members: band 4.1, ezrin, radixin, and moesin Fibroblast growth factor Formin homology 2 Familial hypertrophic cardiomyopathy Filamin Familial partial lipodystrophy Frontometaphyseal dysplasia Fluorescence resonance energy transfer Globular actin

Abbreviations

GADS GAN GDP GEF GFAP GFP GH GPCR GTD GTP HCM HDAC HE HGPS HHS HPP HRD HS HUGO HUVEC ICDs IF IGF1 IHCs INM ITK KAP KASH KO KSP LAP1 LAT LBR LC3 Lck LCR LGMD 1B LIMK LIS LS mAb MACF MADA MADB MAPK

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Grb2-related adaptor downstream of Shc Giant axonal neuropathy Guandinediphosphate Guanine nucleotide exchange factor Glial fibrillary acidic protein Green fluroscent protein Growth hormone G-protein coupled receptors Globular tail domain Guandine triphosphate Hypertrophic cardiomyopathy Histone deacetylase Hereditary elliptocytosis Hutchinson-Gilford progeria syndrome Heart-hand syndrome Hereditary pyropoikylocytosis Hypoparathyroidism/mental retardation/facial dysmorphism Hereditary spherocytosis Human Genome Organisation Human umbilical vein endothelial cells Implantable cardioverter-defibrillators Intermediate filament Insulin growth factor 1 Inner hair cells Inner nuclear membranes Interleukin-2-inducible T cell kinase Kinesin associated protein Klarscicht, ANC-1; Syne Homology Knock Out Kinesin spindle protein Lanina-associated polypeptide 1 Linker of activated T cells Lamin B receptor Light chain 3 subunit Lymphocyte specific protein tyrosine kinase Locus control region Limb girdle muscular distrophy type 1B LIM Kinase Lissencephaly Larsen Syndrome Monoclonal antibody Microtubule-Actin Crosslinking Factor Mandibuloacral dysplasia type A Mandibuloacral dysplasia type B Mitogen-associated potein kinase

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MAPs MARK 4 MCAK MDCL MDR MEFs MFs MHC miRNA MLCK MMM MNS MRI mRNA MRP MT MTBR MTOC MZ NCK NE NER NF NFH NFL NFM NFTs NGF NIFID NM NMDA NMR NSCLC NTI OHCs ONM OPDSDs PARDS PD PH PHA PHF PHN PKC PLCγ1

Abbreviations

Microtubule-associated proteins Microtubule affinity-regulating kinase 4 Mitotic centromere-associated kinesins LMNA-related congenital muscular dystrophy Multi-drug resistance Embryonic fibroblasts Microfilaments Major histocompatibility complex MicroRNA myosin light chain kinase Multi-mini core myopathy Melnick-Needles Syndrome Magnetic resonance imaging Messenger RNA Multi-drug resistance-associated protein Microtubules Microtubule binding repeat Microtubule-organising centers Marginal zone Non-catalytic region of tyrosin kinase Nuclear envelope Nucleotide excision repair Neurofilaments Neurofilament, heavy polypeptide Neurofilament, light polypeptide Neurofilament, medium polypeptide Neurofibrillary tangles Nerve-growth factor Neuronal intermediate filament inclusion disease Nemaline myopathy Non-N-methyl-D-aspartic acid Nuclear magnetic reasonance Non small cell lung cancer N-terminal insert Outer hair cells Outer nuclear membranes The optopalatodigital spectrum disorders Periaxiplasmic ribosomal plaques Parkinson’s disease Pleckstrin homology Pelger-Huet anomaly Paired helical filaments Periventricular heterotopia Protein Kinase C Phospholipase Cγ1

Abbreviations

PNH PNS PP PPIase pRb pSMAC PTMs PTMs RBC RCM RD RNA RTK SAH SBH SBMS SCAs SCCs SCTS shRNAi siRNA SLD SLP-76 SM SMC SOD1 SREBP1 SREs SRF STMN STOPs SUN SVZ TBA TBC TCR Tms TNAP TOD TOG TTCP TTL TTLL TZDs

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Periventricular nodular heterotopia Peripheral nervous system Pre-plate Peptidyl-prolylisomerase Phosphorylated retinoblastoma Peripheral supramolecular activation cluster Post-translational modifications Post-translational modifications Red blood cells Restrictive cardiomyopathy Restrictive dermopathy Ribonucleic acid Receptor tyrosine kinases Single alpha helix Subcortical band heterotopia Spectrin-based membrane skeleton Spinocerebellar ataxias Squamous Carcinoma Cells Spondylocarpotarsal syndrome Short hairpin RNAi Small-interfering RNA Stathmin-like domain Src homology 2 domain containing leukocyte phosphoprotein of 76kDa Smooth Muscle Smooth muscle cell Superoxide dismutase 1 Sterol regularoty element binding proteim 1 SRF-binding elements Serum response factor Stathmin Stable tubulin only proteins Sad1 and UNC-84 Subventricular zone Tubulin Binding Agents Tubulin specific chaperone T cell receptor Tropomyosins Tissue non-specific phosphatase Terminal osseous dysplasia Tumour overexpressed gene Tubulin-specific carboxypeptidase Tubuline tyrosine ligase Tubuline tyrosine ligase-like proteins Thiazolinediones

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UTRs VASp VDA VDAC VGSC WAS WASp WAVE WIP XLD-PH XLN γ-CYA γ-SMA γTuSC α-CAA α-SKA α-SMA β-actin β-CYA βMyHC γ-actin 3R 4R WT

Abbreviations

Untranslated regions Vasodilator-stimulated phosphoprotein Vascular disrupting agents Voltage-dependent anion channel Sodium-channel clusters Wiskott-Aldrich Syndrome Wiskott-Aldrich syndrome protein WASp-family verprolin-homologous protein-2 WASp interacting protein X linked periventricular nodular heteroptopia X linked neutropenia Gamma-cytoplamic actin Gamma-smooth muscle actin γ-tubulin small complex Alpha-cardiac muscle actin Alpha-skeletal muscle actin Alpha-smooth muscle actin Beta-actin Beta-cytoplasmic actin β-myosin heavy chain Gamma-actin Any one of the three possible 3 microtubule binding repeat tau isoforms (3R0N, 3R1N, 3R2N) Any one of the three possible 4 microtubule binding repeat tau isoforms (4R0N, 4R1N, 4R2N) Wild type

Part I

Basics of the Cytoskeleton

Chapter 1

ACTIN Vera Dugina, Richard Arnoldi, Paul A. Janmey and Christine Chaponnier

Abstract Eukaryotic cells contain three distinct cytoskeletal filament systems, including actin, that exhibit very different assembly properties, supramolecular architectures, dynamic behavior, and mechanical properties. Actin, which is involved in a plethora of functions, is the most abundant protein in most cell types and is impressively conserved across species. The highest concentrations of actin (about 20% of total protein) are found as stable microfilament systems assembled within myofibrillar contractile structures in striated muscles. In addition to its specialized role in muscle contraction, actin is present in all muscle and nonmuscle cells, where it plays a variety of roles thanks to its ability to assemble and disassemble depending on cell requirements. Actin plays an important role in maintaining cell structure and function by conferring mechanical strength and enabling intracellular contraction and/or tension. In nonmuscle cells, microfilaments are involved in cell motility and cytokinesis. The dynamics of the actin cytoskeleton are maintained by two factors: (1) the ability of actin to undergo reversible transformation from the monomeric state (G-actin) to the polymeric state (F-actin) and (2) the interaction of actin with actin-binding proteins (ABPs), that can inhibit or stimulate actin polymerization, sever the polymers, crosslink actin filaments into bundles or in filamentous three-dimensional networks, and bind them to cell membranes. Considering the multiple cellular functions of actin, alterations in the organization of microfilaments will result in disorganized cell arrangement and orientation, uncontrolled cell growth, and abnormal responses to the environment. Higher vertebrates express six different highly conserved actin isoforms. Over the last decades, numerous studies have tried to elucidate the specific expressions, localizations, regulations, properties, and functions of the different isoactins. The understanding of their specific underlying mechanisms would be of major relevance not only for fundamental research but also for clinical applications, since modulations of actin isoforms are directly or indirectly correlated with severe pathologies. C. Chaponnier () · V. Dugina · R. Arnoldi Department of Pathology and Immunology, Faculty of Medicine, University of Geneva, 1 rue Michel-Servet, 1211 Geneva, Switzerland e-mail: [email protected] V. Dugina A. N. Belozersky Institute of Physical and Chemical Biology, Moscow State University, Moscow, Russia P. A. Janmey Institute for Medicine and Engineering, University of Pennsylvania, Philadelphia, PA, USA M. Kavallaris (ed.), Cytoskeleton and Human Disease, DOI 10.1007/978-1-61779-788-0_1, © Springer Science+Business Media, LLC 2012

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1.1

V. Dugina et al.

Introduction

The dynamic formation and dissolution of actin filament networks and their coupling to the cell membrane, molecular motors, and other intracellular structures are essential for cell motility, transport, and many other aspects of cell biology. In addition to their assembly into polymer networks that determine the cell’s mechanical properties, actin subunits and filaments bind hundreds of intracellular ligands and perform many cellular functions, some of which are only beginning to be identified. The recent finding of prokaryotic analogs of actin (e.g., MreB; [1, 2]) suggests functions beyond those generally involved in eukaryotic cell movements and viscoelasticity. In vertebrates, six actin isoforms, that is, two striated muscle (α-skeletal, α-SKA and αcardiac, α-CAA), two smooth muscle (α- and γ-SMA) and two cytoplasmic (β-CYA and γ-CYA) are encoded by distinct genes [3]. Muscle actins are tissue-specific and organized into contractile units, whereas β- and γ-CYA are ubiquitous and essential for cell survival [4]. All actin isoforms exhibit similar primary structures and the main differences among them are located at the N-terminus of the molecules.

1.2 Actin The actin monomer is a moderate size (42 kDa) globular protein that is characterized by a large cleft in its middle in which a divalent metal ion, usually Ca2+ or Mg2+ , and a nucleotide, ATP or ADP, are bound by specific interactions with amino acid side chains that line the cleft, as shown in Fig. 1.1. The N-terminus, where most differences among actin isoforms lie, is a flexible chain that extends from the body of the monomer. Actin was first identified as part of the acto-myosin complex responsible for producing the contractile force in skeletal muscle [5]. Actin is one of the most strictly conserved proteins, with a very similar amino acid sequence in species ranging from plants to animals. Actin monomers are modified by several different modifications, both co- and post-translational. Actin is acetylated on its N-terminal residue, and nearly all actins are modified at His 73 by methylation [6]. Post-translational phosphorylation of monomeric actin has also been reported [7]. Actin can also be ADP-ribosylated by Clostridium perfringens iota toxin and this modification prevents filament elongation [8]. A comprehensive listing of natural and mutant actin modifications is found in [9]. A recently discovered post-translational modification of both actin and several ABPs is arginylation.1 This modification reduces the surface charge of the filament and appears to destabilize the cytoskeleton in vivo [10, 11]. Monomeric actin binds to intracellular and nuclear proteins such as DNase 1 and potentially regulates some cellular events, but most of actin’s functions are due to its polymerization into filaments, as shown in Fig. 1.2. Actin filament structural models suggest that actin’s N-terminal residues protrude from the filament surface 1

Arginylation is a post-translational modification mediated byArg-tRNA protein transferase (ATE1) which transfers arginine (Arg) from tRNA onto proteins.

1 ACTIN

5

Fig. 1.1 Structure of monomeric actin (PBD code: 1J6Z) showing the nucleotide and metal ion-binding cleft in the middle and the unstructured N-terminus (red). Black numbers refer to the conventional assignment of actin’s four subdomains. The C-terminus, where most fluorescent labels are covalently attached, emerges from the back of subdomain 1

Fig. 1.2 Structural model for F-actin adapted ([12] (PDB code: 3G37)). The N-termini of three adjacent monomers are labeled as N

(Fig. 1.2; [12]), suggesting that the differences among actin isoforms are likely to affect the binding of other proteins to the filament. The polymerization of actin into filaments is usually modeled as analogous to the polymerization of low molecular weight monomers into polymers. Kinetic schemes based on mechanisms extended from traditional polymer chemistry have

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been remarkably powerful for quantifying the rates and extents of actin polymerization and for predicting the effects of proteins that bond subunits or the ends of polymers [13, 14]. Actin polymerization in vitro is well approximated as a nucleationelongation reaction, with additional effects caused by the hydrolysis of nucleotides that proceeds during polymerization [15]. At the lowest level of approximation, actin assembles in three stages: formation of actin nuclei, elongation of filaments from one or both filament ends, and maturation to a steady state where subunit assembly and disassembly from the two ends balance. Stage one: filament nucleation. Even under polymerizing conditions, the affinity of actin monomers for each other is very low, but when trimers or larger oligomers form, they are stable and act as seeds onto which additional monomers can add [16]. The structure of the actin monomer is asymmetric, and monomers bind each other head-to-tail (as well as side-to-side) so the nuclei for polymerization are also asymmetric, leading to a polarity in the filament. By convention, based on the structure of F-actin complexed with a myosin fragment that gives the filaments an arrow-head appearance, one end of the filament is called the “barbed end” and the other the “pointed end.” These two ends are not only structurally distinct but the kinetics and affinity with which monomers add to these ends are also different. Stage two: filament growth. Elongation of actin filaments occurs both by subunit addition to the two filament ends and by annealing of filaments with each other endto-end. Actin polymerization occurs mainly by addition of monomers to the ends of the nucleus or growing filament. The on- and off-rates of subunits for the two ends of the filaments are different. The barbed end assembles more rapidly than the pointed end when there is an excess of monomer in solution. In addition to elongation by subunit addition, end-to-end annealing of short filaments produces a longer filament that preserves the asymmetry of the two ends [17]. The extent to which this reaction contributes to cytoskeletal assembly in vivo is uncertain [18, 19]. Under typical conditions for maintaining unpolymerized actin (or G-actin) in vitro, the nucleotide is ATP and the metal ion is Ca2+ in the monomer, and this monomeric subunit or its complex with proteins like profilin that help direct it to the barbed end of a nucleus or growing filaments is the form of actin that associates to elongate a polymer. As polymerization proceeds, ATP is hydrolyzed to ADP-Pi and later the phosphate is released and Ca2+ is exchanged for Mg2+ . As a result, the configuration of the actin monomer unit undergoes subtle changes, and its structure within the filament is thought to be somewhat strained with respect to its lowest energy state. ATP hydrolysis is not required for polymerization, since ADP-actin also polymerizes, but nucleotides and divalent cations strongly affect polymerization rate constants and the affinities of the filament ends for monomer units. The actin filament, also called F-actin, is a helical filament with a pitch of 37.5 nm, a diameter of approximately 8 nm, a shape closer to a thick ribbon than a cylinder, and a length that can be many microns, and is regulated by a number of proteins that bind the filament ends. The two ends of the filament are structurally and functionally distinct. The barbed end has higher affinity for actin monomers, but under most conditions also faster on- and off-rate constants compared to the barbed end.

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Stage three: filament maturation, steady state, and disassembly. Elongation of an actin filament to a steady state length is not the end of the assembly process. The conformation of the subunit that adds to the filament is different from that of the subunit that dissociates from it. Much of this difference results from changes in the nucleotide and metal ion bound within the subunit, but other less well-characterized changes in subunit packing or conformation occur as the filament ages, and these changes can affect the affinity of various actin filament-binding proteins. Some issues related to the structural plasticity of F-actin have recently been reviewed to make a compelling case for their biological significance [20].

1.2.1

Nucleotide Hydrolysis by Actin

Under the conditions thought to exist in the cytoplasm, actin monomers that are competent to add to a filament bind ATP in their nucleotide-binding cleft, and subunits that dissociate from a filament generally have ADP. Nucleotide hydrolysis resulting from actin polymerization was identified in early studies of actin chemistry (reviewed in [21]), but the biological function of this hydrolysis is still partly unknown. The nucleotide hydrolyzes only after the monomer is incorporated into the filament, but precisely how far into the filament or at which time the hydrolysis happens is still a matter of significant research and discussion [15, 22]. Most data suggest that the energy produced by ATP hydrolysis imparts internal stress within the polymer and thereby makes its disassembly more favorable under some conditions. An additional consequence of the slow hydrolysis of nucleoside triphosphate and the release of phosphate remaining bound to the filament is that newly formed filaments are chemically distinguishable from old filaments. Many proteins, for example, the actin-severing proteins cofilin [23, 24] and gelsolin [25], selectively bind ADP-actin, and therefore newly assembled actin at the growing edge of the cytoskeleton is relatively protected from severing compared to older actin filaments in the cell interior [26]. The distinction between old and new filaments that alters ABP affinity does not appear to involve direct binding to the nucleotide, but rather suggests that the conformations of monomers within a polymer retain enough flexibility or structural plasticity to present different surfaces on the filament exterior depending partly on the bound nucleotide [20].

1.2.2 Actin-Binding Proteins Actin filaments are not intrinsically self-associating, but in vivo they form a wide variety of super-filament structures that are determined by bridging interactions mediated by multivalent counterions and large classes of ABPs [27, 28]. There are currently more than 275 different ABPs reported [27], (en.wikipedia.org/wiki/Actin-binding protein, February 2012). The specific biochemical features of many of these proteins have been summarized in several excellent recent reviews [27, 29–31].

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Mechanical Properties of the Cytoskeleton

The unique physical properties of the actin-based cytoskeleton are largely due to the unusual stiffness of actin filaments. These properties include formation of solidlike gels at very low concentrations (1,000) reflects the importance of this antibody. The availability of such an antibody has motivated consideration of α-SMA as a marker of differentiation, not only in the typical vascular SMCs during development, during atheromatous process, and in various pathological situations, but interestingly also in a large number of other cells such as myofibroblasts, myoepithelial cells, myoid cells, and pericytes [109]. These cells could be classified as SM-like cells because of the organization of a contractile apparatus detected at the EM level and the expression of α-SMA. The large number of studies conducted on myofibroblasts, designed to reveal their mechanisms of differentiation, their contractile activity and their origin, demonstrate the importance of using the α-SMA antibody as a tool. First described in woundhealing granulation tissue [132, 133], and subsequently in numerous pathological settings including hypertrophic scars, fibromatoses, and stromal response to neoplasia [134], the myofibroblast is a mesenchymal cell combining features of fibroblasts and SMCs. Under the influence of mechanical stress (enhanced stiffness and tension of the extracellular matrix), and the combined action of ED-A fibronectin and TGF-β, fibroblasts evolve into differentiated myofibroblasts [135]. This transition is accompanied by the development of α-SMA-positive stress fibers and the formation of “supermature” focal adhesions [136] at the cell-matrix interface. The myofibroblast appears to originate not only from the fibroblast but also from transdifferentiation of endothelial or epithelial cells [137]. Therefore, the immunodetection of α-SMA has been, and still is, a critical tool for analysis of wound healing, development of fibrotic diseases, stromal reaction, and also basic research to understand the mechanisms of differentiation and possibly to manipulate contractile activity during excessive wound healing or fibrotic diseases [138]. Interestingly, a subpopulation of cardiomyocytes expressing α-SMA has recently been detected in the adult rat and human cardiac conduction systems [139]. This type of investigation further supports the concept of a different type of contractile activity in α-SMA-containing cells compared with sarcomeric actin-containing cells, implying specific functional activities for actin isoforms (see below stem cell differentiation-peptide-siRNA in cardiac development, [124]).

1.5.2.2

γ-Smooth Muscle Actin

In embryonic chicken gizzard muscles, γ-SMA, β- and γ-CYAs are synthesized [140, 141]. With development, the relative amount of γ-SMA rises at the expense of

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the cytoplasmic isoforms and the ratios of γ-SMA to β-CYA accumulated in chicken gizzard at the time of hatching are 3.6 at the protein level and 2.2 at the mRNA level. According to Vandekerckhove et al. [3] and Hirai et al. [142], in adult chicken gizzard, the γ-isoform (essentially γ-SMA) is dominant (about 80%), the β-isoform is also readily detected, and the α-isoform is absent. Further 2D-PAGE studies from Izant et al. [143] and Stromer et al. [144] showed that in addition to larger amounts of γ-SMA, smaller amounts of α-SMA are present in gizzard. As far as γ-SMA is concerned, uterine smooth muscle is an interesting tissue since during the course of pregnancy it undergoes dramatic physiological adaptations resulting in major myometrial hypertrophy and remodeling. According to studies from Cavaille et al. [145] and Skalli et al. [146], rat as well as human pregnant myometria is characterized by a considerable increase in the relative amount of γ-SMA relative to α-SMA. According to Shynlova et al. [147], the rat myometrium did not display significant changes in α-SMA gene expression, whereas γ-SMA levels increased by up to 32-fold at the mRNA level, and 17-fold at the protein level compared to nonpregnant samples, followed by a rapid return to nonpregnant levels in the postpartum period. So far a detailed study of γ-SMA expressing cells is still partial because of the lack of a specific antibody till very recently (Arnoldi and Chaponnier, unpublished).

1.5.3

Cytoplasmic Actins

The cytoplasmic actins (β and γ) play crucial roles during key cellular processes like adhesion, migration, polarization, and cytokinesis. In most nonmuscle cells of vertebrates, about half of actin is present in its monomeric state, suggestive of a highly dynamic polymerization-depolymerization activity. Nonmuscle cells such as epithelial cells express only β- and γ-CYAs, although in various proportions [148]. β- and γ-CYAs differ only by four amino acids located at position 1, 2, 3, and 9. Albeit specific functions have been suggested mainly for muscle actin isoforms, largely based on studies of mutations, knock-out mice and actin-dependent pathologies [100–102] little is known about the possible specific roles of β- and γ-CYA. A hypomorphic β-CYA allele was lethal at the embryonic level [149], whereas γ-CYA knock-out mice can survive to adulthood (for a review see [103]). Delayed embryonic development, as well as impaired cell growth and survival, was nevertheless observed in γ-CYA null mice [150]. Distinct localizations of β-CYA and γ-CYA mRNAs have been reported in myoblastic and fibroblastic cells [92], in osteoblasts [151], and in neurons [152]. γ-CYA mRNA was diffusely distributed in these cultured cells, while β-CYA mRNA localized to the leading edge of fibroblasts [153, 154]. The localization of β- and γ-CYA proteins in distinct subcellular compartments has been addressed by several studies, but with discordant results, likely because of variable experimental conditions and interpretations. The use of polyclonal antibodies against γ-CYA, not recognizing β-CYA, but cross-reacting with SM actins [155, 156], probably through the common N-terminus sequence AcEEE, could have affected the interpretation. Mesenchymal cells in culture, such as

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Fig. 1.7 β- and γ-CYAs segregation during mitosis. Dividing HaCaT cells were stained for β- (green), γ-CYA (red), and for DNA (blue, DRAQ5). β-CYA is enriched in the equatorial region during mitotic telophase (contractile ring), whereas γ-CYA only displays a submembranous staining

fibroblasts, can express α-SMA in addition to cytoplasmic actins. Thus, a highly selective criterion is required to establish antibody specificity. Carefully selected monoclonal antibodies (mAbs) highly specific for γ-CYA, that is, not recognizing β-CYA and SM actins, were used for cytoplasmic actin isoform distribution and functional investigations [111]. Whereas β-CYA mAbs were produced many years ago [110], the unavailability of a specific γ-CYA mAb has precluded the precise comparative appreciation of the sorting of the two isoforms. Besides, a lack of staining for β-CYA in stress fibers has designated this structure as being formed of γ-CYA filaments [157]. Even with validated antibodies, the lack of staining may be due to epitope unavailability. Specific cell fixation conditions appear to be mandatory to uncover the actin epitope (N-terminal sequence; [111, 112]). Using newly generated monoclonal antibodies to β- and γ-CYA, we have recently shown that the two isoforms have distinct patterns of organization: β-CYA predominates in stress fibers, circular bundles, contractile rings (Fig. 1.7), and at cell-cell contacts. Conversely, γ-CYA is mainly organized in cortical and lamellar networks. In accordance with our results, Ervasti’s group described a membrane localization of γ-CYA in muscle cells [116] and in stereocilia [158], suggesting that γ-CYA can have a membrane shape supportive function. Furthermore, selective moderate depletion of these isoforms revealed functional diversities of β- and γ-CYA [111]. β-CYA siRNA-treated cells are highly spread, displaying broad lamellipodia and decreased stress fiber organization; in contrast, γ-CYA siRNA-treated cells acquire a contractile phenotype with thick actin bundles and decreased lamellipodial structures. Moreover, as revealed by kymograph analysis, the leading edge of β- or γ-CYA-depleted fibroblasts differs from control during cell migration: β-CYA-depleted cells show smooth and persistent protrusions with reduced ruffling and retraction activity, whereas γ-CYA-depleted cells produce filopodia-like protrusions and cell edge contractions. Finally, long-timescale random migration assays reveal that depletion of β- or γ-CYA results in distinct effects on fibroblast locomotion. β-CYA-depleted cells display a more sustained directional migration with reduced speed compared to control, indicating that β-actin may play

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a role in cell body translocation and rear retraction. In contrast, permanently transfected myoblasts overexpressing β-CYA were shown to display increased areas of protrusion and increased cell locomotion [159]. Whether these apparently opposite results are due to the difference of cell types used and/or to the relative content in actin isoforms remains to be clarified. Furthermore, in our hands, depletion of γ-CYA results in lamellipodia inhibition and in impaired directional migration, suggesting a crucial role for γ-CYA in protrusive activities during cell migration. In conclusion, according to these results, β-CYA would play a preferential role in contractile activities whereas γ-CYA would mainly participate in the formation of a network necessary for cell shape flexibility and motile activity. Preferential association between specific actin isoforms and different ABPs would support the hypothesis about distinct microfilament populations within a cell; however, the interactions between actin isoforms and ABPs in networks and parallel arrays, as well as details of the molecular mechanisms of their regulation remain largely uncharacterized. GFP-actin-transfected cells could have been an alternative/complementary approach to immunostaining. However, keeping in mind that the acetylated N-terminus is crucial for the identity of actin isoforms, the addition of GFP at this extremity of the molecule would likely render the GFP-actin not isoform-specific. The alternative of fusing GFP to the C-terminus of actin, although giving better results [111, 160], is not fully satisfactory. Probably, both types of fusion would block protein folding. In addition, the size of the GFP protein would impede accessibility of ABPs at the Cand N-terminal domains. In conclusion, the subtlety of structural differences among actin isoforms has prevented several conventional methods for studies of intracellular protein localization and function, and therefore antibodies specific for actin isoforms should still in the future provide much important information.

1.6

Intracellular Peptide Delivery

Knock-out mice and siRNA are well-recognized approaches to decipher protein functions. In the case of actin isoforms, results are relatively difficult to interpret. Effectively, the depletion of one isoform, in a long-term (weeks/months) delay as in knock-out mice and, to a lesser extent, in a short-term (days) delay when using siRNA, results in a compensation of the deleted isoform by another isoform’s expression, without functional substitution. Keeping in mind the dynamic activity of actin polymerization, short-term (min/hour) approaches could be an alternative to investigate a specific isoactin organization function. Rho and Rac inhibitions have been shown to affect, respectively, β-CYA and γ-CYA organization. Another, even more direct, approach is based on the hypothesis that ABPs specific for individual isoforms would interact with the isoactin N-terminus. If so, the N-terminus peptide of one isoactin, after introduction into the cell, should trap the specific ABP and circumvent the functional ABP-isoactin association. The first investigation based on this hypothesis was conducted by delivering the α-SMA N-terminus peptide (AcEEED) into myofibroblasts.

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The α-SMA-positive myofibroblasts have been shown to exert a high contractile activity, in vitro, compared to fibroblasts, by measuring collagen gel contraction and wrinkle formation on a silicone substrate [161]. This property is absolutely crucial for the healing process, but becomes deleterious when out of control, such as in hypertrophic scars, fibrocontractive diseases, and stromal reactions to neoplasms [134]. The specific N-terminal domain Ac-EEED in the α-SMA protein plays a crucial role in: (1) α-SMA polymerization [162], (2) promoting high contractile activity of myofibroblasts [161] and (3) incorporating α-SMA into stress fibers [160]. These studies demonstrated that application of a membrane-permeable peptide, containing the Ac-EEED tetrapeptide as functional domain, leads to removal of α-SMA from stress fibers and reduces wound contraction when applied topically. Treatment with this peptide reverts myofibroblasts into nonfibrogenic fibroblasts [161, 163]. To investigate whether actin isoforms have specific functions during cardiac development and cardiomyocyte contractility, α-SMA and α-SKA expression (using siRNA) and organization (using peptide delivery) were hampered during embryonic stem cell differentiation toward cardiomyocytes. Three isoforms are sequentially expressed during cardiac development. α-SMA is first and transiently expressed, followed by α-SKA and finally α-CAA. The sequence of actin isoform expression displays a similar pattern in the in vitro model as in mouse heart embryogenesis. Treatment of embryoid bodies with the α-SMA-fusion peptide during a time window preceding spontaneous beating prevents proper cardiac sarcomyogenesis, while α-SKA-fusion peptide has no effect. The depletion of α-SMA in embryonic stem cells, using RNA interference, also affects cardiac differentiation. The application of both fusion peptides on beating embryoid bodies impairs frequency. These results suggest specific functions for actin isoforms in cardiogenesis and cardiomyocyte contractility. A larger panel of N-terminus isoactin fusion peptides (α-SMA, γ-SMA, β-CYA, γ-CYA, and α-SKA as control) has been used to investigate the inhibition of vascular smooth muscle contractility in a peptide-specific manner and in a stimulus-dependant manner [164]. A significant inhibition of contractility was observed with the two γ peptides in both phenylephrine (PE) and phorbol-induced contractions, whereas α-SMA was only effective on phorbol-induced contractions. This difference may be due to distinct mechanisms of contraction [164]. The peptide approach deserves further validations on various cell types to analyze isoactin functions.

1.7

Conclusions

Although actin is a small protein with no splice variants and only a few highly conserved and similar isoforms, it is the most abundant protein in most eukaryotic cells, and is involved in so many functions that it still remains relatively unexplored, in particular at the level of its different isoforms. Whether the presence of different isoactin patterns reflects functional differences or is simply the result of differential gene expression is, as yet, unknown. The most obvious differences among the three

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classes of isoforms: α, β, and γ, are changes in their electrostatic charge, which lead to altered isoelectric points that permitted classification of these isoforms even before the proteins’ primary structures were known. This difference in charge and the fact that the N-terminus, where most of the charges reside, protrudes from the filament surface suggest that functional differences might relate to docking of proteins, including motors, where the binding energy is in part dependent on charge-charge interactions. Biochemical studies of actin-ABPs interactions in vitro are limited because only α-SKA can be efficiently purified in large quantities. Therefore, most of the available biochemical data relate to ABPs interactions with α-SKA rather than with the non-muscle actins. In this chapter, we have discussed major progress due to the development of specific antibodies as well as the biochemical and molecular biological methods of protein depletion. Future experiments using classical methods, in particular immunodetection with the complete panel of specific isoactin antibodies, as well as new peptide technology, mass-spectroscopy, high-resolution laser confocal microscopy, correlated fluorescence microscopy and cryoelectron tomography, etc. will enable more quantitative and higher resolution studies of the molecular architecture of actin isoform-ABP complexes and their regulation and function in the cell. Another challenge will be the identification of the ABPs specific for each actin isoform. The understanding of actin isoform-specific functions would provide useful tools for the diagnosis and the treatment of various pathologies, such as cardiovascular, fibrocontractive, and oncological diseases. Acknowledgments The research in the authors’ laboratories is supported by SNF grant # 310030_125320 (CC, RA) and Scientific & Technological Cooperation Programme SwitzerlandRussia (CC, VD), Russian Foundation of Basic Investigation grant # 10–04-00227-a (VD) and US NIH GM083272 (PAJ). We thank David Slochower for help with Figs. 1.1 and 1.2.

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123. Hewett TE, Grupp IL, Grupp G, Robbins J (1994) Alpha-skeletal actin is associated with increased contractility in the mouse heart. Circ Res 74:740–746 124. Clement S, Stouffs M, Bettiol E, Kampf S, Krause KH, Chaponnier C, Jaconi M (2007) Expression and function of alpha-smooth muscle actin during embryonic-stem-cell-derived cardiomyocyte differentiation. J Cell Sci 120:229–238 125. Moll R, Holzhausen HJ, Mennel HD, Kuhn C, Baumann R, Taege C, Franke WW (2006) The cardiac isoform of alpha-actin in regenerating and atrophic skeletal muscle, myopathies and rhabdomyomatous tumors: an immunohistochemical study using monoclonal antibodies. Virchows Arch 449:175–191 126. Kumar A, Crawford K, Close L, Madison M, Lorenz J, Doetschman T, Pawlowski S, Duffy J, Neumann J, Robbins J, Boivin GP, O’Toole BA, Lessard JL (1997) Rescue of cardiac alpha-actin-deficient mice by enteric smooth muscle gamma-actin. Proc Natl Acad Sci U S A 94:4406–4411 127. Martin AF, Phillips RM, Kumar A, Crawford K, Abbas Z, Lessard JL, de Tombe P, Solaro RJ (2002) Ca(2+) activation and tension cost in myofilaments from mouse hearts ectopically expressing enteric gamma-actin. Am J Physiol Heart Circ Physiol 283:H642–649 128. Blau HM, Baltimore D (1991) Differentiation requires continuous regulation. J Cell Biol 112:781–783 129. Weintraub H, Tapscott SJ, Davis RL, Thayer MJ, Adam MA, Lassar AB, Miller AD (1989) Activation of muscle-specific genes in pigment, nerve, fat, liver, and fibroblast cell lines by forced expression of MyoD. Proc Natl Acad Sci U S A 86:5434–5438 130. Sassoon D, Lyons G, Wright WE, Lin V, Lassar A, Weintraub H, Buckingham M (1989) Expression of two myogenic regulatory factors myogenin and MyoD1 during mouse embryogenesis. Nature 341:303–307 131. Lassar AB, Buskin JN, Lockshon D, Davis RL, Apone S, Hauschka SD, Weintraub H (1989) MyoD is a sequence-specific DNA binding protein requiring a region of myc homology to bind to the muscle creatine kinase enhancer. Cell 58:823–831 132. Gabbiani G, Ryan GB, Majne G (1971) Presence of modified fibroblasts in granulation tissue and their possible role in wound contraction. Experientia 27:549–550 133. Majno G, Gabbiani G, Hirschel BJ, Ryan GB, Statkov PR (1971) Contraction of granulation tissue in vitro: similarity to smooth muscle. Science 173:548–550 134. Schurch W (1999) The myofibroblast in neoplasia. Curr Top Pathol 93:135–148 135. Tomasek JJ, Gabbiani G, Hinz B, Chaponnier C, Brown RA (2002) Myofibroblasts and mechano-regulation of connective tissue remodelling. Nat Rev Mol Cell Biol 3:349–363 136. Dugina V, Fontao L, Chaponnier C, Vasiliev J, Gabbiani G (2001) Focal adhesion features during myofibroblastic differentiation are controlled by intracellular and extracellular factors. J Cell Sci 114:3285–3296 137. Hinz B, Phan SH, Thannickal VJ, Galli A, Bochaton-Piallat ML, Gabbiani G (2007) The myofibroblast: one function, multiple origins. Am J Pathol 170:1807–1816 138. Hinz B, Gabbiani G (2010) Fibrosis: recent advances in myofibroblast biology and new therapeutic perspectives. F1000 Biol Rep 2:78 139. Orlandi A, Hao H, Ferlosio A, Clement S, Hirota S, Spagnoli LG, Gabbiani G, Chaponnier C (2009) Alpha actin isoforms expression in human and rat adult cardiac conduction system. Differentiation 77:360–368 140. Saborio JL, Segura M, Flores M, Garcia R, Palmer E (1979) Differential expression of gizzard actin genes during chick embryogenesis. J Biol Chem 254:11119–11125 141. Kuroda M (1985) Change of actin isomers during differentiation of smooth muscle. Biochim Biophys Acta 843:208–213 142. Hirai S, Hirabayashi T (1983) Developmental change of protein constituents in chicken gizzards. Dev Biol 97:483–493 143. Izant JG, Lazarides E (1977) Invariance and heterogeneity in the major structural and regulatory proteins of chick muscle cells revealed by two-dimensional gel electrophoresis. Proc Natl Acad Sci U S A 74:1450–1454

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Chapter 2

Microtubules Pavel Dráber and Eduarda Dráberová

Abstract Microtubules, assembled from heterodimers of α- and β-tubulin, are hollow tubes of about 25 nm in diameter, participating in essential cellular functions such as maintenance of cell shape, cell division, cell motility, and ordered intracellular transport. Tubulin dimers form protofilaments running lengthwise along the microtubule wall with the β-tubulin facing the microtubule plus end conferring a structural polarity. α- and β-Tubulins are highly conserved and consist of isotypes encoded by different genes. Numerous posttranslational modifications of tubulin subunits diversify the surfaces of microtubules and provide a mechanism for their functional specialization. A third member of the tubulin family, γ-tubulin, plays a role in microtubule nucleation. Microtubules display dynamic instability characterized by alternating phases of growth and shrinkage separated by catastrophe and rescue events. The dynamic nature of microtubules is dependent on many microtubule-regulatory proteins.

2.1

Introduction

Microtubules are cylindrical cytoskeletal polymers indispensable for many vital cellular activities such as maintenance of cell shape, division, migration, and ordered vesicle transport powered by motor proteins. They are also essential in organizing the spatial distribution of organelles in interphase cells. Microtubules can be organized into microtubule-based organelles with a specialized function, including the radial cytoplasmic network, axonemes, centrioles, midbodies during cytokinesis, and the mitotic/meiotic spindles. Singlet microtubules are the most ubiquitous form of the polymer, however, microtubules can also form doublets (in cilia) or triplets (in centrioles and basal bodies; [1, 2]). Microtubule structures appearing during the cell cycle are shown in Fig. 2.1. The basic building blocks of microtubules are heterodimers of globular α- and β-tubulin subunits. Tubulins are arranged in a head-to-tail fashion to form 13 protofilaments that constitute microtubules with outer diameter around 25 nm. Microtubules are thus inherently polar and contain two structurally distinct P. Dráber () · E. Dráberová Laboratory of Biology of Cytoskeleton, Institute of Molecular Genetics, Academy of Sciences of the Czech Republic, Vídeˇnská 1083, 142 20, Prague 4, Czech Republic e-mail: [email protected] M. Kavallaris (ed.), Cytoskeleton and Human Disease, DOI 10.1007/978-1-61779-788-0_2, © Springer Science+Business Media, LLC 2012

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Fig. 2.1 Changes in microtubules during the cell cycle. Microtubule structures (green) undergo marked morphological changes during the cell cycle. Human osteosarcoma cell U2OS in a interphase, b metaphase, c anaphase, and d telophase was stained for αβ-tubulin dimer with polyclonal antibody (green) and for γ-tubulin with monoclonal anti-γ-tubulin antibody TU-30 ([55]; red). DNA is stained blue. Scale bar, 10 µm

ends: a slow-growing minus end, exposing α-tubulin subunits, and a fast-growing plus end, exposing β-tubulin subunits [3, 4]. Typically in mammalian cells, the microtubule minus ends are stably anchored in microtubule-organizing centers (MTOC), whereas the plus ends are highly dynamic and switch between phases of growth and shrinkage. There are, however, exceptions to this organization. In dendrites of nerve cells, some microtubules are oriented with minus ends away from the cell body. Microtubules vary considerably in their stability. While microtubules that form the axonemes in cilia and flagella are stable, cytoplasmic microtubules turn over rapidly. Microtubule dynamics help remodel the microtubular network during the cell cycle. Although the structure of microtubules is conserved among various cell types, it can be adapted to highly divergent tasks by mechanisms that are not yet fully understood. Incorporation of alternative tubulin isotypes and posttranslational modification of tubulin subunits can regulate microtubule properties. Intracellular microtubule organization is further controlled by the distribution of nucleation sites and by the activity of microtubule-regulatory proteins. This chapter will focus on microtubule fundamentals, briefly reviewing tubulin and microtubule structure, microtubule dynamics, tubulin isotypes, microtubule nucleation, tubulin posttranslational modifications, and proteins that stabilize or destabilize microtubules.

2 Microtubules

2.2

31

Microtubule Structure

In mammals, tubulin heterodimers represent 3–4% of the total protein content in cells and reach up to 20% in brain. The secondary and tertiary structures of the α- and β-monomers are essentially identical, as expected from their identity of >40% over the entire sequence of above 445 amino acids of the sequence [5]. α- and β-Tubulins are globular proteins with a molecular weight approximately 55 kDa and isoelectric points between 5.2 and 5.8 [6]. Each monomer is formed by three sequential and functionally distinctive domains: the nucleotide-binding N-terminal domain, intermediate domain, and C-terminal domain whose C-terminal tail (CTT) part is exposed on the surface of microtubules [7]. Dimers of α- and β-tubulin are stable and rarely dissociate at the 10–20 µM concentration of tubulin found in cells. Each tubulin monomer binds one molecule of GTP, nonexchangeably in α-subunit (N-site) and exchangeably in β-subunit (E-site). Tubulin dimers also bind divalent cations. Within the microtubule, each tubulin heterodimer forms extensive noncovalent bonds with its neighbors. These bonds form longitudinally as well as laterally between dimers in a protofilament, linking adjacent protofilaments. The N-site is buried at the monomer-monomer interface within the dimer, explaining the nonexchangeability at that site. On the other hand, the nucleotide at the E-site is partially exposed on the surface of the dimer, allowing its exchange with the solution. The cylindrical and left-handed helical microtubule wall typically comprises 13 parallel protofilaments in vivo (Fig. 2.2a). Microtubules assembled from purified tubulin in vitro have a broad distribution of protofilament numbers centered on 14 [8]. A major advance step in understanding the microtubule functions is marked by the solution of its structure, based on docking the high-resolution structure of brain tubulin, studied by electron crystallography [5, 9], into lower-resolution microtubule maps imaged by electron cryomicroscopy [10, 11]. These studies have confirmed that tubulin dimers form a B lattice, where the main lateral contacts across protofilaments are between subunits of the same type (i.e., α-α, β-β). Most of the studied microtubules appear to have a seam along their length in which lateral contacts are reversed (i.e., α-β, β-α; [12]). This is due to 12-nm helical pitch in combination with the 8-nm longitudinal repeat between αβ-tubulin dimers (Fig. 2.2b). There are fenestrations of about 1.5 × 2 nm between the contact regions of protofilaments, thus giving direct access to the microtubule lumen [13].The microtubular surface displays a surprisingly large number of binding sites, with numerous proteins binding to the outside surface and a multitude of small ligands binding to the inside of microtubules [14]. Some structural interactions with other molecules including nucleotides, drugs, microtubule-associated proteins (MAPs), and motor proteins have been predicted [3].

2.3

Microtubule Dynamics

Assembly (polymerization) and disassembly (depolymerization) of microtubules is driven by the binding, hydrolysis, and exchange of GTP on the β-tubulin monomer. GTP hydrolysis is not required for microtubule assembly per se but is necessary

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Fig. 2.2 Microtubule structure and dynamics instability. a Microtubules are composed of stable αβ-tubulin heterodimers that are aligned in a polar head-to-tail fashion to form protofilaments. b The cylindrical and helical mcrotubule wall typically comprises 13 protofilaments in vivo. A discontinuity in the structure of the microtubule wall (lattice seam) is marked by red dashed line. c Dynamic instability of microtubules. Polymerization of microtubules is initiated from a pool of GTP-loaded tubulin subunits. GTP hydrolysis changes the conformation of a protofilament from a slightly curved tubulin-GTP to a more intensely curved tubulin-GDP structure. The curved tubulinGDP is forced to remain straight when it is part of the microtubule wall. Growing microtubule sheets presumably maintain the “cap” of tubulin-GTP subunits to stabilize the straight tubulin conformation within the microtubule lattice 1. Closure of the terminal sheet structure generates a metastable, blunt-ended microtubule intermediate 2, which might pause, undergo further growth or switch to the depolymerization phase. A shrinking microtubule is characterized by fountain-like arrays of ring and spiral protofilament structures 3. The polymerization–depolymerization cycle is completed by exchanging GDP of the disassembly products with GTP 4. Reprinted by permission from Macmillan Publishers Ltd: Nature Reviews in Molecular Biology (Akhmanova and Steinmetz 2008), copyright (2008)

for switching between alternating phases of growth and shrinkage separated by catastrophe (transition from growth to shrinkage) and rescue (transition from shortening to growth) events. Polymerization is typically initiated from a pool of GTP-loaded tubulin subunits (Fig. 2.2c 1; [15]). Growing microtubule ends fluctuate between slightly bent and straight protofilament sheets. GTP hydrolysis

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and release of inorganic phosphate occur shortly after incorporation, and is promoted by burial and locking of the partially exposed nucleotide as a result of the head-to-tail assembly of dimers. It has been postulated that GTP hydrolysis changes the conformation of protofilament from a slightly curved tubulin-GTP to a more profoundly curved tubulin-GDP structure [16]. This nucleotide-dependent conformational model predicts that the curved tubulin-GDP is forced to remain straight when it is part of the microtubule wall. Growing microtubule sheets maintain a “cap” of tubulin-GTP subunits to stabilize the straight tubulin conformation within the microtubule lattice [17]. A loss of this cap results in rapid depolymerization. A closure of the terminal sheet structure generates a metastable, blunt-ended microtubule intermediate (Fig. 2.2c 2), which may pause, undergo further growth or switch to the depolymerization phase. A shrinking microtubule is characterized by fountain-like arrays of ring and spiral protofilament structures (Fig. 2.2c 3). This conformational change, presumably directed by tubulin-GDP, may destabilize lateral contacts between adjacent protofilaments. The polymerization–depolymerization cycle is completed by exchanging GDP of the disassembly products with GTP (Fig. 2.2c 4). These characteristics result in dynamic instability [18], an essential feature of microtubules that allows them to search through the cell for targets, such as the chromosomal kinetochores, the cell cortex, and actin cytoskeleton [19, 20]. In some cells, microtubules that are not anchored to the centrosome undergo a special form of turnover called treadmilling. If the concentration of tubulin dimer exceeds a critical concentration (Cc ), the dimers polymerize into microtubules, whereas microtubules depolymerize at concentrations below the Cc . With differing concentrations at the opposite microtubule ends, and when the microtubules are at, or near, the steady state, higher Cc at the minus end results in shortening at this end, whereas lower Cc at the plus end of microtubule results in net growth [21]. When the overall concentration of soluble tubulin is maintained in between the two different Cc at opposite ends, continuous flow of subunits through microtubules will occur. One of the major sites of microtubule treadmilling is the mitotic spindle. In this situation the ends of microtubules, although tethered to kinetochores and spindle poles, remain free for subunit exchange and rapid flow of tubulin from plus to minus ends. This flow in spindle microtubules has been termed flux [22]. The stability and dynamics of microtubules are actively regulated by a number of cellular factors [19] as well as a variety of ligands, some of them with important anticancer properties. A host of well-known drugs (e.g., vinblastine, colchicine, and paclitaxel) potently suppress the dynamic instability and treadmilling dynamics of microtubules, and can thereby perturb cellular processes dependent on these dynamics [23, 24].

2.4 Tubulin Isotypes In mammals both α- and β-tubulin consist of isotypes encoded by different genes and differing in amino acid sequences. Alignment of amino acid sequences of the α- and β-tubulin isotypes revealed that most of the divergence is contained in the last

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Table 2.1 Overview of human tubulin isotypes Isotypea

Gene name

Protein ID (NCBI)

Length (amino acids)

C-terminal sequence (beyond residue 430)

α1A α1B α1C α4A α3C/D α3C/D α3E α8 α-like 3

TUBA1A TUBA1B TUBA1C TUBA4A TUBA3C TUBA3D TUBA3E TUBA8 TUBAL3

NP_006000 NP_006073 NP_116093 NP_005991 NP_005992 NP_525125 NP_997195 NP_061816 NP_079079

451 451 449 448 450 450 450 449 446

DYEEVGVDSVEGEGEEEGEEY DYEEVGVDSVEGEGEEEGEEY DYEEVGADSADGEDEGEEY DYEEVGIDSYEDEDEGEE DYEEVGVDSVEAEAEEGEEY DYEEVGVDSVEAEAEEGEEY DCEEVGVDSVEAEAEEGEEY DYEEVGTDSFEEENEGEEF DLAALERDYEEVAQSF

βI βII βII βIII βIVa βIVb βV βVI

TUBB TUBB2A TUBB2B TUBB3 TUBB4 TUBB2C TUBB6 TUBB1

NP_821133 NP_001060 NP_821080 NP_006077 NP_006078 NP_006079 NP_115914 NP_110400

444 445 445 450 444 445 446 451

EEEEDFGEEAEEEA DEQGEFEEEEGEDEA DEQGEFEEEEGEDEA EEEGEMYEDDEEESEAQGPK EEGEFEEEAEEEVA EEEGEFEEEAEEEVA NDGEEAFEDEEEEIDG VLEEDEEVTEEAEMEPEDKGH

a The nomenclature of α-tubulins follows the recent revision [27], and that of β-tubulins is based on recent reviews [28, 172]

20 amino acids [25], a region of the protein that lies on the exterior of the microtubule and is the putative binding site for MAPs [26]. Differences among isotypes are often highly conserved in evolution, suggesting that they have functional significance. In humans, eight α-tubulin and seven β-tubulin isotypes, were identified ([25, 27, 28]; see Table 2.1). In addition, other very different forms of tubulin have been discovered, designated as γ, δ, ε, ζ, η, θ, ι, and κ [29]. Interestingly, all have been found either in the centrosome or a very similar basal body. Some of these tubulins play a significant role in the assembly of these two organelles [30]. Together with α- and β-subunits, these tubulins constitute the tubulin superfamily. Several of them are widespread among eukaryotes (α, β, γ, δ), while other are more restricted [29, 30, 31, 32]. Different α- and β-tubulin isotypes often differ in their cellular and tissue distribution. Besides, purified isotypes display different properties including microtubule assembly, conformation, GTPase, dynamics, and ability to interact with antitumor drugs [24, 28]. Vertebrate β-tubulin isotypes have fairly distinct tissue distributions [25, 33]. βI-Tubulin expression is essentially ubiquitous. The distribution of βIVbtubulin is also widespread among tissues and cell types but is especially prominent in axonemes (cilia and flagella; [34]). βII-tubulin particularly abounds in brain, peripheral nerves, and muscles, but is also expressed to a lesser degree in other tissues. It has also been described in tumor cell nuclei of various cancer types [35, 36, 37]. βIII-Tubulin occurs largely in neurons and testicular Sertoli cells, and in low amounts in a very small number of other tissues. Specific expression of βIII-tubulin in neuronal cell is depicted in Fig. 2.3a, b. βIVa-Tubulin is expressed only in brain, while βVI-tubulin is restricted to hematopoietic-specific cell types (megakaryocytes and

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Fig. 2.3 Differential expression of βIII-tubulin and γ-tubulin. a, b Primary culture of rat neurons and glial cells stained for tubulin with polyclonal antibody (a green) and for neuron-specific βIIItubulin with monoclonal antibody TU-20 [173] (b red). Nuclei are shown in blue. c, d Comparison of γ-tubulin expression in primary culture of human astrocytes (c) and human glioblastoma cells T98G (d) using monoclonal antibody TU-30 [55]. Note that γ-tubulin in astrocyte is concentrated in centrosome, while in glioblastoma cells a high amount of γ-tubulin is also present in cytoplasm. Fluorescence images were captured and processed in exactly the same manner. Scale bars, 20 µm

platelets). The distribution of βV is still largely unknown. On the other hand, tissue distribution of α-tubulin isotypes seems to be much less complex when compared to β-tubulin. In many cases, the isotype distribution among tissues and even among different cell types within the same tissue is complex [38]. In addition, the pattern changes during development [39]. Interestingly, expression of tubulin isotypes is altered in drug-resistant and tumorous cells [40–42]. There is some evidence suggesting that cells alter the synthesis of certain tubulin isotypes in order to overcome drug exposure. Tubulin isotype expression profiling has been assessed by RT-PCR of mRNA [43, 44], specific antibodies [43] or by mass spectrometry [45]. While antibodies directed against C-terminal, isotype-defining sequences of most β-tubulin isotypes are readily available, antibodies to α-tubulin isotypes are scarce.

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βIII-Tubulin

Compared to other β-tubulin isotypes, βIII-tubulin possesses certain distinctive properties, which may account for its unique function(s) [28]. Unlike the βI, βII, and βIV isotypes, βIII-tubulin lacks in this regard the widely conserved and oxidationsensitive residue cys239, which is replaced by ser239 [46]. It has been hypothesized that the absence of cys239 may permit αβIII-tubulin dimers to assemble in the presence of free radicals [28]. In addition, the βIII-tubulin contains an uncommon cys124 residue, in contrast to other β-tubulin isotypes which share the ser/ala124 residue [46]. The remarkable phylogenetic conservation of βIII-tubulin across various vertebrate species indicates that cys124 and ser239 may have some functional roles. Unlike the βII- and βIV-tubulin isotypes, βIII-tubulin is phosphorylated at a serine in the C-terminus [47]. Also, in contrast to other β-tubulin isotypes, the presence of thr429 in βIII-tubulin strongly favors microtubule assembly [48]. When tubulin is reduced and carboxymethylated, βIII-tubulin shows a unique electrophoretic mobility on polyacrylamide gels [49]. Finally, despite its restricted and highly selective (predominantly neuronal) cell type distribution in normal organs and tissues, the βIII isotype is widely, albeit differentially, expressed in a broad range of human tumors of neuronal and nonneuronal origin. Abnormal βIII-tubulin expression is probably associated with more aggressive and drug-resistant cancers [33, 50, 51].

2.5

Microtubule Nucleation

One of the key components required for microtubule nucleation and stabilization is γ-tubulin [52], a highly conserved, albeit minor, member of the tubulin superfamily concentrated in interphase cells mainly in MTOCs [53]. There is about 30% identity between γ-tubulin, with molecular weight approximately 48 kDa, and tubulin dimers. In mitotic cells the γ-tubulin appears on spindle poles and it is also distributed along spindle fibers [54, 55]. During cytokinesis it is found in midbodies [56]. Localization of γ-tubulin in different phases of the cell cycle is shown in Fig. 2.1. The γ-tubulin is associated in complexes with other proteins. The human γ-tubulin small complex (γTuSC; around 280 kDa) comprises two molecules of γ-tubulin and one molecule each of GCP(γ-tubulin complex protein) 2 and 3 [57]. It has a form of Yshaped flexible structure with γ-tubulins located on the two arms [58, 59]. The large γ-tubulin-ring complex (γTuRC) derives from 5 to 7 γ-TuSCs by condensation and association with proteins GCP4, GCP5, GCP6 [60], and GCP-WD/NEDD1 [61]. Electron microscopic tomography indicates that the associated proteins absent in γTuSC form the cap of the ring structure [62]. Hundreds of γTuRC-like rings were found in pericentriolar material of centrosomes, and the presence of these rings correlated with the ability of centrosomes to nucleate microtubules [63]. Apart from the nucleation from MTOC, γTuRCs are also involved in the regulation of microtubule minus-end dynamics [64]. There is some evidence that γ-tubulin may associate with

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the microtubule wall [65, 66] and with cellular membranes [67–69] where it can participate in noncentrosomal microtubule nucleation [67, 70, 71]. γ-Tubulin complexes apparently also play a role in the regulation of microtubule plus-end dynamics and in the spindle assembly checkpoint signaling [72, 73]. Whereas multiple gene families encode α- and β-tubulin, only two functional genes exist in mammalian cells (TUBG1, TUBG2) that code very similar γ-tubulins [74]. The γ-tubulin is posttranslationally modified [66, 75, 76], and phosphorylation [77–79] as well as monoubiquitination [80] of γ-tubulin have been described. Complexes of γ-tubulin with protein tyrosine kinases of Src family [78, 81, 82], polo-like kinase [83], microtubule affinity-regulating kinase 4 (MARK 4; [84]) or phosphoinositide 3-kinase [71, 85, 86] have been documented. Collectively taken, the data strongly suggest that kinases might be involved in the regulation of γ-tubulin interactions. Increased γ-tubulin expression has been reported in cells of breast carcinoma [87, 88] and gliomas [89–91]. The γ-tubulin abnormalities in cancer cells can result in dysfunction of centrosomes due to the presence of supernumerary centrosomes, and/or in aberrant nucleation due to ectopic γ-tubulin localization [92]. Differential subcellular distribution of γ-tubulin in human astrocytes and glioblastoma cells is shown in Fig. 2.3c, d. Numerous proteins and protein complexes, including ninein, augmin, Cep192/SPD2, AKAP450/CG-NAP, pericentrin/kendrin, and CDK5RAP2/centrosomin contribute to the anchoring of γTuRC to MTOCs. Localization of these factors can be specific for a particular cell type or cell cycle stage [72, 93].

2.6 Tubulin Posttranslational Modifications High-resolution isoelectric focusing that separates polypeptides differing in their net charge has revealed that tubulin subunits can be resolved into more than 20 isoforms, far more than expected from the number of isotypes that are actually expressed [94, 95]. This fact reflects extensive posttranslational modifications (PTMs) of both tubulin subunits. Most PTMs of tubulin subunits take place after polymerization into microtubules and modified tubulins are nonuniformly distributed along microtubules. Strongly modified stable microtubules are concentrated in specialized organelles, such as centrioles and cilia or in axons of neurons. Currently known PTMs are summarized in Table 2.2. Well-characterized PTMs include acetylation, detyrosination, polyglutamylation, and polyglycylation.

2.6.1

Tubulin Acetylation

Acetylation of the ε-amino group of lys40 of α-tubulin [96, 97] is the sole modification that occurs on the amino acid moiety that extends to microtubule lumen [5]. The acetyl-lys40 of α-tubulin is therefore obscured in unfixed cells [98]. Several

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Table 2.2 Overview of tubulin posttranslational modifications Modificationa Subunit Residue(s) Forward enzyme Acetylation Detyrosination 2 Glutamylation Glycylation Phosphorylation

α α α α, β α, β α, β

K40 C-terminal Y Penultimate E Multiple E in CTT Multiple G in CTT S172 β-tubulin

Glycosylation Arginylation Methylation on K Methylation on E Palmitoylation Sumoylation Ubiquitylation Nitration on Y

α, β α, β α α, β α, β α α α

E434 α-tubulin C376 α-tubulin Multiple K

MEC-17 CCP1

Reverse enzyme HDAC6, SIRT2 TTL

TTLL1, 4, 5, 6, 7, 9, 11, 13 CCP5 TTLL3, 8, 10 Cdk1 PSK, Syk, Fes

Parkin

a

See text for references CTT C-terminal tail

additional acetylation sites on α-tubulin and one on β-tubulin were found by mass spectrometry, and some of them are exposed on the outer surface of microtubules [99]. However, lys40 of α-tubulin seems to be the main acetylation site [100]. The acetyllys40 mark appears with some delay after microtubule assembly and is an indicator of the microtubule age [101, 102]. MEC-17 protein related to GCN5 histone acetyltransferases was identified as the acetyltransferase that exclusively acetylates lys40 of α-tubulin [103]. Deacetylation of acetyl-lys40 on α-tubulin is catalyzed by HDAC6 [104, 105] and SIRT2 [106] deacetylases. Acetylation is a characteristic feature of stable microtubules.

2.6.2

Tubulin Detyrosination and Generation of 2-Tubulin

The reversible detyrosination/tyrosination of α-tubulin is the best characterized tubulin modification [107]. Detyrosination removes the gene-encoded C-terminal tyrosine residue on α-tubulin after incorporation of tubulin dimer into microtubule [108, 109]. Detyrosination is likely to be generated by the cytosolic carboxypeptidase 1 (CCP1), since mutation of this enzyme in mice substantially decreases the level of microtubule detyrosination [110]. It remains to be determined whether or not CCP1 has α-tubulin detyrosination activity in vitro. Retyrosination of tubulin heterodimers released from microtubules is generated by tubulin tyrosine ligase (TTL) [111]. Detyrosinated α-tubulin can be further converted into 2-tubulin by irreversible removal of penultimate glutamate residue [112]. This PTM can limit the amount of α-tubulin that undergoes recycling because 2-tubulin released from microtubules cannot revert to an unmodified state. Detyrosinated α-tubulin is characteristic of stable microtubules and 2-tubulin is found on very stable microtubules.

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39

Tubulin Glutamylation and Glycylation

Polyglutamylation [113] and polyglycylation [114] are two related posttranslational polymeric modifications (polymodifications) that involve the attachment of polypeptide side chains made of glutamates and glycines to specific glutamate residues in the CTT (C-terminal tail) of α- and β-tubulin. Glycylation is restricted to cilia and flagella in mammalian cells [115], whereas polyglutamylation is a more widespread modification. Polymodifications are catalyzed by a large family of tubulin tyrosine ligase-like proteins (TTLL) that show homology with TTL [100]. TTLL enzymes have characteristic specifities, displaying preference for α- or β-tubulin, and for the generation of short or long side chains [116]. Generation of polymodification side chains is based on two biochemically distinct reactions: initiation and elongation. First, a glutamate or glycine residue is attached to the γ-carboxyl group of the acceptor glutamate and second, additional glutamate or glycine residues are added via isopeptide bond. A competition exists between the two polymodifications because the glutamic acid- and glycine-ligases attach side chains to the same glutamate in CTTs [117]. Cytosolic carboxypeptidase-like protein 5 (CCP5) was identified as the tubulin deglutamylase [118].

2.6.4

Tubulin Phosphorylation

Phosphorylation of serine residues within the CTT of β-tubulin in microtubules has been reported, although the enzymes responsible for this modification have not been identified [119–121]. Tyrosine residue in the CTT of α-tubulin can be phosphorylated by the nonreceptor tyrosin kinase Syk [122, 123]. Outside the CTT region, the β-tubulin can be phosphorylated at ser172 by Cdk1/cyclin B complex that regulates the entry into mitosis. This phosphorylation occurs on the unpolymerized tubulin dimer and inhibits its incorporation into assembling microtubules [124]. Tubulin can be phosphorylated in vitro by serine/threonine-protein kinase PSK [125] or nonreceptor tyrosine kinase Fes [126], but the in vivo relevance and modification sites are unknown. Phosphorylation seems to be a minor modification for microtubules. The reason might be that tubulin itself is a highly anionic protein and the introduction of a negatively charged phosphate group might not be sufficient to change the structure or charge of the protein. On the other hand, polyglutamylation introduces the formation of bulky side chains and multiple charged residues, and provides thereby a sufficient structural change on microtubules [127].

2.6.5 Additional Tubulin Modifications Tubulin is ubiquitylated on the α-subunit [128]. It was proposed that the acetyl-lys40 α-tubulin is targeted for degradation through ubiquitylation, presumably following the disassembly of acetylated microtubules [129]. Palmitoylation of tubulin can

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explain the association of tubulin with membranes [130, 131]. Glycosylation of the α-and β-tubulin by sialyloligosaccharides [132] as well as arginylation [133] of the two tubulin subunits has been reported. Moreover, lysine methylation on α-subunit [134], methylation on glutamic acid in both tubulin subunits [135], sumoylation of α-tubulin [136], and tyrosine nitration of α-tubulin [137] were documented.

2.6.6

Tubulin Code

Diverse PTMs are proposed to form the biochemical “tubulin code” that can be read by factors interacting with microtubules. Specific microtubule regions can be distinguished biochemically and functionally by the presence of PTMs on tubulins [2, 127]. It appears that modifications could participate in targeting the molecular motors and MAPs to defined subsets of microtubules inside a cell. Microtubules in various subcellular compartments can be functionally distinguished by means of divergent PTM. Subsets of microtubules that orient toward the leading edge are more stable and richer in acetylation and detyrosination [138]. Similarly, axons, dendrites, synapses, and growth cones in neurons contain different PTMs on microtubules [139]. Tubulin detyrosination affects the dynamics of the plus end of microtubules by inhibiting the binding of microtubule plus-end tracking proteins (+TIPs) comprising a Cytoskeleton-Associated Protein Glycine-rich (CAP-Gly) microtubule-binding domain. These proteins localize to the ends of tyrosinated microtubules but not to the ends of detyrosinated microtubules [140]. Among the PTMs, polyglutamylation has the highest potential for generation of complex signals in microtubules by varying the: (1) tubulin subunit specificity (α- versus β-tubulin), (2) side chain length, and (3) modification sites on the tubulin molecule [141]. The extent to which microtubules become polyglutamylated might affect the binding of positively charged MAPs that are attached along microtubules as MAP2 and tau. On the other hand, glutamylation can also have a microtubule-destabilizing effect through the regulation of microtubule-severing proteins, such as katanin and spastin. Hyperglutamylated microtubules are more sensitive to spastin-mediated severing in vitro [142]. Microtubule-stabilizing and destabilizing activities of tubulin glutamylation can be regulated by the length and position of the glutamylate side chain and the availability of microtubule effectors [100]. It has been reported that some kinesin motor proteins associate preferentially with microtubules that are enriched with acetylation and detyrosination. This property makes it possible for kinesin motors to segregate membrane trafficking events between stable and dynamic microtubule populations [143].

2.7

Microtubule-regulatory Proteins

Dynamic remodeling of microtubules is essential for many cellular processes such as cell division, migration, and differentiation. Many proteins interact with microtubules within the cell and are involved in basic functions like microtubule growth,

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stabilization, destabilization, and interactions with other cellular organelles. The mechanochemical ATPases kinesins and dyneins (microtubule motor proteins)—not discussed in this chapter in detail—use microtubules as pathways for intracellular transport. A wide variety of microtubule-regulatory proteins provide the functional diversity of microtubules [93, 144, 145]. Regulation can occur on many levels, including regulation of tubulin monomer folding through the action of tubulin-folding factors or microtubule nucleation already dealt within Sect. 2.5. The following section will focus on proteins that are primarily involved in regulating microtubule stability and dynamics.

2.7.1

Microtubule-Stabilizing MAPs

MAPs of the MAP2/tau family include proteins MAP2(A, B, C), MAP4, and tau. The MAP2 and tau are found in neurons, whereas MAP4 is present in many other tissues but is generally absent in neurons. All three members of the family have alternative splice forms; all isoforms share a conserved carboxy-terminal domain containing 18 residue microtubule-binding repeats and an amino-terminal projection domain of varying size. These structual MAPs are thermostable and contain three or four microtubule-binding repeats separated by 13 or 14 residue inter-repeat regions. Each repeat binds to a tubulin subunit along the length of a single protofilament. Microtubule binding is regulated by phosphorylation at the KXGS motif within the repeats, and the phosphorylated proteins dissociate from microtubules [146]. The projection domains are capable of cross-linking with membranes, other microtubules, or intermediate filaments [147]. All members of the family are best known for their microtubule-stabilizing activity and for their proposed roles in the regulation of microtubule networks in axons and dendrites of neurons. Accumulating evidence suggests various other functions, such as binding to microfilaments, recruitment of signaling proteins, and regulation of microtubule-mediated transport. Data from studies of motor proteins moving along the tau-decorated microtubules revealed that dynein tends to reverse the direction, whereas kinesin rather shows the tendency to detach at patches of bound tau. The differential modulation of dynein and kinesin motility suggests that MAPs can spatially regulate the balance of microtubuledependent axonal transport [148]. Tau is also implicated in Alzheimer’s disease and other dementias, since hyperphosphorylation promotes the aggregation of tau into paired helical filaments found in these neurodegenerative diseases [149]. The ability of MAP2 to interact with both microtubules and microfilaments might be crucial for neuromorphogenic processes, such as neurite initiation, where microtubules and microfilaments become reorganized in a coordinated manner. Various upstream kinases and interacting proteins have been identified that regulate the microtubule-stabilizing activity of MAP2/tau family proteins [150]. Similarly, proteins of the MAP1 family, that include MAP1A, MAP1B, and MAP1S, belong to the microtubule-stabilizing MAPs and bind along the microtubule lattice. MAP1A and MAP1B are predominantly expressed in neurons, where they supposedly play an important role in the formation and development of axons

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and dendrites. After their initial expression, MAP1A and MAP1B polypeptides are cleaved into light and heavy chains, which are then assembled into mature complexes together with the separately encoded light chain 3 subunit (LC3). Both MAP1A and MAP1B are well known for their microtubule-stabilizing activity, but MAP1 proteins can also interact with other cellular components. Furthermore, the activity of MAP1A and MAP1B is controlled by upstream signaling mechanisms, including the MAP kinase and glycogen synthase kinase 3β pathways [151]. A variety of other proteins stabilize microtubules in distinct ways. Stable tubulin only proteins (STOPs) bind to microtubules and make them resistant to depolymerization by cold. STOPs are expressed in both neuronal and nonneuronal cell types; a lack of these proteins is associated with defects in synapse function [152]. Tektins cross-link and stabilize axoneme microtubules in cilia and flagella [153].

2.7.2

Microtubule-Severing Proteins

Microtubule-severing proteins cut the existing microtubules, thus creating new minus and plus ends. The generation of short microtubule fragments by the action of these proteins is important for reorganization of the microtubule cytoskeleton without the necessity of complete microtubule disassembly [154]. Katanin is an AAA protein (ATPases Associated with various cellular Activities) engaged in mitosis, neuronal differentiation, and flagellar physiology. It is a heterodimer with a 60-kDa catalytic subunit and a regulatory 80-kDa subunit involved in subcellular targeting. The presence of microtubules stimulates the katanin ATPase activity, resulting in disassembly at different positions along the microtubule. Katanin is effective in form of a hexameric ring attached to the microtubule lattice. Spastin, another member of the AAA family, also severes microtubules. Its importance is strenghtened by the fact that mutations of the spastin gene are the major cause of hereditary spastic paraplegia. Fidgetin and VPS4 are another katanin-related microtubule-severing proteins [155].

2.7.3

Microtubule-Disassembly Promoters

Apart from severing, microtubule disassembly is promoted by two additional mechanisms; sequestration of tubulin dimers, and by activity of microtubule depolymerases. Stathmin family proteins are small proteins that bind to tubulin dimers and lower the pool of free tubulin available for polymerization, thus decreasing the microtubule growth rate and increasing the frequency of spontaneous depolymerization. Binding of two tubulin subunits to stathmin results in the formation of ternary tubulin-stathmin T2 S complex. Other proteins of the stathmin family can also bind two tubulin heterodimers through their stathmin-like domain (SLD; [156]). Moreover, stathmin can destabilize microtubules by increasing the switching frequency (catastrophe frequency) from growth to shortening at plus and minus ends through binding directly to microtubules [157]. The activity of stathmin is downregulated by multisite phosphorylation in response to a number of cellular signals. Inhibition of stathmin by

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mitotic kinases promotes the assembly of mitotitc spindle. Interestingly, many malignant cells express unusually high levels of stathmin (hence its synonym Op18, for oncoprotein 18). Its overexpression correlates with abnormal cell motility and tissue invasion of human sarcomas in vivo [158]. Two classes of kinesins, kinesin-13 (kinesin with internal motor domain; Mitotic Centromere-Associated Kinesin; MCAK) and kinesin-8 (KIF18A; Kip3 in yeast) bind directly to microtubule ends and use ATP hydrolysis to catalytically depolymerize microtubules. They represent microtubule depolymerases which bind to and stabilize the bent protofilament conformation, a structural intermediate during depolymerization. MCAK, though it is a kinesin, does not move in a directed manner on the microtubule lattice. On the other hand, Kip3 is the most processive motor protein discovered so far; it moves on average 12 microns before falling off [159].

2.7.4

Microtubule-Assembly Promoters

The XMAP215/Dis1/TOG family of proteins associate with microtubule plus ends and accelerate growth, perhaps by reducing the dissociation of incoming tubulin dimers and stabilizing the GTP cap [159]. These microtubule polymerases contain TOG domains (named so after their discovery in the protein ch-TOG; Colonic, Hepatic Tumor Overexpressed Gene). The TOG domain has about 250 amino acids made up of six pairs of parallel α-helices (HEAT repeats) folding into a paddlelike structure. Tandemly arranged TOG domains mediate binding to tubulin and are probably responsible for microtubule growth-promoting activity of these proteins [160]. XMAP215 (detected in frog) is a processive polymerase that accelerates growth tenfold. There are two ways in which XMAP215 might catalyze polymerization. XMAP215 could stimulate polymerization either by acting as an adaptor that brings multiple tubulins to the growing end or by changing the structure of the growing end so that soluble tubulin can bind more easily. Structurally, XMAP215 is a long thin molecule that could potentially bind multiple tubulin heterodimers and form small curved protofilaments. High-resolution tracking of microtubule growth in vitro shows that in the presence of XMAP215 the microtubule growth is saltatory increasing the length in steps of up to 60 nm [161].

2.7.5

Microtubule Plus-End Tracking Proteins

+TIPS are specialized MAPs that are conserved and specifically accumulate at growing microtubule plus ends. +TIPs represent a structurally and functionally diverse group of proteins and typically target the growing, but not shrinking, microtubule ends. +TIPs interact with microtubule ends and with each other to regulate microtubule dynamics. Moreover, +TIP networks are also profoundly involved in coordinating complex aspects of cell architecture. At present, more than 20 different +TIP families have been identified. +TIPs are usually multidomain and/or multisubunit

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Fig. 2.4 Intracellular distribution of plus-end tracking protein EB1. a Human osteosarcoma cells U2OS were fixed and stained with antibodies for EB1 (green) and tubulin (red). Nuclei are shown in blue. b–d Higher magnification of the region marked in (a). Note the distribution of EB1 at distal ends of microtubules. Scale bars, 10 µm

proteins that range in size from a few hundred up to thousands of residues [15, 162]. Although +TIPs have a common ability to track growing microtubules in vivo, they do not have a common microtubule-end-binding domain; furtheromore, the molecular mechanisms by which +TIPs recognize microtubule ends differ. +TIPs can be classified into several classes according to their sequence homologies and the domains and/or mechanisms involved in microtubule tracking [163]. MCAK and TOG proteins, discussed in the previous section, also belong to +TIPs. End-binding (EB) family proteins contain a highly conserved N-terminal domain that adopts a calponin homology (CH) fold and is responsible for microtubule binding. The C terminus of EB proteins harbors an α-helical coiled-coil domain that mediates parallel dimerization of EB monomers. It further comprises the unique EB homology (EBH) domain and an acidic tail encompassing a C-terminal EEY/F motif, resembling those of α-tubulin and Cytoplasmic Linker Protein of 170 kDa (CLIP-170). Both the EBH domain and the EEY/F motif enable the EB proteins to physically interact with an array of +TIPs to recruit them to microtubule ends [164]. EB proteins are generally accepted to represent core components of +TIP networks because they autonomously track growing microtubule plus ends independent of any binding partners. EB proteins usually promote microtubule dynamics and growth, and suppress catastrophes in cells [165]. Distribution of EB1 in interphase U2OS cells is shown in Fig. 2.4. The first reported +TIP was CLIP-170 [166] that contains CAP-Gly domains, small globular modules that contain a unique conserved hydrophobic cavity, and several characteristic glycine residues. CAP-Gly domains use their hydrophobic cavity

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to confer interactions with microtubules and EB proteins by specifically recognizing the C-terminal EEY/F sequence motifs. Other prominent examples of CAP-Gly proteins are CLIP-115 and the large subunit of dynactin complex p150glued . Mammalian CLIPs represent anticatostrophe factors and promote microtubule rescue [162]. The largest group of +TIPs comprises large and complex, often multidomain, proteins containing low-complexity sequence regions that are rich in basic, serine and proline (basic-S/P) residues. They share a small four-residue motif Ser-x-Ile-Pro (SxIP, where x denotes any amino acid), which is specifically recognized by the EBH domain of EB proteins [167]. Noteworthy representants of this diverse class of +TIPs are the Adenomatous Polyposis Coli (APC) tumor suppressor, MicrotubuleActin Cross-linking Factor (MACF), and Stromal Interaction Molecule 1 (STIM1). SxIP motif is present also on motor protein MCAK and CLIP-Associated Protein (CLASP) that belong, respectively, to mirotubule depolymerases and to TOG family proteins. APC, MACF, and CLASP are involved in stabilization of microtubules and interaction of microtubules with various cellular structures [163]. STIM1 participates in the extension of endoplasmic reticulum together with growing microtubule ends [168], and participates in remodeling of microtubule organization [169]. +TIPs also promote organization of specialized microtubule arrays, such as mitotic spindles [170]. Interestingly, many +TIPs accumulate at centrosomes and other microtubule organizing centers where they might take part in microtubule nucleation and anchoring [171]. Acknowledgments This work was supported in part by Grants 204/09/1777 and P302/101/1701 from the GA CR, Grant LC545 from Ministry of Education, Youth and Sports of the Czech Republic and by the Institutional Research Support (AVOZ 50520514).

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145. Wade RH (2009) On and around microtubules: an overview. Mol Biotechnol 43:177–191 146. Ozer RS, Halpain S (2000) Phosphorylation-dependent localization of microtubule-associated protein MAP2c to the actin cytoskeleton. Mol Biol Cell 11:3573–3587 147. Chapin SJ, Bulinski JC (1992) Microtubule stabilization by assembly-promoting microtubuleassociated proteins: a repeat performance. Cell Motil Cytoskeleton 23:236–243 148. Dixit R, Ross JL, Goldman YE, Holzbaur EL (2008) Differential regulation of dynein and kinesin motor proteins by tau. Science 319:1086–1089 149. Baas PW, Qiang L (2005) Neuronal microtubules: when the MAP is the roadblock. Trends Cell Biol 15:183–187 150. Dehmelt L, Halpain S (2005) The MAP2/Tau family of microtubule-associated proteins. Genome Biol 6:204 151. Halpain S, Dehmelt L (2006) The MAP1 family of microtubule-associated proteins. Genome Biol 7:224 152. Bosc C, Andrieux A, Job D (2003) STOP proteins. Biochemistry 42:12125–12132 153. Amos LA (2008) The tektin family of microtubule-stabilizing proteins. Genome Biol 9:229 154. Zhang D, Rogers GC, Buster DW, Sharp DJ (2007) Three microtubule severing enzymes contribute to the “Pacman-flux” machinery that moves chromosomes. J Cell Biol 177:231–242 155. Roll-Mecak A, McNally FJ (2010) Microtubule-severing enzymes. Curr Opin Cell Biol 22:96–103 156. Cassimeris L (2002) The oncoprotein 18/stathmin family of microtubule destabilizers. Curr Opin Cell Biol 14:18–24 157. Steinmetz MO (2007) Structure and thermodynamics of the tubulin-stathmin interaction. J Struct Biol 158:137–147 158. Baldassarre G, Belletti B, Nicoloso MS, Schiappacassi M, Vecchione A, Spessotto P, Morrione A, Canzonieri V, Colombatti A (2005) p27Kip1 -stathmin interaction influences sarcoma cell migration and invasion. Cancer Cell 7:51–63 159. Howard J, Hyman AA (2007) Microtubule polymerases and depolymerases. Curr Opin Cell Biol 19:31–35 160. Slep KC (2009) The role of TOG domains in microtubule plus end dynamics. Biochem Soc Trans 37:1002–1006 161. Kerssemakers JW, Munteanu EL, Laan L, Noetzel TL, Janson ME, Dogterom M (2006) Assembly dynamics of microtubules at molecular resolution. Nature 442:709–712 162. Galjart N (2010) Plus-end-tracking proteins and their interactions at microtubule ends. Curr Biol 20:R528-R537 163. Akhmanova A, Steinmetz MO (2010) Microtubule +TIPs at a glance. J Cell Sci 123:3415– 3419 164. Slep KC (2010) Structural and mechanistic insights into microtubule end-binding proteins. Curr Opin Cell Biol 22:88–95 165. Komarova Y, De Groot CO, Grigoriev I, Gouveia SM, Munteanu EL, Schober JM, Honnappa S, Buey RM, Hoogenraad CC, Dogterom M, Borisy GG, Steinmetz MO, Akhmanova A (2009) Mammalian end binding proteins control persistent microtubule growth. J Cell Biol 184:691–706 166. Perez F, Diamantopoulos GS, Stalder R, Kreis TE (1999) CLIP-170 highlights growing microtubule ends in vivo. Cell 96:517–527 167. Honnappa S, Gouveia SM, Weisbrich A, Damberger FF, Bhavesh NS, Jawhari H, Grigoriev I, van Rijssel FJ, Buey RM, Lawera A, Jelesarov I, Winkler FK, Wuthrich K, Akhmanova A, Steinmetz MO (2009) An EB1-binding motif acts as a microtubule tip localization signal. Cell 138:366–376 168. Grigoriev I, Gouveia SM, van der Vaart B, Demmers J, Smyth JT, Honnappa S, Splinter D, Steinmetz MO, Putney JW, Jr., Hoogenraad CC, Akhmanova A (2008) STIM1 is a MT-plusend-tracking protein involved in remodeling of the ER. Curr Biol 18:177–182 169. Hájková Z, Bugajev V, Dráberová E, Vinopal S, Dráberová L, Janáˇcek J, Dráber Pe, Dráber P (2011) STIM1-directed reorganization of microtubules in activated mast cells. J Immunol 186:913–923

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170. Goshima G, Nedelec F, Vale RD (2005) Mechanisms for focusing mitotic spindle poles by minus end-directed motor proteins. J Cell Biol 171:229–240 171. Bettencourt-Dias M, Glover DM (2007) Centrosome biogenesis and function: centrosomics brings new understanding. Nat Rev Mol Cell Biol 8:451–463 172. Verdier-Pinard P, Pasquier E, Xiao H, Burd B, Villard C, Lafitte D, Miller LM, Angeletti RH, Horwitz SB, Braguer D (2009) Tubulin proteomics: towards breaking the code. Anal Biochem 384:197–206 173. Dráberová E, Lukáˇs Z, Ivanyi D, Viklický V, Dráber P (1998) Expression of class III β-tubulin in normal and neoplastic human tissues. Histochem Cell Biol 109:231–239

Chapter 3

The Kinesin Superfamily Linda Wordeman

Abstract The mammalian genome possesses 45 unique genes that code for kinesins. Kinesins are motor molecules, ATPases, which are specialized for the transport of cellular materials along the surface of cellular microtubules. Microtubules consist of linear polymers of repeating 8-nm-long tubulin dimers, each of which comprises one binding site for the kinesin motor domain. Kinesins “walk” from one binding site to the next, hydrolyzing one ATP with every step. In addition to their transport roles, these enzymes also remodel microtubules, engineer mitotic spindle assembly, and assist with chromosome segregation in dividing cells. Thus far, kinesins have been identified to operate in every conceivable microtubule-based process in the cell. Their diversity has enabled researchers to study disparate microtubule-based processes in isolation by selective disruption of individual kinesin motors. Functional characterization of microtubule-dependent activities with such high precision would not be possible using microtubule drugs, most of which globally disrupt all microtubule processes simultaneously. For this reason, kinesins have recently become attractive targets for the development of chemotherapeutic drugs.

3.1

Introduction

The kinesin superfamily consists of 45 unique genes coding for motor proteins that use the energy from ATP hydrolysis to translocate along microtubules [1]. The highly conserved core motor domain of kinesins consists of an approximately 340 amino acid globular domain, possessing a diagnostic ATP-binding domain that is structurally quite similar to myosins and small G-proteins (Fig. 3.1a; [2]). Hydrolysis of ATP results in the motor cycling between low- and zero-affinity and high-affinity binding states to the microtubule. The majority of kinesins are dimeric and they utilize this dimeric structure to “walk” along the surface of the microtubules by coordinately alternating the motor that is tightly bound to the surface of the microtubule (Fig. 3.1b). This property makes this class of proteins superbly adapted for directed long-distance transport of materials along neuronal tracts [3]. Kinesins also L. Wordeman () Department of Physiology and Biophysics, University of Washington, 1705 NE Pacific St., 98195-7290 Seattle, WA, USA e-mail: [email protected]

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Fig. 3.1 Structure and mechanism of kinesin motors. a The structure of the kinesin motor domain resembles the small g-proteins. Switch I, yellow; switch II, red. Kinesin motor is oriented with the microtubule-binding side facing front. b Dimeric motile kinesins coordinate motor domain binding with ATP hydrolysis to translocate unidirectionally along the tubulin lattice. “T,” ATP; “D,” ADP; “O,” no ATP bound. c Depolymerizing kinesins diffuse on the surface of the microtubule to reach the end. At the end the motors bind strongly to the terminal dimer in the ATP state, hydrolysis leads to the release of the detached tubulin dimer. “T,” ATP; “T*,” ATP transition state; “D,” ADP, “?”, identity of the nucleotide-bound state during diffusion is controversial

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use the same mechanism to transport materials over short distances in small cells, sort materials within cells in a microtubule-dependent manner, and also to move and reorganize the microtubules themselves as occurs during mitotic spindle formation. To accommodate these diverse activities the motors are considerably less well conserved outside of the 340 amino acid motor domain. However, the 45 unique kinesin genes can be classified into 14 (or perhaps 15, the exact number is controversial) functional families based on sequence conservation both inside and outside of the motor domain [3, 4]. Functionally, these family-specific domains adapt family members to associate with certain cargos, localize to particular regions of the cell, translocate in different directions, or with distinct kinetics along the microtubule. In the case of the family of microtubule-depolymerizing kinesins, the motor domain stabilizes a curved tubulin conformation that results in detachment of the tubulin dimer from the microtubule (Fig. 3.1c; [5]). The functional and structural diversity of the kinesin superfamily has led to a concerted effort to identify small molecule chemotherapeutic inhibitors to individual kinesins. Such inhibitors could be more specifically targeted to dividing cells than existing inhibitors such as paclitaxel or vinblastine, which is broadly specific for microtubules in all tissues and whose administration leads to side effects associated with disruptions in axonal transport. In contrast to microtubules, a number of mitotic kinesins are not utilized, or even expressed, in postmitotic differentiated tissue. The search for small molecule inhibitors targeting these kinesins is greatly facilitated by thorough investigations of kinesin function and regulation. The goal of this review is to present a summary of kinesin mechanism of action with an emphasis on how the mechanistic specializations of some kinesins lead to functional specialization and human diseases.

3.2

Nomenclature

Forty-five unique kinesin genes have been identified at specific chromosomal loci in humans and mice [1]. As many of these genes are conserved between organisms, kinesins have been classified based on sequence homology both within and outside of the 340 amino acid kinesin core motor domain. Based on this categorization scheme, kinesins have been broadly classified into 14 kinesin families [4] consisting of consensus monophyletic groups conserved among published phylogenetic analyses [6–10]. These families are summarized briefly in Table 3.1. Individual members of a specific family may have distinct roles in either mitosis or interphase cells or both. In general the broad function will depend upon the enzymatic activity, or what in the motor field is more commonly described as the “motility” of the motor. Most kinesins are plus-end-directed motile (or “walking”) motors. This refers to the fact that microtubules are intrinsically polar molecules. The end-to-end assembly of dimers of alpha- and beta-tubulin occurs linearly to create a polymeric tube that consists of one end in which beta-tubulins are exposed (traditionally called the “plus-end”) and one end in which alpha-tubulins are exposed (traditionally called

Kif4, XKLP1 Kif11, Eg5, BimC Kif20, Kif23, MKLP1/2 CENP-E Kif18A/B, Kip3 Kif6, Kif9 Nod, Kid, Kif22 Kif26A/B, Smy1

Dimer Bipolar tetramer [97] Dimer Dimer Dimer Unknown Unknown Unknown Unknown Dimer Dimer

Plus-end motilityb [92, 93]

Plus-end motility [96] Plus-end motility [99] Plus-end motility Plus-end motilityb [62, 63, 65] Unknown Plus-end motilityc [92, 109] MT binding [112]

Plus-end motilityb [116, 117]

Depolymerizer [55, 121]

Minus-end motilityb [32, 122, 123]

Kinesin-4

Kinesin-5 Kinesin-6 Kinesin-7 Kinesin-8 Kinesin-9 Kinesin-10 Kinesin-11

Kinesin-12

Kinesin-13

Kinesin-14

a

H heavy chains (containing the ATPase domain), L light chains, KAP kinesin-associated protein Molecule interconverts between monomer and dimer b Also reported to modulate microtubule polymerization c Nod may lack motility [128, 129]

NCD, Kar3, KifC1/2, Kif25

Kif2A/B/C, MCAK

Kif12, Kif15, Xklp2

Kif1, Kif13, Kif14, Unc104, Kif16

Monomer/dimera [37, 88]

Kinesin-3

Common names Kinesin heavy chain, Unc116

Plus-end motility

Plus-end motility [84]

Kinesin-2

Structure 2H, 2L; Heterotetramer Kif3A/B/C, Kif17

Plus-end motility [30, 31]

Kinesin-1 2H, KAP; Heterotrimer

Activity

Family

Table 3.1 Summary of 14 kinesin families Cellular functions Intracellular transport, signaling [80–83] Intracellular/flagellar transport, sensory cells [46, 85–87] Intracellular/axonal transport, mitosis/cytokinesis [89–91] Mitosis/congression, spindle assembly [94, 95] Mitosis/spindle assembly [98] Mitosis/cytokinesis [100–102] Mitosis/congression [103, 104] Mitosis/congression [64, 105] Ciliar/flagellar transport [106–108] Mitosis/congression [110, 111] Secretion [113, 114], signal transduction [112], cell adhesion [115] Mitosis/spindle assembly [79, 118–120] Chromosome segregation, neurite extension [16] Intracellular transport [124, 125], mitosis/spindle assembly [126, 127]

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the “minus-end”). Each 8-nm-long tubulin dimer represents one binding site for a kinesin motor. The classic names for the respective ends of the microtubule have accordingly worked their way into the nomenclature for kinesin activity because these motors hydrolyze ATP to translocate either unidirectionally toward the plus-end or toward the minus-end. Recent studies have shown that the kinesin motor domain can also be specialized to exploit ATP hydrolysis to depolymerize microtubules, to modulate microtubule assembly, and finally to bind and release from microtubules in the absence of motility. The activities associated with the 14 kinesin families are conserved across species and, for many kinesin families, so are their key cellular functions. For example, the kinesin-5 family consists of plus-end-directed motile kinesins possessing a bipolar tetrameric structure that enables them to simultaneously translocate on two antiparallel microtubules. This activity slides the two microtubules apart and establishes an outward force for the assembly of the bipolar mitotic spindle [11]. In every species from yeast to mammals this family performs this same key role [12]. The individual kinesin family functions are subject to increased diversity via splice variants [13], which will lead, in some cases, to the same motor transporting different cargos in different tissues. Alternatively, kinesins from the same family may utilize a similar activity for different purposes in different tissues. The kinesin13 member MCAK/Kif2C, which is not present in terminally differentiated tissue, uses microtubule-depolymerizing activity to facilitate microtubule turnover in mitotic spindles and correct errors in chromosome attachment [14, 15]. Another member of the same family, Kif2A, uses the same microtubule disassembly activity to control collateral branch extension during neural development [16]. Thus, the activities of the 14 kinesin families are conserved although their cellular functions may not, superficially, appear to be conserved.

3.3

Structure and Activity

Members of the kinesin superfamily can be identified by an approximately 360 amino acid globular domain possessing a phosphate-binding (p-loop) motif characteristic of the nucleotide-binding pocket of small G-proteins. High-resolution structural studies have revealed considerable structural and enzymatic similarity between small G-proteins, kinesins, and myosins [17]. An evolutionary relationship between small G-proteins (the ancestor) and myosins and kinesins is predicted, despite the lack of sequence conservation, because they possess spatial conservation between core structural elements: switch I, switch II, and the p-loop ([2]; Fig. 3.1a). Kinesins hydrolyze one ATP for each 8-nm step along the surface of the microtubule [18, 19]. During this regime, the switch II helix mediates the interaction between the kinesin motor with the tubulin dimer and communicates the ATP hydrolysis state. A flexible linker attaching the motor domain to the rest of the molecule docks against the motor upon ATP binding, an interaction that is thought to promote strain within the dimeric molecule. This interaction serves to fling the rear

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head forward where it can encounter another binding site further down the lattice (Fig. 3.1b). This is the basis for the “hand-over-hand” mechanism of translocation utilized by motile kinesins [20]. The majority of kinesins are dimers (Table 3.1). This quaternary structure facilitates motility by alternating ATP hydrolysis and microtubule binding in a mechanism not fully understood, to ensure that one head will bind to the microtubule while the other head releases. In this manner, some highly processive kinesins can take hundreds of 8-nm steps along the surface of the microtubule without falling off. Similar to myosins, high processivity is not required for all cellular functions. Some kinesins, such as those of the kinesin-14 family are not very processive as single molecules [21], however, can succeed as effective minus-end-directed transporters when operating in groups [22], when bound to other microtubule-interacting proteins [23] or on bundles of microtubules [24]. The cycle of binding and release from tubulin coupled toATP hydrolysis is adapted in the kinesin-13 family to facilitate microtubule disassembly (Fig. 3.1b). Many microtubule-binding proteins, including motors, will exhibit unbiased, bidirectional diffusion when associated with microtubule lattice [25]. True to form, the kinesin-13 family members will associate with microtubule lattice and diffuse along it in the presence of bound nucleotide, however, without nucleotide hydrolysis. Binding of ATP, similar to motile kinesins, converts the motor into a high-affinity microtubule-bound state with the caveat that the preferred structure is to a curved microtubule protofilament ([26, 27]; such as would be seen in depolymerizing microtubules [28, 29]). The motor and its associated tubulin dimer then break away from the microtubule polymer and the hydrolysis of ATP leads to the release of the motor from the detached tubulin dimer in a manner very analogous to the motile kinesins’ release of one head (Fig. 3.1c). Although kinesin-13s appear to be dimers, it is unclear if the heads are coordinated as is seen with motile kinesins.

3.4

Kinesin Regulation

Kinesins participate in all microtubule-based functions within the cell. These functions can be broadly divided into two areas: intracellular transport and cell division. The duties of kinesins in intracellular transport are complex, overlapping and, in some instances, incompletely characterized (see [3] for an excellent review). Historically, the kinesin-1 family was originally identified and biochemically purified as a microtubule-dependent ATPase that was correlated with anterograde fast axonal transport [30, 31]. This was the first plus-end-directed microtubule motor to be identified and was clearly the fast axonal transport motor. The second kinesin to be identified was a kinesin-14 family member identified by genetic means by its participation in cell division [32, 33]. These subsequent studies demonstrated that kinesins are likely to be a family of related microtubule-dependent ATPases that participate in a variety of cellular processes including cell division. A general rule of thumb for many microtubule motors is that their motility on microtubules is often subject to intramolecular autoinhibition that is relieved by cargo

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Fig. 3.2 Autoinhibition is a common feature of kinesins. a Kinesin-1 family members are autoinhibited by the C-terminal tail (yellow) until the light chains (blue) associate with cargo freeing the motor to interact with the microtubule. b Kinesin-2 family members exhibit two-stage inhibition of motility mediated by the C-terminal tail domain (yellow)

binding (or putative posttranslational modifications). Kinesin-1 has been studied extensively in this respect. The two light chains of the heterotetramer regulate the autoinhibition that the C-terminus of the motor-containing subunits confers on the microtubule-binding activity of the motor domain ([34]; illustrated in Fig. 3.2a). Recently, similar autoinhibition has been documented for a number of kinesin-related proteins [35–39]. The kinesin-3 family member Unc104, which has been implicated in the longdistance transport of synaptic vesicle precursors, exhibits an unusual switch in quaternary structure between a monomeric version, which is incompetent for rapid, long-distance transport to a dimeric, highly processive rapid motor for axonal transport. The transition from monomer to dimer can be triggered by a local increase in the concentration of the motor on cargo. In contrast the mammalian orthologue for this protein, Kif1A, is expressed as an inactive dimer. The basis for this inactivity is twofold: both the interaction with microtubules and the processivity are differentially autoinhibited by distinct nonmotor regions of the molecule. Thus, similar to kinesin-1, rapid, long-distance motility in mammalian Kif1A is triggered by cargo binding. A similar method of regulation has also been reported for Kif17, a kinesin-2 family member important for ciliary transport (Fig. 3.2b).

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Precise mapping of the domains responsible for robust motility is important because these domains are key regions that are regulated by cellular second messengers and cellular binding partners. For example, Kif17 is a ciliary transport motor important for vertebrate photoreceptor development and maintenance and channel targeting in the cilia of olfactory sensory neurons. This motor is negatively regulated by two different domains: the extreme C-terminal tail and a coiled-coil domain within the tail (Fig. 3.2b). Thus, by extension, at least two binding partners might be required for full activation of motor activity enabling finer cellular control of the motor. In fact, it has been shown that importin-B is required for Kif17 to enter the base of the cilium where RanGTP is then required for full activation of motor activity. It is likely that all kinesins will exhibit some form of intramolecular autoregulation that is controlled by phosphorylation, other posttranslational modifications and/or interaction with cellular binding partners.

3.5 3.5.1

Kinesins and Disease Transport

It is becoming increasingly evident that disruptions in the axonal transport leads to peripheral neuropathies. Charcot-Marie Tooth (CMT) disease is one of the most common peripheral neuropathies and can result from myelinopathies, derived from defects in Schwann cells (CMT1), or axonopathies, derived from defects leading to dying back of the peripheral neuronal axons (CMT2). One family afflicted with axonopathic Charcot-Marie Tooth disease (CMT2A) has genetically mapped to a point mutation in Kif1B (a kinesin-3 family member) that abolished the ATPase activity of the motor [40]. kinesin-2 member. Neuropathies can also result from mutations in the transport machinery that does not specifically target motors. For example, loss of mitofusin, an outer mitochondrial membrane protein that couples mitochondrial transport to kinesins via the Miro-Milton complex also leads to CMT2A [41]. Ironically, while kinesins are often considered in the context of long-range rapid axonal transport, they are also employed in nonneuronal cells for all manner of subcellular transport jobs including membrane remodeling and vesicle transport. Thus, it is not surprising to discover that a myelinopathic CMT resulting from the loss of myotubularin-related protein, similarly uncouples a scaffold for membrane remodeling from Kif13B (a kinesin-3 family member) in Schwann cells [42]. Recent studies suggest that abnormal accumulations of amyloid beta (Aβ) protein, a histopathological characteristic of Alzheimer’s syndrome, leads to increased phosphorylation of the light chains of kinesin-1 resulting in the uncoupling between kinesin-1 and its cargo [43]. This disruption of kinesin-dependent fast axonal transport is likely to contribute to the neuronal dysfunction and death seen in Alzheimer’s patients. Mouse and fly models have shown that any disruption in the smooth flow of axonal traffic, particularly in large peripheral neurons, leads to aggregates of cytoskeletal proteins analogous to a traffic jam on a large freeway [44, 45]. Such

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aggregates, which are hallmarks of many neuropathic diseases such as Alzheimer’s and Amyotrophic lateral sclerosis (ALS), further exacerbate proper neuronal function leading, eventually, to death of the neuron and/or its processes. Although many people are familiar with the role of the microtubule motor protein dynein in ciliar and flagellar movement, kinesins are also critically important for ciliar function. In this case, they serve a transport function, moving rafts of essential proteins and vesicular proteins to the distal tip of the cilum [46]. Dynein, which is a minus-end-directed motor, transports material back to the base of the cilium in addition to its role in ciliar and flagellar bending. Ciliar dysfunction leads to a vast spectrum of disorders such as kidney disease, disruption of cellular signaling, and developmental malformations. Kinesin-2 family members, which are essential for ciliary assembly and maintenance, are often implicated in ciliar disease. For example, the loss of either Kif3A or Kif17 (both members of the kinesin-2 family) will lead to defects in ciliogenesis and signaling [47, 48]. Of particular interest, in this respect, is the function and maintenance of the primary cilium. This nonmotile cilium that protrudes from the surface of most mammalian cells, integrates signaling responses, mediates mechanotransduction in the renal tubule, influences embryonic pattern formation, and mediates signal transduction in the growth and maintenance of stem cell populations. Experimental disruption of Kif3A in mice leads to randomization of the left-right axis (a ciliopathic effect also known as situs inversus) and ultimate embryonic lethality from a number of other problems. However, when the loss of Kif3A was restricted to renal tubular epithelial cells, the defect was limited to polycystic kidney disease [48, 49]. In contrast, global disruption of Kif17 primarily affected photoreceptor development in zebrafish embryos [47]. An explanation for the tissue-specific effects of the two kinesin-2 family members, both of which are important for ciliogenesis, is that the function of Kif3A is essential in the proximal region of the cilium whereas Kif17 manifests more activity in the distal region of the cilium. The outer segments of rods and cones are specializations of the distal region of the primary cilium. Thus, as Kif17 is required for transport more specifically in the outer segments of the rods and cones, the loss of Kif17 manifests a distinct phenotype relative to Kif3A.

3.5.2

Cell Division

Kinesins are equally well-known for their essential roles in cell division and ensuring accurate chromosome segregation as they are for intracellular transport. Mitotic kinesins have been touted as excellent targets for anticancer therapies because, unlike general microtubule-targeting drugs such as paclitaxel and vinblastine, drugs targeting mitotic kinesins would be limited to dividing cells, avoiding peripheral neuropathies commonly associated with drugs that broadly disrupt microtubules. All of the kinesin families possess members that either directly participate in cell division or participate in growth control and apoptotic processes. Thus, there are numerous kinesins with potential utility as markers for tumor progression or as therapeutic targets for anticancer drugs (reviewed in [50, 51]).

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Cell Division: Microtubule Dynamics

Dynamic microtubules, capable of rapid remodeling, are essential for mitotic spindle assembly and function, cell motility, and neuronal growth cone advancement. Dynamic microtubules are so important for cell division and mitotic spindle function that suppression of microtubule dynamics leads to cell cycle arrest and apoptosis. Thus, drugs such as paclitaxel, that suppress microtubule dynamics, have proven to be highly efficacious for the elimination of dividing cancer cells [52]. Less appreciated is the role of microtubule dynamics in metastasis and vascularization. Drugs that suppress microtubule dynamics have also proven effective in controlling metastasis and expansion of the microvasculature [53]. For these reasons, kinesins that regulate microtubule polymerization and depolymerization are both important biomarkers and key targets for cancer therapy that have the potential to increase the clinical arsenal of therapeutic targets beyond microtubules [54]. Members of the kinesin-13 family, especially MCAK/Kif2C, are potent regulators of microtubule dynamics. As described above, they use the energy from ATP hydrolysis to disassemble microtubules from both ends of the polymer [55]. MCAK/Kif2C appears to be maintained particularly in dividing cells and discarded upon differentiation [56]. In cells, MCAK/Kif2C increases microtubule turnover in both interphase and mitosis, which facilitates migration and chromosome movement and suppresses erroneous microtubule attachments during cell division [14, 57]. MCAK/Kif2C is overexpressed in a variety of tumors and high MCAK/Kif2C levels are correlated with a poor prognosis for recovery [58, 59]. Suppression of MCAK expression has the potential to limit tumor growth in breast cancer and may be a useful early predictor of tumor metastasis as the protein has been detected in the peripheral bloodstream of colon cancer patients [60]. Another family of kinesins, the kinesin-8 family, has a key role in regulating microtubule dynamics. Kinesin-8 family members are highly processive, plus-end-directed motile kinesins that can “walk” for many hundreds of steps along a microtubule without detaching. Although implicated in disassembling microtubules [61–63] this family of kinesins has a still controversial effect once it reaches the microtubule end, perhaps involving suppression of MT dynamics [63–65] rather than overt disassembly as is seen with the kinesin-13 family members. The kinesin-8 family member Kif18A is essential for proper congression of chromosomes to the metaphase plate. Data mining of human breast cancer microarrays suggests that, like MCAK, overexpression of Kif18A is correlated with invasive breast cancer and a poor prognosis for survival. Furthermore, suppression of Kif18A expression inhibited cell division and metastatic potential in breast cancer cell lines [66]. Study of this family of kinesins is still at an early stage, however, it is likely to be a tantalizing target for anticancer drug development. Cellular overexpression of kinesins that do not overtly regulate microtubule dynamics have been demonstrated to influence the ratio between microtubule dimers and polymer. While the mechanism by which this occurs is not understood, a recent study has shown that upregulation of kinesins, even nonmitotic, kinesins, that increased the pool of free tubulin resulted in breast cancer cell lines becoming more

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resistant to docetaxel [67]. This study underscores the important influence that the intrinsic homeostasis of tubulin subunits with microtubule polymer has on treatment outcomes from therapies utilizing anticancer drugs that target microtubules.

3.5.2.2

Cell Division: Mitotic Spindle Function

The majority of kinesin superfamily members play important roles in the assembly and operation of the mitotic spindle. However, an individual kinesin family may include a number of unique genes or alternatively spliced variants that operate exclusively in either transport or cell division. This tissue-specific and functional diversity makes mitotic kinesins prime candidates for anticancer drugs that have the potential to avoid neuropathies associated with broad spectrum drugs targeting microtubules (reviewed in [50]). Targeting mitotic kinesins will also be efficacious for the development of multidrug combinations, second-line treatment of relapses, and molecular targeted therapy. The greatest progress along these lines has been in developing small molecule inhibitors of the kinesin-5 family member Kif11/Eg5. Monastrol, the first specific inhibitor of a mitotic kinesin that was active in cells, was identified in 1999 [68]. Since that time, more potent and selective Monastrol derivatives [69] and other chemically diverse compounds (reviewed in [70]) have been identified that specifically inhibit Kif11. Ispinesib, the first kinesin-5 family inhibitor to make it to phase I and phase II clinical trials, has exhibited acceptable toxicity levels both singly and in combination with docetaxel [71], and demonstrated an ability to permeate the blood-brain barrier [72]. Animal and cell mouse models show antitumor activity associated with Ispinesib, albeit at high drug levels [73, 74]. Collectively, these studies suggest that the further chemical optimization of kinesin-5 family inhibitors and the screening and development of other mitotic kinesin inhibitors is a promising line of investigation toward increasing the clinical arsenal of cancer therapies. Data mining of expression profiles of normal and tumorigenic tissues have revealed altered expression profiles in a number of mitotic kinesins, pointing the way toward more potential therapeutic targets. The kinesin-5 family may have led the way because the mitotic monasters formed in cell culture when kif11/Eg5 is inhibited are strikingly obvious in phenotype-based screens. However, altered expression of several mitotic kinesins, such as KID/Kif22 (a kinesin-10 family member; [75]), Kif14 (a kinesin-3 family member; [76]), Kif4A (a kinesin-4 family member; [77]) and CENP-E (a kinesin-7 family member; [78]), has been correlated with tumorigenesis. In addition, a relatively understudied kinesin, Kif15 (a kinesin-12 family member; [79]) has recently been attributed with mitotic functions. Its role in cell division was only discovered when another key mitotic kinesin was impaired. Creative development of functionally informative combinatorial screens is likely to identify more mitotic kinesins in the future. Such kinesins are likely to impact chromosomal instability during tumorigenesis even if they appear, superficially, to contribute redundantly to cell division.

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3.6

Conclusion

Over 45 unique kinesin genes have been identified since the first kinesin was biochemically purified from the cytoplasm of the giant axon of the squid. This diversity is even greater if one considers the numerous tissue-specific splice variants that have been identified. In addition to their roles as intracellular transporters of vesicles and protein complexes along microtubules, kinesins are also intimately involved in the assembly and function of cilia, flagella, and the mitotic spindle during cell division. By extension from these diverse microtubule-dependent cellular activities, deregulation of kinesin function has been implicated in peripheral neuropathies, Alzheimer’s disease, kidney disease, and cancer. The kinesin superfamily, with its tissue specificity and functional specialization, represents a rich area for the development of small molecule inhibitors. An immediate goal for the future is to develop functional screens to identify more classes of kinesin inhibitors. Chemical refinement of identified inhibitors is significantly advanced by a precise data on kinesins’ mechanism of action, kinetics, and regulation. Single-molecule studies of kinesins in vitro identify key domains regulating kinesin function that might be discovered no other way, however, by systematic molecular dissection and sensitive kinetic assays. Reductionistic studies of kinesin mechanism provide insight into the activities that the motors contribute to cellular functions. Such studies also aid in the development of sensitive functional screens for inhibitors (or activators). History has shown that even those compounds that do not make it to phase-III clinical trials usually end up being of great use to the scientific community as functional probes in model systems. Molecular probes directed at kinesin family members will substantially increase the arsenal of clinical therapies available to patients for some of the world’s most intractable diseases. Acknowledgments I am indebted to Mike Wagenbach for preparing Fig. 3.1a. The research in Linda Wordeman’s laboratory is supported by the National Institutes of Health (GM069429) and the National Science Foundation (MCB-1041173).

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Chapter 4

Basics of the Cytoskeleton: Myosins Omar A. Quintero, Judy E. Moore and Christopher M. Yengo

Abstract The myosin superfamily of molecular motors plays essential roles in a wide variety of cellular processes by virtue of their ability to generate force and motion through an ATP-driven cyclic interaction with actin filaments. We provide an overview of the structure, function, and biophysical properties that are common to most characterized myosins and also include examples of how myosins are adapted to perform specific cellular functions. Since many myosins are implicated in disease conditions, a complete understanding of their cellular roles and biophysical properties is critical for developing treatments for these diseases.

4.1 The Human Myosin Superfamily Approximately 40 myosin genes representing 12 classes have been identified in the human genome [1–4]. Two of those genes appear to be pseudogenes that are not translated [5, 6]. The two classes containing the most myosin genes are the class I and class II myosins with 8 and 15 genes, respectively. All other classes contain no more than three members, and class VI, X, XVI, and XIX contain a single gene in the genome. A thorough class-by-class review of the individual myosin classes found in humans can be found in the book, Myosins: A Superfamily of Molecular Motors [7]. Figure 4.1 shows an unrooted phylogenetic tree generated from amino acid sequence alignment of the motor domains of the 40 human myosin genes and indicates their Human Genome Organization (HUGO) gene name. As the names of multiple myosin genes have been revised, correlation of data in the literature can often become complicated. Multiple articles exist that can serve as guides to the recent changes in the myosin nomenclature ([1, 4, 8]; Fig. 4.1). C. M. Yengo () Department of Cellular and Molecular Physiology, The Penn State College of Medicine, 500 University Dr, Hershey, PA 17033, USA e-mail: [email protected] O. A. Quintero Department of Biology, University of Richmond, Richmond, VA 23173, USA J. E. Moore Department of Biology, University of North Carolina Charlotte, Charlotte, NC 28223, USA

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Fig. 4.1 An unrooted phylogenetic tree of the myosin superfamily in humans. The 38 myosin genes and 2 pseudogenes (boxed name, dashed line) found in the human genome, identified in [4] were aligned based on their core myosin motor domains, as identified by Pfam (http://pfam.janelia. org/) [281]. The alignment was generated using ClustalX2 [282] with the default settings, and the tree representing the topology of relationship between myosin genes was generated using Figtree 1.3.1 (http://tree.bio.ed.ac.uk/software/figtree/). Genes are represented by their official HUGO human gene names (http://www.genenames.org/). Black dots represent nodes validated by bootstrapping values of greater than 90%. This tree is an update of the tree found in [1]

There is some conservation in the overall arrangement of domains within the human myosin superfamily. With a few exceptions, the conserved myosin motor domain is found at the N-terminus. Class III, IX, XVI, and XVIII myosins have kinase domains [9], Ras association domains [10], ankyrin repeats [11], and PDZ domains [1] N-terminal of their motor domains, respectively. In addition, a number of myosins have an SH3-like domain at the N-terminus of the motor domain [3]. The motor domain is followed by a light-chain-binding region that is thought to function as a lever arm [12], amplifying the conformational change associated with the power stroke [13, 14]. The lever arm/neck region contains a variable number of IQ motifs [15], depending upon the class of myosin, that are thought to provide increased rigidity to the alpha-helical neck upon light chain binding. The IQ motifs bind EF hand proteins such as myosin essential and regulatory light chains [15, 16], calmodulin [17], and calmodulin-like protein [18]. The interactions of these proteins allow for myosin activity that is calcium-dependent. In some instances, light chain phosphorylation also regulates myosin activity [19], and some light chains may interact with actin [20].

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Although the members of the myosin superfamily share a conserved ATPhydrolyzing motor domain, there is a considerable amount of variability in the C-terminal tail region. In many cases the tail region of myosins determines the other molecules with which the motor protein can interact. In the case of class II, V, VI, VII, IX, X, XV, and XVIII, portions of the tail domain contain regions that allow dimerization of the myosin via coiled-coil motifs [3]. Other domains mediate interactions with membrane lipids [21–26], microtubules [27, 28], actin filaments [29], integrins [30], small GTPases [31, 32], and other proteins. The enzymatic properties of the motor domain coupled with the biochemical interactions of the neck region and tail domain determine the cellular roles of each myosin. Although myosins were originally identified as components of the contractile machinery in muscle, the vast majority of human myosin genes have been shown to serve a wide variety of cellular functions. This chapter highlights some cellular roles for myosins in humans, but is not an exhaustive description of all myosin functions.

4.2

Molecular Mechanism of Motor-Based Force Production

Over the past century, research has advanced our understanding of how motor proteins such as myosin convert the energy from the hydrolysis ofATP into directed movement through a cyclic interaction with actin filaments. Understanding this mechanism requires high-resolution structural data of a molecular motor in different stages of its ATPase cycle both bound and unbound from its track. In addition, a complete understanding of the kinetics and thermodynamics of each of the steps in the ATPase cycle must be elucidated. Much of what is known about the structure and function of myosin motors was originally studied in muscle myosins while over the past few decades studies of nonmuscle myosins have greatly contributed to our understanding of the chemomechanical cycle of these fascinating force generators. The rotating cross-bridge model as a molecular description of myosin-actin interactions in muscle contraction began to emerge in the 1950s [33–36] and was based on analysis of interference and phase-contrast microscopy images. Globular heads of myosin molecules form the cross-bridges that were proposed to function as independent force generators by binding, rotating, and dissociating from the actin filaments with which they overlap. These observations and inferences remain the foundation for our understanding of actomyosin motor activity, not only for muscle myosins but for the entire myosin superfamily. AF Huxley proposed that myosin generates force by relaxation of a strained elastic element within the motor during the power stroke [37, 38]. The Lymn-Taylor scheme (Fig. 4.2) describes the ATPase cycle of the myosin cross-bridge which contains four distinct nucleotide-bound states [39] and can be classified as either the “weak” (ATP and ADP-Pi) or “strong” (ADP and nucleotide-free) actin-binding states. The swinging lever arm hypothesis explains the large conformational changes seen at the cross-bridge as resulting from amplification of small changes at the motor’s nucleotide-binding region (reviewed in [40]). The motor region of a myosin molecule converts the chemical energy released by ATP hydrolysis into mechanical energy as it undergoes cyclical interactions with an actin filament, first binding ATP then hydrolyzing it and sequentially releasing the

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Fig. 4.2 Diagram of the actomyosin (AM) contractile cycle. Force generation occurs by movement of the lever arm while myosin is attached to the actin filament and the recovery stroke occurs while it is detached from actin. The cycling between strong and weak binding states is mediated by nucleotide-dependent conformational changes in the actin-binding region. The kinetics, thermodynamics, and specific conformational changes of each step have not been completely elucidated. (We thank Anja Swenson for help in making this figure)

products (Fig. 4.2). Evidence seems to most strongly support that the biochemical steps are tightly coupled with mechanical/conformational stages so that each individual power stroke is the result of the hydrolysis of one ATP molecule, although this characteristic has been widely debated (reviewed in [41]). ATP binding (step 1) to nucleotide-free actomyosin (AM) greatly reduces myosin’s affinity for actin and causes rapid detachment of the cross-bridge (M.ATP) from the actin filament. ATP hydrolysis (step 2) is accomplished during the detached state or weakly bound state (M.ADP.Pi), but the hydrolysis products Pi and ADP remain linked to the motor until rebinding to actin (step 3). The release of phosphate (step 4) is associated with a transition from weak to strong actin binding as well as a conformational change that swings the lever arm, the force-generating power stroke phase. The release of ADP from actomyosin occurs in two steps, with the first step associated with an isomerization from a strong (AM.ADPs ) to a weak (AM.ADPw ) ADP affinity state (step 5). The subsequent ADP release (step 6) results in formation of the nucleotide-free or rigor state. Binding of another ATP molecule to rigor myosin releases the motor from the actin filament and begins another ATPase cycle. The crystal structures of myosin in combination with cryoelectron microscopy and image reconstruction of the actomyosin complex [12, 42–64] provide a framework

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Fig. 4.3 Crystal structure of skeletal muscle myosin II [61] modeled in the near rigor form [283] with the functional domains and motifs highlighted (adapted from [284]). The lever arm is formed by a long C-terminal alpha-helix (light blue) that is stabilized by associated light chains (gray). The actin-binding cleft separates the 50-kDa domain into upper (purple) and lower (white) domains. The nucleotide-binding pocket is formed by the N-terminal (blue) and the upper 50-kDa domains. The switch II region (orange), switch I and P-loop (not shown) coordinate nucleotide binding and hydrolysis. The relay helix/loop (yellow) communicates conformational changes from the nucleotide-binding pocket to the SH1 helix (red) which then alters the position of the lever arm through interactions with the converter domain (green). (See text for further details. Image rendered using PyMOL [285])

for understanding the mechanism of the power stroke driven by changes in the orientation of the lever arm with respect to the actin filament. Major structural elements of the myosin motor are highly strongly conserved, with subtle differences between the classes that have been identified from high-resolution crystal structures (Fig. 4.3). Four functional domains—the N-terminal domain, upper 50-kDa domain, lower 50kDa domain, and converter/lever arm, and four connectors—the SH1 helix, rigid relay, switch II, and the strut, make up the motor’s structure, along with various surface loops. The motor’s transducer region is recognized as the core of the motor, a sevenstranded beta sheet whose deformations during key steps in the cycle are proposed to play a role in subdomain coupling [49]. Between the N-terminal and the upper 50-kDa domain is the nucleotide-binding pocket, which orients the nucleotide with the gamma phosphate coordinated deep in the pocket. Several flexible loops (“switches”) associated with the ATP-binding pocket respond to ATP binding, hydrolysis, and product release by changing conformation, and thus are proposed to be critical for transmitting information to other regions of the motor. A large cleft that separates the upper and lower 50-kDa subdomains is more open in the weak binding states and more closed in the strong binding states suggesting this cleft is critical for mediating actin affinity. The converter domain is thought to amplify conformational changes from the nucleotide-binding region via the switch II-rigid relay loop to produce a large conformational change in the lever arm, providing the structural basis

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of the power stroke (100 Å or 30–40◦ for a muscle myosin [40] and up to 90◦ for some myosin I subclasses; [65]; Fig. 4.3). Myosin’s nucleotide-binding pocket is a Walker fold, a fold common to the ATPase domains of kinesins, G-proteins, helicases, and ATP synthase [66–68]. Thus, it is likely that these proteins use a common mechanism to communicate structural changes in the active site to other functional regions in the protein. Switch I directly interacts with the nucleotide’s phosphates within the binding pocket and moves from an “open” position to at least partial closure upon ATP binding (step 1 in Fig. 4.2), and in some way adjusting to allow a water molecule into the pocket for hydrolysis of the γ-phosphate bond [69]. Switch II is a loop located deep in the cleft between the upper and lower 50-kDa domains that coordinates the interactions with the γ-phosphate of ATP. Because of the location of switch II it is proposed to communicate hydrolysisassociated changes at the nucleotide-binding pocket to the actin-binding region (reviewed in [70]). The switch II loop is connected to a long alpha-helix, known as the switch II helix, that changes conformation resulting in movement of the rigid relay loop and inducing rotation of the converter domain during the lever arm recovery stroke ([71]; step 1 in Fig. 4.2). It is not known if the reverse process occurs during the power stroke (step 4 in Fig. 4.2). Evidence suggests that either or both switch I and II may shift upon actin binding to create a back door for phosphate release [41, 69, 70], while the nucleotide-binding pocket remains closed [72]. The P-loop also forms interactions with the γ-phosphate and may play a role in communication between the nucleotide pocket, transducer region, and a long alpha-helix that stretches into the actin-binding region to the hypertrophic cardiomyopathy (HCM) loop [49]. Loop 1 is a surface loop located at the entrance to the nucleotide-binding pocket and is thought to play a role in mediating the rate of ADP binding/release, although the specific mechanism is unresolved. For example, smooth muscle myosin has an alternatively spliced region in loop 1 that results in the addition of seven amino acids in this loop. The plus-insert isoform increases the rate of ADP release and leads to a twofold faster motility in phasic, compared to tonic smooth muscle myosin [73–76]. Many studies have converged on a working model for how myosin alternates between weak and strong actin-binding states during the contractile cycle. The area surrounding the actin-binding cleft has four actin-binding loops, one on the lower 50-kDa domain (loop 3), one spanning the outer cleft (loop 2), and two on the surface of the upper 50-kDa domain (loop 4 and the HCM loop). The highly charged loop 2 participates in an electrostatic association with actin in the weak binding states, such as the post-hydrolysis M.ADP.Pi state [77–79]. A shift to strong stereospecific binding is thought to be associated with closure of the actin-binding cleft, which may occur before or after inorganic phosphate release and the force-generating power stroke [71] (steps 3 and 4 in Fig. 4.2). Actin binding by both the HCM loop and a surface hydrophobic triplet on the lower 50-kDa domain seems to be required for cleft closure and phosphate release [78]. Dissociation of the post-rigor actomyosin complex upon ATP binding, on the other hand, requires actin-binding cleft opening (step 1 in Fig. 4.2). This opening is promoted by the attraction of switch I, switch II, and the P-loop to the bound nucleotide and subsequent rotation of the motor’s upper and lower 50-kDa domains ([80]; Fig. 4.3).

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The force generated by a single myosin molecule has been measured with optical tweezers and found to be in the range of 1.5–10 pN [81–83]. The velocity of movement of actin filaments generated by myosin cross-bridges can be measured with the in vitro motility assay. Skeletal muscle myosin can induce actin filament sliding in the in vitro motility assay at a velocity of 8,000 nm/s [84], which is similar to the contractile velocity measured with isolated muscle fibers (6,000 nm/s; [85, 86]). The in vitro motility rate and in vivo contractile velocity correlate well with the maximal ATPase rate measured in different muscle myosin isoforms, which was noted in early studies [87]. However, this correlation is very poor when nonmuscle and smooth muscle myosins are included. The correlation breaks down when the myosin motors being compared have a different duty ratio, the fraction of the ATPase cycle that myosin spends bound to actin [83]. For example, myosin V has a high duty ratio that which allows it to stay attached to actin for a larger fraction of its ATPase cycle and walk processively along actin filaments, which is important for its function as an organelle transporter. Conversely, skeletal muscle myosin has a low duty ratio which allows it to function well in an ensemble of motors integrated into the highly organized sarcomere structure. The concept of duty ratio is critical for understanding the molecular mechanism of force generation and allows comparison of the macroscopic and microscopic parameters [83].

4.3 The Myosin Tail Although the delineation of myosin classes is strictly based on phylogeny of the core motor domain, it is the C-terminal tail region of the molecule that demonstrates greatest diversity in sequence length, composition, and organization. Its subdomains and motifs form associations with specific cargoes or other binding partners for localization or transport, and also are responsible for the dimeric quaternary structure and filament characteristics of certain myosin classes. Moreover, tail domains of many myosins appear to play important regulatory roles by modulating the activity of their motor domain. A myosin’s functional capacity, then, depends on the fine-tuning of both motor and tail and also the interactions between them.

4.3.1

The Coiled-Coil-Forming Region

The so-called two-headed myosins—including all members of class II, V, VI and probably VII, and X—are dimeric as functional molecules instead of (for some, in addition to) functioning in the monomeric condition. Pairs of identical myosin monomers dimerize by association of their proximal tail which is composed of an α-helical coiled-coil-forming subdomain that serves as this linkage, although the unique MYO6 may be an exception to this rule [88]. For the class II myosins, which encompass all filament-forming subtypes, virtually the entire tail region forms a long rod-like junction between the two monomers. The sequence of this dimerization

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region comprises multiple heptad repeats with hydrophobic residues at positions 1 and 4 in each heptad which spontaneously interact in a coiled coil [89]. A thick filament for a class II myosin is assembled by association of these twisted rods with one another via their charged distal C-terminal residues into a subclass-specific organizational structure [90, 91]. Dimerization of certain unconventional myosins (non-filament-forming) allows their motors to coordinate each one’s biochemical/mechanical cycle with the other so that they alternate in a hand-over-hand walking motion along an actin filament or bundled actin to transport cargo. This has been directly observed using single-molecule detection of fluorescently tagged motors of MYO5A, MYO6, and leucine-zipperlinked MYO7A and MYO10 [92–95]. It is thought that coordination of the two motors for processive motion is enhanced by communicating the forces exerted between the actin-attached leading and trailing heads, a process designated as “gating” whereby strain induces modification of the timing of ADP release [96–100]. Because ADP release is the rate-limiting step for high duty ratio motors, this mechanism ensures that one myosin head will be attached to the actin filament at all times and prevents diffusion away from actin during processive walking [101, 102]. The impact of dimerization on the mechanochemistry of filamentous class II myosins is more speculative. However, it has been reported for skeletal muscle myosins that the cross-bridge pairing appears to enhance binding of the second head to the actin filament when the muscle is stretched. Apparently, the strain on one attached cross-bridge during the stretch effectively pulls the second head into close proximity to the actin filament allowing for cooperative force generation, a possible explanation for the rapid increase in force well-known as a physiological response to sudden muscular stretch [103].

4.3.2

Other Tail Subdomains

There are numerous other subdomains within the tails of members of the myosin superfamily that localize a particular myosin within a cellular region, attach to a cargo for its transport, or regulate the activity of the motor. Some of these are specific to one myosin class or subclass while others are more broadly distributed, and many of these subdomains are homologous to binding elements in other proteins. Class I myosins regulate membrane-cytoskeletal interactions of many types by their dual ability to bind actin and membranes. They localize to nearly all membranebound organelles and especially structures near the plasma membrane by use of specific tail sequences. Certain isoforms have tail SRC homology 3 (SH3) domains, a protein element known to bind other proteins via proline-rich regions, and also found as an N-terminal insert on muscle myosins. Class I myosins have a pleckstrin homology domain (PH), a membrane-binding element that associates with phosphoinositides [104]. Still other tail sequences found within this class include proline-rich segments and ATP-independent actin-binding motifs [105]. The filopodia-associated MYO10 tail has three PH domains that bind phosphoinosides of plasma membrane [22, 106]. MYO10 also uses a myosin tail homology (MyTH4)-FERM domain that

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most likely serves a cytoskeleton-plasma membrane linking function similar to its role in myosin VII and XV [107, 108]. SH3 domains also are present in the tails of myosins VII and XV where, as in myosin I and II, they may bind proline-rich protein sequences intramolecularly and/or those of target proteins [109]. Both these classes of myosin associate with the bundled actin at the core of stereocilia within the inner ear, and may bind actin-associated proteins with their SH3 domains [109]. In addition to the SH3 domain both also have two paired MyTH4-FERM domains. The FERM component binds integrins of the plasma membrane. While MyTH4 domains are common to several myosin classes their cellular role is currently being studied and this domain was found to mediate interactions with microtubules [27, 110, 111]. The various isoforms of myosins V and VI have globular cargo-binding domains (CBD) at their C-termini that utilize specific adapter proteins to link them with localization targets or vesicular cargo. The CBD sequences do not include the identifiable protein-binding motifs described for other myosin classes [109, 112]. MYO6’s short lever arm and surprisingly long step size have been attributed to its use of the CBD instead of its medial alpha-helical region as the portion of the tail responsible for dimerization [88]. The proximal tail region of MYO6 appears to uncoil into a single alpha-helix (SAH) upon dimerization by the CBD, extending the effective length of its lever arm [113]. These CBDs inactivate their motor by binding it in the absence of cargo to assume an inactive, folded conformation [88, 109, 112]. MYO3A, expressed in sensory receptors of the eye and inner ear, has two unique tail domains designated 3THD1 and 3THD2 and also has a tail IQ (calmodulinbinding) motif. THD1 binds espin-1, an actin-bundling protein that is transported in association with this myosin to the tips of stereocilia, while THD2 contains an ATP-independent actin-binding motif that has been speculated to assist in processive movement of monomeric MYO3A [114, 115].

4.4

Regulation

A myosin’s activity must be tightly regulated in order to optimize efficiency in its cellular role. Any explanation of a myosin’s cellular role is incomplete without a description of the regulatory mechanisms it employs to maintain homeostasis. There are three broad, somewhat overlapping, categories of regulatory mechanisms reported for various members of this superfamily. These include calcium concentration, phosphorylation state, and tail -motor interactions. For the conventional myosins, calcium plays a central role in the tight regulation of motor activity. The stimulus for contraction in cardiac and skeletal muscle results in sudden calcium influx in the vicinity of the contractile apparatus, binding of calcium ions to troponin C of the thin (actin-based) filament and subsequent induction of a structural change that allows actomyosin cycling to begin [116, 117]. For smooth muscle myosin and the nonmuscle class II myosins, calcium influx

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results in activation of myosin light chain kinase (MLCK), phosphorylation of the myosin regulatory light chain by activated MLCK, resulting in motor conformational change to an active state [118]. Cessation of the stimulus prevents further activity by various means involving dephosphorylation of the light chain, calcium removal, and motor and/or tail phosphorylation [118]. Regulation for the unconventional myosins employs calcium concentration dependence, phosphorylation state, and cargo binding to tail domains. Calcium concentration influences motor activity of myosin V by its effects on calmodulin light chain binding, on cargo binding to the globular tail domain (GTD), and on tail-motor interactions [119–121]. The calcium-dependent structural changes are characterized by an extended conformation (active) and a folded (inactive) conformation in which the myosin V GTD interacts with the motor domain [121]. Independent of calcium concentration and in the absence of cargo, tail-motor interactions can downregulate myosin V and VII activity [109, 122–124]. For many myosin I isoforms, motor phosphorylation of a serine or threonine residue at the highly conserved TEDS site (so called because the position must be occupied by a phosphorylatable threonine or serine or must be glutamic or aspartic acid) on the motor’s cardiomyopathy loop serves to upregulate motor activity [125–127]. In MYO6, the role of TEDS phosphorylation is less clear, but appears to influence the motor’s kinetic properties in a calcium concentration-dependent manner [128–130]. The myosin III kinase domain autophosphorylates Limulus myosin III motors on or near loop 2, downregulating their activity by a reduction in actin affinity due to charge reduction [131, 132], while the exact phosphorylation sites in mammalian MYO3A are unknown. Several myosin classes are functional as both monomers and dimers, with different cellular roles for each condition. MYO6 appears to function mainly as an anchor in its monomeric state and walks processively once cargo binding instigates dimerization [88, 124, 133–135]. Myosin VII and X also are likely to alternate between monomeric stabilizers and dimeric transporters depending upon the availability of cargo [102, 124]. A “loose” coiled coil in myosins VI, VII, and X’s proximal or medial tail regions can unfold to become SAH under strain. This feature appears to be a mechanical component of the molecule that allows for extension of the effective lever arm. The ability to unfold into a SAH [136] appears to modify myosin X and VII for efficient walking on bundled actin so that they remain on the surface rather than following a single filament into the interior of the bundle [137]. This ability to partially disassemble the coiled coil also appears to be important for regulation in smooth muscle and some striated myosins by allowing them to assume an inactivated folded conformation [138].

4.5

Contractile Roles of Myosin

The first myosin to be well characterized and carefully studied was muscle myosin, a class II myosin responsible for muscle contraction. A total of 15 class II myosins exist in the human genome [1, 4] encoding 8 found in skeletal muscle, 2 found

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in cardiac muscle, one smooth muscle myosin, three nonmuscle myosins, and one pseudogene related to myosin found in the masticatory muscle of carnivores [6]. The physiological and contractile properties of specific muscle fibers vary with the specific isoform of myosin found in each fiber type [139, 140]. In addition, the associated proteins found in the sarcomere, such as the myosin-associated light chains, also vary with fiber type contributing to the physiological function of each muscle type [141]. For example, MYH7B, MYH15, and MYH13 are expressed exclusively in extraocular muscles [142, 143]. Although fibers usually express one myosin isoform predominantly, muscle tissue is often a combination of multiple fiber types and this distribution determines the properties of a particular muscle [141]. When myosin isotype expression is heterogeneous within a fiber, it is often in response to external cues [144] such as during development, or in response to injury [145]. Two cardiac myosin genes, MYH6 and MYH7, are preferentially expressed in either atrial or ventricular myocardium; relative expression of each gene varies under differing developmental and disease states [146], suggesting different contractile roles in each region of the heart [147, 148]. Mutations in MYH7 (cardiac myosin heavy chain beta) are the most common cause for familial hypertrophic cardiomyopathy (FHC) [149]. Although there is only one smooth muscle myosin gene, alternative splicing leads to the expression of two separate splice-forms [150] with amino acid differences in both the motor [73, 151] and the tail [152–154]. In addition, differential expression of light chains associated with smooth muscle myosin [155] provides specificity to the contractile properties of the genitourinary, gastrointestinal, and vascular systems. The three class II nonmuscle myosin genes are involved in contraction of the cytoskeleton during cell migration. At the leading edge of the cell, nonmuscle myosins play roles in the maturation of focal adhesions and stress fiber formation [156, 157] and lamellipodial extension [158]. Nonmuscle myosin II also incorporates into actomyosin structures during cell division [159], driving the contraction of the cytokinetic furrow during cytokinesis [160, 161].

4.6

Myosins in Cytoskeletal Dynamics and Assembly

In addition to their roles in adhesion and lamellipodial protrusion, nonmuscle myosin II genes are also involved in the formation and regulation of cell-cell adhesion complexes including tight junctions and their associated apical actin belt [162–164], in supporting cell spreading on fibronectin [165], and in cell polarization persistence of directed cell migration [166]. In addition, fate specification of stem cells via sensing of substrate stiffness involves nonmuscle myosin II [167]. Multiple myosins have been shown to play roles in regulation of the cellward flow of polymerized actin, known as retrograde flow, including MYH9, MYH10, and MYO1C [168–170]. Class III, X, and XV myosins localize to and may be involved in regulation of the formation of actin-based protrusions including filopodia and stereocilia. MYO3A localizes to the tips of hair cell stereocilia and may function in conjunction with espin-1 to mediate stereocilia formation/stability, as expression of certain MYO3A

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constructs in COS7 cells stimulates filopodial elongation [114] and filopodial density [131]. MYO15A also localizes to the tips of actin projections, and mice deficient in MYO15A have abnormally short stereocilia and are deaf [171]. MYO15A is thought to function in concert with whirlin to regulate stereocilia length, and in the COS7 expression system, increased levels of MYO15A and whirlin at the filopodia tip correlated with increased filopodia length [172]. Mutations in either MYO3A or MYO15A can cause nonsyndromic forms of deafness (DFNB3 and DFNB30, respectively), while Usher IB, the most common cause of deaf-blindness in humans, can arise from mutations in MYO7A [173, 174]. MYO10 has also been shown to localize to filopodial tips and may shuttle cargo to the filopodial tips as single molecules [175]. MYO10 function regulates both filopodia number and length when expressed in cultured cells [176], possibly through interactions with VASP [177, 178], although some of the function of MYO10 may be VASP-independent [179]. Myosins may also serve as negative regulators of actin dynamics. The GTP-bound forms of Rho proteins are regulators of actin-based processes such as contractility, adhesion, and lamellipodial protrusion [180]. Localization of Rho-GAP domains to regions of dynamic actin via myosin motor activity may allow regulation of actin dynamics; and class IX myosins contain a Rho-GAP domain in their tail [181, 182], capable of stimulating the GTP hydrolyzing activity of Rho GTPases [183] and thereby regulating Rho-mediated cytoskeletal rearrangements. In addition to interactions and regulation of the actin cytoskeleton, some myosins may regulate the intermediate filament and microtubule cytoskeletons as well. MYO5A has been shown to be capable of transporting intermediate filament subunitcontaining complexes in neurons [184]. It is also possible that MYO5A may play a role in cross-talk between the actin and microtubule cytoskeletons during cell migration [185] as it has also been shown to bind microtubules directly [28] and indirectly [186]. MYO10 also interacts with microtubules [27] and functions in podosome positioning in osteoclasts [187], meiotic spindle assembly [27], mitotic spindle pole integrity [188], and mitotic spindle pole orientation [188, 189]. Class II MYH9 [190] [191] localizes to spindle poles during mitosis, and chemical disruption of myosin light chain phosphorylation or myosin II function lead to contractile ringspecific and spindle rotation-specific defects in egg activation following fertilization of mouse oocytes [192].

4.7

Myosins in Membrane Dynamics

Interactions between phospholipid membranes and myosins is a well established myosin function, as multiple classes of myosins are known to bind to membranes or membrane-bound organelles [193]. Endocytic processes such as macropinocytosis [194], phagocytosis [195, 196], and some aspects of clathrin-mediated endocytosis [197] have all been shown to involve multiple myosin classes. Dendritic cells undergo macropinocytosis as part of their immune surveillance role, and cells deficient in MYO6 display increased macropinocytic uptake, suggesting MYO6 somehow serves as a restrictor of such processes [198]. Class II nonmuscle

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myosins, which can be directly inhibited with pharmacological agents, have also been shown to play a role in macropinocytosis in both macrophages [199] and nerve cells [200]. In a mechanism similar to the possible antagonistic roles of nonmuscle class II myosins in retrograde flow, selective inhibition of MYH9 and MYH10 resulted in either increased macropinocytosis or decreased macropinocytosis, respectively [201]. Myosins function in a variety of stages of the phagocytic process. MYO10 is recruited to the phagocytic cup playing a role in pseudopod extension during Fcreceptor-mediated phagocytosis [202], and cup closure may involve MYO1E [203]. Similarly, a number of clathrin-mediated endocytic processes involving receptors require myosin function, such as MYH9 function in chemokine receptor internalization [204] and MYO6 in neurons and in kidney proximal tubules [205, 206]. As MYO6 is a minus-end-directed motor, its motor activity may function to exert an inward tension on the newly forming vesicle [207] and mediate transport to early endosomes [208]. Myosins also function in organelle traffic and at multiple points along the exocytotic pathway. MYO6 associates with the Golgi [209]; and loss of MYO6 function results in disruption of Golgi morphology and reduced protein secretion [210]. Melanosome positioning in the cell periphery has been shown to be MYO5A-dependent [211, 212], and MYO5A also carries smooth endoplasmic reticulum [213, 214], secretory vesicles [215], and other membranous organelles as cargo [216, 217]. MYO5B plays a role in recycling of plasma membrane proteins such as EGF-receptor [218], transferrin receptor [219], aquaporin 2 [220], and other receptors [221, 222]. MYO5C participates in the traffic of secretory vesicles in exocrine tissues such as the lacrimal gland [223, 224]. When examined in neurons, class V myosins are important for long-term potentiation [225, 226] and long-term synaptic depression [227] through trafficking and positioning of particular membranous cargoes. Most recently, MYO19 was shown to be involved in mitochondrial transport/positioning. Immunostaining indicates that endogenous MYO19 localized to mitochondria and expression of GFP-MYO19 led to increased mitochondrial dynamics in A549 non-smallcell lung cancer (NSCLC) cells. Expression of a GFP-MYO19 tail construct (lacking the motor domain) resulted in decreased mitochondrial run lengths in cultured neuronal cells [228]. Myosins have also been implicated in roles where they serve as a dynamic linkage between the plasma membrane and the cytoskeleton in protrusive structures, such as microvilli, filopodia, and stereocilia [229]. In regions where multiple actin protrusions exist, links between the actin cytoskeleton and the plasma membrane are required to maintain the membrane coverage of individual protrusions and prevent the fusion of multiple projections resulting in a loss of membrane surface area. MYO1A [230, 231], MYO7A [173], and MYO6 [174] may serve to regulate membrane tension in bundled-actin structures, as defects in those genes result in aberrant membrane association with bundled-actin structures. In addition to maintaining membrane tension, recent studies demonstrate that MYO1A powers the generation of alkaline phosphatase-containing vesicles at the tips of microvilli in the intestinal brush border, resulting in the release of these enzymes into the gut [232, 233].

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Myosins and Nucleic Acids

In addition to their traditional cytosolic roles, recent studies have shown that myosins may also be involved in nucleic acid transport and implicated in a variety of functions in conjunction with nuclear actin. Although the majority of experimental evidence for class V myosin function in messenger RNA transport comes from work in nonmammalian model systems such as yeast [234, 235] and Drosophila [236], evidence does exist for localization of mRNA in mammals via certain splice-forms of MYO5A. MYO5A is present in polyribosome mRNA/protein complexes known as periaxoplasmic ribosomal plaques (PARPS [237, 238]), and has also been localized to other ribonuclear complexes as well [239]. Specific mRNA localization in dendritic spines was shown to be dependent on functional MYO5A, as both RNAi and dominant negative experimental approaches led to impaired localization of messenger ribonucleoprotein complexes in dendritic spines [240]. In primary fibroblasts isolated from MYO5A null mice, mRNA localization was disrupted, and could be rescued by exogenous expression of MYO5A [241]. Many myosin classes have been localized to the nucleus and implicated in a variety of functions along with nuclear actin. Particular isoforms of MYO1 [242, 243], MYO2 [244], MYO5 [245], MYO6 [246], MYO16 [11], and MYO18 [247] have been localized to the nucleus, although their specific functions have not all been identified. MYO6 localizes to the nucleus upon DNA damage and is thought to be a mediator of the p53 prosurvival pathway [246]. MYO16 function has been linked with progression through S-phase of the cell cycle [248]. MYO1 functions in initiation of rRNA transcription via RNA polymerase I [249–252], while MYO5B also appears to interact with RNA polymerase I [253]. Both MYO1 [243, 254] and MYO6 [255] are involved in initiation of RNA polymerase II-mediated transcription. Class I myosins are thought to play roles in the processing [256] and transport of preribosomal complexes to the nuclear pore for export [257, 258]. Nuclear MYO1 functions in the arrangement of chromatin within the interphase nucleus [259–261].

4.9

Other Cellular Roles for Myosin

Research continues to elucidate functions for myosin genes that might not have been anticipated by the founders of the field. For example, MYH9 appears to play a central role in the proper formation of platelets as a variety of mutations in MYH9 result in platelet-related diseases [262]. Multiple reports indicate that one of the transmembrane receptors for the complement protein, surfactant protein-A (SP-A) is encoded by the MYO18A gene [263, 264] and coordinates a number of cellular responses in macrophages [265, 266], T lymphocytes [267], and alveolar type II cells [263]. Just as other cytoskeletal structures are associated with infection and pathogenesis [268, 269], MYO10 is involved in the budding of the Marburg virus [270] and MYO5A in herpes virion secretion [271]. Multiple myosin genes have also been linked to cancer. MYO18B has been implicated as a tumor suppressor with decreased expression levels in particular lung, ovarian, and colorectal cancers [272–275]. With

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increased expression levels in some prostate, gastroesophageal, and lung cancers, MYO6 function may be central to cancer cell development [276]. In fact, the clinical behavior of ovarian cancers has been correlated to levels of MYO6 expression [277]. MYH11 is a conventional myosin that also has links to cancer, as chromosomal abnormalities resulting in the core-binding factor β protein-MYH11 fusion proteins (CBFB-MYH11) are associated with certain leukemias [278]. Myosin, initially characterized as the contractile force-generating component of muscle, has now been clearly revealed to be an ancient and diverse family [2, 3] involved in performing a wide variety of physiological functions far removed from muscle contraction [193, 279, 280].

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Part II

Actin and Disease

Chapter 5

The Actin Cytoskeleton and Membrane Organisation in T Lymphocytes Rhea Cornely, Thomas Grewal and Katharina Gaus

Abstract The dynamics of the actin cytoskeleton are under the strong influence of signalling events at the membrane and membrane domains. In T cells, the importance of actin-membrane-interactions is critical in the processes of T cell activation and migration. If the coordination of actin and the T lymphocyte membrane are out of order, it can compromise the human immune system and lead to immunodeficiencies.

5.1

Introduction

Lymphocytes play an important role in the immune response and are made up of B and T lymphocytes. B cells bind antigens, for example, fragments of bacterial protein, to their major histocompability complex (MHC) class II. CD4+ T lymphocytes or T helper cells bind the antigen and the major histocompability complex with their T cell receptor (TCR) and CD4 co-receptor, respectively, and thus become activated. While activation of T lymphocytes via the TCR ultimately leads to T cell proliferation, the activation of a chemokine receptor on a T cell by a chemoattractant will trigger migration towards the source of chemoattractant and possibly to an encounter of an antigen-presenting B cell. Both receptor systems share elements of common signalling cascades including the key signalling molecules for the restructuring of the actin cytoskeleton. Here, we focus on the signalling pathways and the role of membrane domains in the regulation of the actin cytoskeleton in T cell activation and migration. Actin plays a central role in T cell activation. The interaction of a TCR with the major histocompability complex of an antigen-presenting cell triggers polymerisation and restructuring of the filamentous actin (F-actin) cytoskeleton. Genetic mutations that affect proteins involved in TCR-induced actin restructuring can potentially lead to immune deficiencies due to poorly activating T lymphocytes. At least three known types of immunodeficiency syndromes, Wiskott-Aldrich Syndrome (WAS), K. Gaus () · R. Cornely Centre for Vascular Research, University of New South Wales, 2052 Sydney, Kensington (NSW), Australia e-mail: [email protected] T. Grewal Faculty of Pharmacy, University of Sydney, Sydney, NSW, Australia

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X-linked neutropenia (XLN) and some cases of CommonVariable Immunodeficiency with defective T cell function (T-CVID) are linked to mutations in the sequences for proteins involved in organising F-actin architecture, T cell activation and signalling. Our understanding of the importance of actin regulation in cells of the immune system has grown rapidly in recent times. In most cells, the actin cytoskeleton serves the purpose of stabilising cell shape and in assisting in anchoring cells to neighbouring cells and the extracellular matrix. The regulation of actin dynamics in cells of the immune system, however, also enables the ligation and signalling of immune and chemokine receptors. Rather than forming a stable actin network, in this context, actin is rapidly remodelled during T cell activation and migration. In T cells, for example, F-actin forms short stick-like structures rather than long fibrous structures. Actin organisation varies at different receptor activation sites and, in fact, defines the immunological synapse, which is the contact side with the antigenpresenting cell and the opposite side of the cell, known as the distal pole. In a migrating cell, the actin-rich leading edge forms the front end of the cell where activated chemokine receptors are located and rigorous assembly and disassembly of actin fibres facilitates directed movement. While many of the proteins involved in actin regulation in T lymphocytes have been identified, the role of membrane domains and hence the targeting of actin to receptor activation sites is less well understood.

5.2 Activation of T Cells Signals for T lymphocyte activation are transmitted at the so-called immunological synapse, which is the contact zone between T cell and an antigen-presenting cell [1]. TCRs recognise peptides bound to major histocompatibility complexes on the surface of an antigen-presenting cell (Fig. 5.1). The recognition is confirmed by the co-receptor CD4, which binds to MHC. TCR ligation triggers the segregation of the T cell membrane and the assembly of multi-molecular TCR signalling clusters. This process is initiated when engaged TCRs form a TCR/CD3 complex, which recruits the Src family kinase, Lck (lymphocyte specific protein tyrosine kinase) and Fyn, and the protein tyrosine kinase, ZAP70 (ζ -chain associated protein kinase of 70kD). Activated by Lck, ZAP70 subsequently phosphorylates the transmembrane protein LAT (linker for activation of T cells), which acts as a scaffold for adaptor proteins Grb2 and SLP76, and signalling enzymes including PLCγ (phospholipase Cγ) and PI3K (phosphoinositol-3-kinase). This leads to TCR-LAT oligomerisation and results in the recruitment of proteins involved in actin regulation like NCK (noncatalytic region of tyrosine kinase), ITK (interleukin-2-inducible T cell kinase), Vav1 and subsequently DNM2 (dynamin 2) and EZH2 (enhancer of zeste homologue 2). The link between TCR-LAT clusters and actin restructuring is necessary to couple TCR engagement to the induction of calcium fluxes (via PLCγ), activation of PI3K, the Ras/MAPK (mitogen-activated protein kinase) signalling pathway, cytoskeleton restructuring [2, 3] and ultimately, cytokine production and proliferation.

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Fig. 5.1 From T cell activation to actin polymerisation. T cells become activated when the TCR detects and binds to an antigenic peptide presented on a major histocompability complex (MHC) of an antigen-presenting cell. Peptide recognition is confirmed by co-receptor such as CD4, which also binds to the MHC. The ligation of the TCR initiates a signalling cascade. The kinase Lck is activated and leads to recruitment and activation of ZAP70 via the TCR ζ chain (Phosphorylation is indicated by small dark circles.). ZAP70 in turn initiates a chain of phosphorylations. ZAP70 phosphorylates LAT, which is associated to GADS and recruits SLP76, which assumes the role of a scaffold to which further adapter proteins and kinases bind. This facilitates the activation of VAV1. VAV1 ultimately causes the activation of WASp and WAVE, two proteins of the same family, which trigger the nucleation core activity for actin filaments of the Arp2/3 complex upon binding

During the initial interaction stages (70% of cases; [115]). The relative lack of cases with Tm and actin mutations may mean that mutations in these genes have a greater effect on protein function and result in early lethalities, or the pathophysiology of patients with Tm and actin mutations is more variable (e.g., hypertrophy is not a consistent feature of Tm-based HCM; [114]) and therefore may be excluded in linkage analyses. One of the defining features of thin filament cardiomyopathies which contrasts with β-myosin heavy chain-based forms is the dissociation between the clinical severity and the degree of ventricular hypertrophy. This indicates that there are disease-specific processes resulting from mutations in thin versus thick filament proteins.

6.3 6.3.1

Mouse Models of Thin Filament Diseases The TPM3(Met9Arg) Mouse Model of NM

The only published mouse model of NM is a transgenic mouse expressing the dominant negative TPM3(Met9Arg) in skeletal muscle [16]. This mouse model has all features of the human disease including the presence of nemaline rods in skeletal muscle, an increase in slow fibers and fast fiber hypertrophy. The fast fiber hypertrophy declines with increasing age coincident with a loss of muscle strength beyond 5–6 months of age. This mimics the late-onset of the disease in patients with this mutation [53]. In the TPM3(Met9Arg) mouse, the increase in slow/oxidative fibers was apparent at 1 month of age and was maintained throughout adulthood, indicating that the normal maturation of the muscle fiber types was altered in this mouse. The number of rod-containing fibers varied widely in the different muscles of the mouse and did not correlate with the level of mutant protein in the muscle. As has been observed in cardiomyocytes [18, 62, 63], the mutant protein incorporated into the thin filament in muscles from the NM mouse [16, 17]. Despite this, in some muscles

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(e.g., soleus and gastrocnemius) there was virtually no disruption to the sarcomeric structure indicating that the presence of the Met9Arg mutation is not sufficient to alter the normal formation of the thin filament.

6.3.2

Cardiomyopathy Models

There are now a large number of animal models of all major forms of familial cardiomyopathy (HCM, DCM, and RCM) and the reader is referred to recent reviews for detailed discussion of these models [9, 100, 115]. Both transgenic and knock-in approaches have been used to model CMs in the mouse, and transgenic rat [33, 34, 58] and rabbit [59] models have also been created. β-myosin heavy chain (βMyHC; [36, 74]), αTm [66, 80, 81, 82], cTnT [71, 101, 102], cTnI [112, 113], cardiac actin [94] CM mutations have all been modeled in the mouse. The animal models phenocopy many of the aspects of the human disorders including both pathological and contractile features. They have provided important insights (summarized in the next section) into the mechanisms for contractile dysfunction and cardiac failure due to sarcomeric protein mutations and are providing excellent model systems for trial of potential therapeutic strategies (summarized in the next section).

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Disease Processes and Causes of Muscle Dysfunction in Thin Filament Diseases Nemaline Myopathy

The precise involvement of nemaline rods in the development of muscle dysfunction is unclear. In patients, the number of rod-containing fibers correlates poorly with the severity of disease. Similar findings have been obtained in the transgenic mouse model where in young animals ( T c.353A > T c.354G > C c.364A > G c.721G > A c.792C > T c.833C > T c.994C > G c.1109T > C

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The actin regulator Twinfilin 2, an actin-capping protein, which localizes at the tips of short- and middle-row cochlear stereocilia, but not in tall-row stereocilia, and regulates the length of the stereocilia F-actin core in developing and mature cochlear hair cells [33]. Twinfilin 2 displays a strong ability to cap actin filament barbed ends as well as an ability to sequester actin monomers [31, 43]. Finally, the scaffolding PDZ protein whirlin is a key molecular component of stereocilia membrane and F-actin growth in the inner ear that is critical for actin cytoskeletal assembly [24, 25].

9.2

Genetic Epidemiology of ACTG1

Whereas mutations in both, γ- and β-actin, were initially suspected to lead to syndromic disorders as they are nonmuscle cytoplasmic actins ubiquitously expressed; this assumption was only true for β-actin. A mutation of β-actin that alters depolymerization dynamics was associated with autosomal dominant developmental malformations, deafness, and dystonia [35]. However, mutations in ACTG1 have been reported to be associated with the autosomal dominant nonsyndromic sensorineural hearing loss DFNA20/26, a locus that has been mapped to chromosome 17q25 in several independent families [6, 29, 46]. At present, only ten mutations in ACTG1 have been identified that cause DFNA20/A26, including four in American families (T89I, K118M, P264L, P332A), two in Dutch families (D51N, T278I), one in a Norwegian family (V370A), one in a Chinese family (I122V), and two in Spanish families (K118N and E241K; [12, 20, 30, 36, 45, 47]; Table 9.1). The global prevalence of DFNA20/A26 is unknown as comprehensive population-based genetic screening has not been conducted but in the Spanish population, for which mutations in ACTG1 appear to be accounting for less than 1% in ADNSHL [30]. All patients with ACTG1 mutations characteristically show postlingual, bilateral, symmetrical, and progressive sensorineural hearing loss primarily affecting high frequencies that progresses later to affect the entire frequency range quickly becoming severe to profound [7, 16, 36]. Only subtle phenotypic differences in patients carrying the different mutations in ACTG1 have been observed and no clear correlations can be established between both the type of mutation and the subdomain affected with the

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Fig. 9.2 Locations of the γ-actin mutations associated with DFNA20/A26 hearing loss. Front view of the crystal structure of the yeast actin monomer [44], modified from Protein Data Bank code 1YAG using PyMol. The positions of the mutations are modeled in space-fill as following: D51N, T89I, K118M, K118N, I122V, E241K, P264L, T278I, P332A, and V370A. Numbers denote the actin subdomains. The amino and carboxilo terminal ends of the protein are represented as N and C, respectively

age of manifestation or severity (Fig. 9.2). Mutations in subdomain 3, P264L, T278I, and P332A, are all associated with hearing loss that appears in the second decade, however, in the case of T89I, K118M, K118N, I122V, and V370A, all located in subdomain 1, the reported age of onset differs widely, ranging from the first to the third decade (Table 9.1). Note that even mutations affecting the same residue, K118M and K118N, are distinctly associated with ages of onset in the second and third decades, respectively. The E241K mutation, in subdomain 4, causes hearing loss with onset in the first decade. Similarly, carriers of the novel D51N mutation, the only reported to affect subdomain 2, display hearing loss that appears in first decade of life, and quickly progresses until moderate to severe in third decade. Apparently, this is the first family with DFNA20/A26 hearing loss that shows vestibular symptoms, including dizziness, occasional tinnitus, or vertigo [12].

9.3

Pathophysiology of DFNA20/A26 Hearing Impairment

Despite the lack of significant phenotypic differences observed between familiar cases with DFNA20/A26 hearing loss, the mutations hitherto described are localized in all subdomains of the protein (Fig. 9.3); hence they might be suspected to

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Fig. 9.3 Audiograms showing the air conduction values obtained from affected patients carrying the different ACTG1 mutations described. Each point represents the average hearing threshold for the right and left ears at the time of recording. Audiograms plot the hearing threshold levels in normalized decibels (dB) (on the vertical axis) against sound’s frequencies in hertz (Hz) (on the horizontal axis) covering a limited range (250−8,000 Hz, in a logarithmic scale) which is the most important for clear understanding of speech

be distinctly impacting the way in which the protein function is altered. Some recent works have focused on deciphering the mechanism of pathogenesis associated with the ACTG1 mutations. Bryan and colleagues studied in yeast the effects of six (T89I, K118M, P264L, T278I, P332A, and V370A) of the human deafness γ-actin mutations on actin function in vivo and in vitro [4]. Four of the six mutant strains

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(K118M, T278I, P332A, and V370A) exhibited growth deficiencies and abnormal mitochondrial morphology. All of the mutant yeast cells exhibited cytoskeleton abnormalities, malformed actin cables, or misdistribution of actin patches. The majority also displayed strain-specific vacuole morphological abnormalities. However, only abnormal polymerization was observed for the V370A mutant actin. On this basis, they suggested that DFNA20/A26 hearing loss could be the result of an altered ability of the actin filaments to be properly regulated by actin-binding proteins rather than an inability to polymerize. This hypothesis was further investigated by studying the effect of the γ-actin mutants on actin/cofilin interaction [3]. In vitro, T89I and V370A mutant F-actins were much more susceptible to cofilin disassembly than WT filaments, conversely, P332A filaments demonstrate enhanced resistance. Interestingly, depression of cofilin action in vivo rescued the inability to grow in glycerol caused by K118M, T278I, P332A, and V370A indicating that by decreasing a system that promotes increased filament turnover the filament instability caused by these mutations could be balanced. In a related study, Morin and colleagues investigate K118N and E241K mutations in a spectrum of different situations with increasing biological complexity by combining biochemical and cell biological analysis in yeast and mammalian cells [30]. They observed that K118N had in vivo a very mild effect on yeast behavior, however, the E241K phenotype was very severe consisting in an aberrant multivacuolar pattern, a highly compromised ability to grow on glycerol

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as a carbon source and the deposition of thick F-actin bundles randomly in the cell. The latter is consistent with the observed highly unusual spontaneous tendency of the E241K mutant to form bundles in vitro. When transfected in mammalian cells, both mutant actins were, however, normally incorporated into cytoskeletal structures with only the presence of some aberrant cytoplasmic aggregates. Interestingly, gene-gun-mediated expression of K118N, K118M, E241K, and V370A in cochlear hair cells revealed no gross alteration of cytoskeletal structures or in the morphology of stereocilia. Collectively, these findings supported the hypothesis that the postlingual and progressive nature of the DFNA20/A26 hearing loss could be the result of a mutant-dependent filament destabilization leading to progressive deterioration of the hair cell cytoskeleton. There are a number of possible explanations for the differences in the behavior of these mutant actins in yeast versus mammalian cells. The most likely fact is that in yeast, the mutant actin is the only actin in the cell, whereas in the mammalian cells, the full effects of the mutations are diluted by the presence of the γ- and βactins normally in the cell. The absence of abnormal effects on hair cell stereocilia revealed by gene-gun experiments could also reflect the fact that the stereociliary actin bundle is a much more stable cytoskeletal structure as it takes part of a complex protein network. Thus, the interaction of different proteins with the actin core, either directly or mediated by scaffolding proteins [19] may mask the underlying structural deficiencies associated with ACTG1 mutations. For example, T89I, in subdomain 1, is in the alpha helix that is thought to participate in binding of fimbrin. Similarly, K118M and K118N, also in subdomain 1, are near the fimbrin-binding domain that has been implicated in interactions with fimbrin, gelsolin [9], and γ-actinin [8, 27]. These proteins are implicated in bundle or gel-like structure formation, interaction with membrane or scaffold proteins, important in actin function. Gelsolin is found in OHC, but not in IHC, where it takes part in the whirlin complex that plays a role in stereocilia elongation [26]. The V370A mutation is in the C-terminal tail of the protein that forms part of a binding site for important actin-binding proteins, including cofilin, profilin, or gelsolin, proteins implicated in length control of F-actin bundles or in gel-like structure formation. E241K is very close to an actin-binding site (243–245) involved in intermolecular contacts along the F-actin helix and could produce changes in monomer interactions that alter filament topology leading to an association of actin filaments, although, as it has been demonstrated in vitro, the propensity to bundle of the E241K mutants can be neutralized by tropomyosin and the E241K filament bundles were hypersensitive to severing in the presence of cofilin [30]. The P264L and T278I mutations, in subdomain 3, are located in the self-assembly site of actin, and may affect polymerization or stability of F-actin. The mutation P332A, in subdomain 3, is located in a three-amino acid loop that is part of the primary contact site for myosin [8]. As it is well known, myosins are actin–based molecular motors that regulate several processes, such as rearrangement of the actin cytoskeleton, regulation of tension of actin filaments, and transport of organelles [28]. Four myosin members, Myosin VI, VIIA, XVA, and IIIA have been reported to play crucial roles in the inner ear hair cells and have also been implicated in hearing

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loss [42]. Myosin VI is expressed in the cuticular plate and is involved in stereocilia formation. At the minus-end, Myosin VI and the protein tyrosine phosphatase receptor Q (PTPRQ) regulate taper formation/maintenance [11]. Myosin VIIA is found in the stereocilia, lateral membrane of the cell, cuticular plate, and synaptic region, interacting with actin filaments. Myosin VIIa forms a complex with twinfilin-2 and this complex inhibits polymerization of actin in stereocilia cores [38]. Myosin XVA localized to the tip of the stereocilia near the barbed end of the actin filament where it may be involved, together with its cargoes whirlin and Eps8, in stereocilia elongation and formation [1, 23]. Finally, Myosin IIIA is localized around the stereocilia tips forming a thimble-like pattern. It is a barbed end-directed motor and its targeting to the tips of stereocilia depends on a motor-driven translocation along the actin filaments [40]. Recent investigations in mice have helped to bring insight on the function that γ- and β-actin play in the inner ear. Two studies have shown that null mice for cytoplasmic γ-actin, ACTG1(−/−), are viable although they suffer increased mortality and show progressive hearing loss during adulthood despite compensatory upregulation of beta-actin [2, 5]. The unexpected viability and normal hearing of young ACTG1(−/−) mice, because the ubiquitous expression of γ-actin, indicates that β-actin can likely build all essential nonmuscle actin-based cytoskeletal structures including mechanosensory stereocilia of hair cells that are necessary for hearing. Although γ-actin-deficient stereocilia form normally, they cannot maintain the integrity of the stereocilia actin core. In the wild-type, γ-actin localizes along the length of stereocilia but redistributes to sites of F-actin core disruptions resulting from animal exposure to damaging noise. In ACTG1(−/−) stereocilia, similar disruptions are observed even without noise exposure [2]. This study interestingly reveals that cytoplasmic γ-actin is required for reinforcement and long-term stability of F-actin-based structures but is not an essential building block of the developing cytoskeleton. In an additional study, Perrin and colleagues compared the functions of β-and γ-actin in stereocilia formation and maintenance by generating mice conditionally knocked out for ACTB or ACTG1 in hair cells [34]. They found that, although cytoplasmic actin is necessary, neither β-actin nor γ-actin is individually essential for normal stereocilia development or auditory function in young animals. However, aging mice with β- or γ-actin-deficient hair cells develop different patterns of progressive hearing loss and distinct pathogenic changes in stereocilia morphology, despite colocalization of the actin isoforms. The loss of β-actin preferentially affects the high frequencies while the loss of γ-actin affects the whole frequency range and progresses more rapidly. These results demonstrate overlapping developmental roles but unique postdevelopmental functions for β- and γ-actin in maintaining hair cell stereocilia. Next challenge is generating animal models, transgenic knock-in mice, for the DFNA20/A26-specific mutations that will shed light on the underlying pathogenic mechanism and will help in developing therapies to palliate the progressive detererioration of hearing that is characteristic of this subtype of hereditary hearing loss.

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20. Liu P, Li H, Ren X, Mao H, Zhu Q, Zhu Z, Yang R, Yuan W, Liu J, Wang Q, Liu M (2008) Novel ACTG1 mutation causing autosomal dominant non-syndromic hearing impairment in a Chinese family. J Genet Genomics 35:553–558 21. Mallavarapu A, Mitchison T (1999) Regulated actin cytoskeleton assambly at filopodium tips controls their extension and retraction. J Cell Biol 146:1097–1106 22. Manor U, Kachar B (2008) Dynamic length regulation of sensory stereocilia. Semin Cell Dev Biol 19:502–510 23. Manor U, Disanza A, Grati M, Andrade L, Lin H, Di Fiore PP, Scita G, Kachar B (2011) Regulation of stereocilia length by myosin XVa and whirlin depends on the actin-regulatory protein Eps8. Curr Biol 21(2):167–172 24. Mburu P, Mustapha M, Varela A, Weil D, El-Amraoui A, Holme RH, Rump A, Hardisty RE, Blanchard S, Coimbra RS, Perfettini I, Parkinson N, Mallon AM, Glenister P, Rogers MJ, Paige AJ, Moir L, Clay J, Rosenthal A, Liu XZ, Blanco G, Steel KP, Petit C, Brown SD (2003) Defects in whirlin, a PDZ domain molecule involved in stereocilia elongation, cause deafness in the whirler mouse and families with DFNB31. Nat Genet 34:421–428 25. Mburu P, Kikkawa Y, Townsend S, Romero R, Yonekawa H, Brown SD (2006) Whirlin complexes with p55 at the stereocilia tip during hair cell development. Proc Natl Acad Sci USA 103:10973–10978 26. Mburu P, Romero MR, Hilton H, Parker A, Townsend S, KikkawaY, Brown SD (2010) Gelsolin plays a role in the actin polymerization complex of hair cell stereocilia. PLoS One 5:e11627 27. McGough A, Way M, DeRosier D (1994) Determination of the a-actinin-binding site on actin filaments by cryoelectron microscopy and image analysis. J Cell Biol 126:433–443 28. Mermall V, Post PL, Mooseker MS (1998) Unconventional myosins in cell movement, membrane traffic, and signal transduction. Science 23:527–533 29. Morell RJ, Friderici KH, Wei S, Elfenbein JL, Friedman TB, Fisher RA (2000) A new locus for late-onset, progressive, hereditary hearing loss DFNA20 maps to 17q25. Genomics 63:1–6 30. Morín M, Bryan KE, Mayo-Merino F, Goodyear R, MencíaA, Modamio-Høybjør S, del Castillo I, Cabalka JM, Richardson G, Moreno F, Rubenstein PA, Moreno-Pelayo MA (2009) In vivo and in vitro effects of two novel gamma-actin (ACTG1) mutations that cause DFNA20/26 hearing impairment. Hum Mol Genet 18(16):3075–3089 31. Nevalainen EM, Skwarek-Maruszewska A, Braun A, Moser M, Lappalainen P (2009) Two biochemically distinct and tissue-specific twinfilin isoforms are generated from the mouse Twf2 gene by alternative promoter usage. Biochem J 417:593–600 32. Pataky F, Pironkova R, Hudspeth AJ (2004) Radixin is a constituent of stereocilia in hair cells. Proc Natl Acad Sci U S A 101:2601–2606 33. Peng AW, Belyantseva IA, Hsu PD, Friedman TB, Heller S (2009) Twinfilin 2 regulates actin filament lengths in cochlear stereocilia. J Neurosci 29:15083–15088 34. Perrin BJ, Sonnemann KJ, Ervasti JM (2010) β-actin and γ-actin are each dispensable for auditory hair cell development but required for Stereocilia maintenance. PLoS Genet 6(10):e1001158 35. ProcaccioV, Salazar G, Ono S, Styers ML, Gearing M, DavilaA, Jimenez R, Juncos J, Gutekunst C-A, Meroni G, Fontanella B, Sontag E, Sontag JM, Faundez V, Wainer BH (2006) A mutation of beta-actin that alters depolymerization dynamics is associated with autosomal dominant developmental malformations, deafness, and dystonia. Am J Hum Genet 78:947–960 36. Rendtorff ND, Zhu M, Fagerheim T, Antal TL, Jones M, Teslovich TM, Gillanders EM, Barmada M, Teig E, Trent JM et al (2006) A novel missense mutation in ACTG1 causes dominant deafness in a Norwegian DFNA20/26 family, but ACTG1 mutations are not frequent among families with hereditary hearing impairment. Eur J Hum Genet 14:1097–1105 37. Rzadzinska AK, Schneider ME, Davies C, Riordan GP, Kachar B (2004) An actin molecular treadmill and myosins maintain stereocilia functional architecture and self-renewal. J Cell Biol 2004 Mar 15 164(6):887–897 38. Rzadzinska AK, Nevalainen EM, Prosser HM, Lappalainen P, Steel KP (2009) MyosinVIIa interacts with Twinfilin-2 at the tips of mechanosensory stereocilia in the inner ear. PLoS One 4(9):e7097

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39. Schneider ME, Belayantseva IA, Azevedo RB, Kachar B (2002) Rapid renewal of auditory hair bundles. Nature 418:837–838 40. Schneider ME, Dosé AC, Salles FT, Chang W, Erickson FL, Burnside B, Kachar B (2006) A new compartment at stereocilia tips defined by spatial and temporal patterns of myosin IIIa expression. J Neurosci 2640:10243–10252 41. Sekerková G, Zheng L, Loomis PA, Mugnaini E, Bartles JR (2006) Espins and the actin cytoskeleton of hair cell stereocilia and sensory cell microvilli. Cell Mol Life Sci 63:2329–2341 42. Van Camp G, Smith RJ. (2011). Hereditary hearing loss homepage. http://hereditaryhearing loss.org/. Accessed 3 December 2011 43. Vartiainen MK, Sarkkinen EM, Matilainen T, Salminen M, Lappalainen P (2003) Mammals have two twinfilin isoforms whose subcellular localizations and tissue distributions are differentially regulated. J Biol Chem 278:34347–34355 44. Vorobiev S, Strokopytov B, Drubin DG, Frieden C, Ono S, Condeelis J, Rubenstein PA, Almo SC (2003) Structure of nonvertebrate actin. Implications for the ATP hydrolytic mechamism. Proc Natl Sci U S A 100:5760–5765 45. van Wijk E, Krieger E, Kemperman MH, De Leenheer EM, Huygen PL, Cremers CW, Cremers FP, Kremer H (2003) A mutation in the gamma actin 1 (ACTG1) gene causes autosomal dominant hearing loss (DFNA20/26). J Med Genet 40:879–884 46. Yang T, Smith R (2000) A novel locus DFNA26 maps to chromosome 17q25 in two unrelated families with progressive autosomal dominant hearing loss. Am J Hum Genet 67(suppl 2):300 47. Zhu M, Yang T, Wei S, DeWan AT, Morell RJ, Elfenbein JL, Fisher RA, Leal SM, Smith RJ, Friderici KH (2003) Mutations in the gamma-actin gene (ACTG1) are associated with dominant progressive deafness (DFNA20/26). Am J Hum Genet 73:1082–1091

Chapter 10

Therapeutic Targeting of the Actin Cytoskeleton in Cancer Teresa Bonello, Jason Coombes, Galina Schevzov, Peter Gunning and Justine Stehn Abstract In cancer, actin filament populations and associated remodelling proteins are involved in driving proliferation, apoptosis and motility. Furthermore, a web of signalling pathways converge with the actin cytoskeleton to regulate these functions. Importantly, the actin cytoskeleton is a heterogeneous assembly of filament populations, each contributing to shared and unique cellular functions. The current range of actin-disrupting compounds are limited in their therapeutic use as they cannot discriminate between functionally specific populations of actin. Universal disruption of actin is likely to be intolerable in a clinical setting. Dissecting the regulation and composition of these filament populations will allow for treatments tailored to target the unique cytoskeletal repertoire of tumour cells. Identifying specific actin filament populations which are indispensible for tumour cell function is the focus of current work.

10.1

Introduction

Oncogenic cellular transformation is characterised by unrestricted proliferation, inappropriate cell survival and the acquisition of a motile and invasive phenotype. These changes are accompanied by a significant reorganisation of the actin cytoskeleton [1]. In culture, the malignant phenotype can be reversed by reinstating actin filamentstabilising proteins to restore cytoskeletal architecture. This implies a direct role for the actin cytoskeleton in oncogenic signalling [2]. Understanding the role of actin in biological processes deregulated in malignancy, including cell cycle progression, apoptosis and cell motility, presents an opportunity to identify points of vulnerability in a tumour cell for therapeutic targeting. Dissecting the role of actin in these cellular processes has been greatly facilitated by a diverse group of naturally derived compounds which directly target actin. These compounds can be broadly

J. Stehn () · T. Bonello · J. Coombes · G. Schevzov · P. Gunning School of Medical Sciences, University of New South Wales, Sydney, NSW, Australia J. Stehn Department of Pharmacology, School of Medical Sciences, Oncology Research Unit, University of New South Wales, Sydney, NSW 2052, Australia e-mail: [email protected]

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categorised as either inhibitors or stabilisers of actin filament assembly. For example, the cytochalasins prevent polymerisation by binding to the barbed end of actin filaments, inhibiting monomer association and dissociation at this end. Latrunculin cytotoxins associate with only the actin monomer, preventing its incorporation into the growing filament. In contrast, jasplakinolide as well as the more recently identified dolastatin 11, doliculide and hectochlorin, markedly enhance the rate of actin assembly, leading in some instances to the formation of amorphous aggregates of actin in cells [3, 4]. Despite a number of actin-targeting compounds showing potent anti-proliferative and anti-migratory activity, their application has not extended to the clinic [5]. Due to the fundamental role of actin in all cellular systems, the direct targeting of actin in this way does not appear to be a viable therapeutic option at this time. Disrupting contractile function in skeletal and cardiac muscle is of particular concern. Alternatively, targeting actin cytoskeletal regulators altered in malignancy may serve to specifically attenuate cytoskeletal-dependent function in tumour cells. Notably, Rho GTPases have a well-documented role in activating signalling pathways required for actin assembly and organisation [6]. Furthermore, a complex web of actin-regulatory proteins exists with altered expression and activity in cancer, many of which act downstream of the GTPases, Rho, Rac and Cdc42. Actin-regulatory proteins organise the actin cytoskeleton into distinct filament populations which drive specific cellular functions [7]. Therefore, the standing challenge is to identify and disrupt specific actin filaments populations that are indispensible for tumour cell function.

10.2 The Actin Cytoskeleton and Proliferation Malignant growth is characterised by uncontrolled cellular replication, resulting from de-regulation of the cell cycle. The rate of cell replication is dependent on the events of the G1 phase, where extracellular-derived signals from mitogens and adhesion to the extracellular matrix (ECM) promote G1-phase progression [8]. As such, cancer cell proliferation is predominantly determined by dysregulation of the G1 phase [9]. The actin cytoskeleton is heavily involved in the events of G1-phase regulation, and therefore may provide new avenues to target cancer cell proliferation. The actin cytoskeleton appears to play an important part in the transduction of extracellular signals which cause cells to progress through G1 [10]. Here, receptor tyrosine kinases (RTK) and G-protein coupled receptors (GPCR) are activated by soluble factors which, in addition to matrix adhesion, results in an increase in the production of the G1-phase mediator cyclin D1. The formation of the cyclin D1-cdk4/6 (cyclin-dependent kinase) complex leads to phosphorylation of pocket proteins comprising the retinoblastoma protein Rb, p107 and p130. Phosphorylated pocket proteins permit the release of E2F transcription factors, which allows expression of cyclin E [8]. In turn, the cyclin E-cdk2 complex further phosphorylates Rb to a hyperphosphorylated state, resulting in a feedback loop that allows for complete dissociation of Rb from E2F and the transcription of more cyclin E [8]. This results

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in the entrance to S-phase and transcription of genes required for DNA synthesis such as cyclin A [11]. It has long been known that drugs that perturb the actin cytoskeleton cause cells to arrest in G1 phase. Disruption of actin by dihydrocytochalasin B was found to induce arrest in the G1 phase in mouse 3T3 fibroblasts despite stimulation by serum, epidermal growth factor (EGF) and fibroblast growth factor (FGF; [12]). Cytochalasin D treatment of African green monkey cells prevented S-phase entry after serum stimulation [13], and this was associated with cytoplasmic rather than nuclear accumulation of the proto-oncogenes c-fos and c-myc [14]. Disruption by cytochalasin B inhibits G1-phase progression in human capillary endothelial cells via cyclin D1 regulation and expression of the cdk-inhibitor p27, and this was suggested to relate to loss of cytoskeletal tension [15]. Jasplakinolide treatment is also associated with reduced proliferation of prostate carcinoma cells [16] and acute myeloid leukemia cells [17]. Such studies indicate the potential for actin-targeting drugs to inhibit mitogenic cell cycle progression and cancer cell proliferation. That drugs which impact actin in fundamentally different ways (cytochalasin B/D and jasplakinolide) are anti-proliferative, emphasises the importance of intact actin filament dynamics in facilitating proliferation, rather than general toxicity effects of the drugs. Furthermore, it is implied that the dynamicity of actin filaments rather than their presence/ absence is important in mediating cell cycle progression.

10.3

Mitogenic G1 Phase

Clearly, the mitogenic phase of the cell cycle, which temporally spans early-to-mid G1 phase, is an actin cytoskeleton-dependent period. Beyond this point, an intact cytoskeleton is not necessary for progression to S phase [18]. The actin dependence of this period is based on the inter-relationship of the cytoskeleton with signal transduction pathways, particularly the mitogen-associated protein kinase (MAPK) pathway [19], which present prime therapeutic targets for cancer [20]. The initial reception of mitogens, upstream of ERK/MAPK, occurs under the influence of actin. Actin is directly associated with growth factor receptors, including EGF [21–23]. Also, EGF-initiated Phosphatidylinositol 3-kinase (PI3K) activity occurs via association with polymerised actin [24], and PI3K signalling initiated by integrin activation is regulated by the actin cytoskeleton and focal adhesion kinase in thrombin-activated platelets [25]. Upon activation, these growth factor receptors promote G1-phase progression and proliferation. As mutations affecting EGF receptor activity are strongly associated with a range of cancers, there has been strong momentum to target such growth factor receptors with kinase inhibition or antibody blockade strategies [26]. These efforts are currently undergoing a phase of refinement to optimise and improve patient response rates [27]. It would be worth investigating whether modulation of the receptor-actin interaction is an avenue by which EGF-based therapy could be refined.

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Growth factor

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Fig. 10.1 Involvement of the actin cytoskeleton in G1-phase signalling. Regulation of growth factor binding sites and adhesion formation is influenced by actin dynamics. Rho GTPases link adhesion and growth factor signalling. Distinct actin-based regulatory pathways are likely to make differing contributions to G1-phase cyclin expression. For example, Rho/stress fibre/ERK-mediated cyclin D1 expression is distinct from and inhibitory of Rac-mediated cyclin D1 expression. Actindependent cyclin E expression appears to be independent of effects on cyclin D1. This raises the possibility that targeting unique actin filament populations can achieve different anti-proliferative therapeutic effects

10.4

G1-Phase Signalling

Downstream of growth factor receptors actin governs the transduction of signalling cascades that contribute to G1-phase progression. The Rho GTPase family, which regulates the actin cytoskeleton, is also required for cyclin D1 transcription [28]. Furthermore, Rho GTPases link adhesion signalling with cell cycle progression [29]. Cyclin D1 expression is induced by sustained ERK signalling associated with actin stress fibre formation and the subsequently increased intra-cellular tension [30]. This stress fibre formation depends on actin polymerisation and cellular contractility induced by Rho A signalling [28]. Integrin-mediated adhesion and tensional signals are combined with mitogenic signals and transmitted by Rho-GTPases, extracellular signalling-related kinase (ERK1/2), and focal adhesion kinase (FAK) to promote cyclin D1 expression (Fig. 10.1, see [31]). However, treatment with clostridium difficile toxin A, which inactivates Rho GTPases, reveals subtleties to this mechanism [30]. It appears that the Rho-mediated formation of stress fibres controls the timing of cyclin D1 expression to the mid-G1 phase through sustaining ERK activity. In the absence

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of Rho, cyclin D1 expression is brought forward to early G1, and is less robust. The expression of cyclin D becomes mediated by the GTPase Rac, instead of Rho, and is stress fibre and MAPK/ERK independent (Fig. 10.1). Such a cytoskeleton-mediated switch suggests that specific actin filament populations, and signalling by different actin regulators, affect the rate of cell cycle progression. Factors which affect cytoplasmic actin integrity and filament interaction with regulators such as Rho-GTPases would have an impact on the ability of a cell to proliferate, through altering cyclin D1 expression. This could be a useful tool in targeting cell cycle mechanisms in specific cancer cells. Indeed, there is evidence to suggest that pharmacologically inhibiting cellular tension and contractility induced by Rho and MLCK, and the associated ERK signalling, can inhibit malignant transformation [32]. Although cyclin D1-cdk4/6 activity is typically at its peak in mid-G1, late G1-early S phase progression is predominantly associated with the cyclin E-cdk2 complex. In mitogen-stimulated 3T3 fibroblasts, disruption of actin filament organisation by cytochalasin B inhibits the induction of cyclin E, phosphorylation of cdk2, and causes p107, an Rb-family pocket protein which inhibits cell cycle progression, to accumulate in the nucleus [33]. However, in this system, cyclin D1 levels are unaltered. This indicates that the specific actin filament populations involved in cyclin D1 expression may differ to those involved in cyclin E expression. It is possible that Rho-sensitive filaments are more likely to be involved in cyclin D1 but not cyclin E expression. This would also indicate that cyclin E could be regulated via disruption of specific actin filaments. Therefore, there could be opportunities to target different actin filament populations to specifically inhibit different G1-phase cyclins. This would be useful in tailoring specific therapies to cancer cells with mutations that effect distinct cell cycle regulatory profiles. There is also evidence of a shorter, distinct period later in G1 just prior to the restriction point, R. Disruption of this period with cytochalasin D decreases cyclin D1 expression, and increases the cdk-inhibitor p27Kip1 [34]. This observation may provide an avenue by which actin-interrupting drugs could be used to target cancer cells that do not depend on mutations leading to increased MAPK-based signalling, but rather other signalling pathways such as PI3K, or in a combined approach affecting multiple signalling pathways [35, 36]. If actin filaments themselves are crucial components of the G1 phase, factors which control actin dynamics are also likely to govern G1-phase progression. Mutations in the actin depolymerising factor destrin leads to aberrant hyperproliferation in corneal epithelia associated with an accumulation of F-actin [37]. The actin depolymerisation factor cofilin causes G1 arrest when over-expressed, whereas knockdown of cofilin reduces G1 accumulation [38, 39]. Cofilin expression is associated with p27 up-regulation, and repression of cyclin D1/cdk4-mediated Rb phosphorylation. Profilin, a regulator of actin assembly, has a similar effect in that its over-expression also causes G1 arrest via p27 expression in breast cancer cells [40]. Tropomyosins maintain and stabilise actin filaments and their expression is cell cycle regulated. The expression of cancer-associated low-molecular weight tropomyosins is up-regulated in the G1 phase [41]. Malignancy is associated with down-regulation of high molecular weight tropomyosins and therefore an increased

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reliance on actin filaments containing low-molecular weight tropomyosins [42]. One high-molecular weight tropomyosin isoform with known tumour-suppressor function, Tm1, inhibits nuclear accumulation of phosphorylated ERK (p42/p44 MAPK) when over-expressed in transformed fibroblasts [43]. The relative abundance of highversus low-molecular weight tropomyosins may therefore define which actin filament populations are involved in G1-phase progression. Specifically, targeting LMW tropomyosin actin filament populations may avoid toxicity associated with general cytoskeletal disruption. It appears, then, that there are multiple opportunities to exploit the actin cytoskeleton to slow excessive proliferation rates. Within the mechanisms that signal G1-phase progression, alteration of the actin cytoskeleton has the potential to impact extracellular signalling pathways. Useful candidates may include signals resulting from extracellular growth factors and substrate adhesion and in modulating the MAPK cascade to limit the expression of cell cycle regulators.

10.5 The Actin Cytoskeleton and Apoptosis The regulation of tissue homeostasis in a multi-cellular organism depends on maintaining a balance between cell proliferation and programmed cell death. The de-regulation of apoptosis-inducing pathways leading to inappropriate cell survival disrupts this balance, and allows for excessive proliferation or growth in nonphysiological environments. Therefore, the ability of tumour cells to evade apoptosis is a basic requirement for cancer development. Many apoptotic signals converge at the level of the mitochondrion, leading to its permeabilisation and the release of cytochrome c and other apoptogenic factors. This in turn initiates a cascade of apoptotic proteases termed caspases, which effectively act to disassemble the cell by cleaving an array of cellular substrates. It is well established that several morphological changes that occur in apoptotic cells, including cell rounding, nuclear condensation and membrane blebbing, are mediated by the actin cytoskeleton downstream of caspase cleavage [44]. There is accumulating evidence, however, that actin is also involved in pathways required to initiate apoptosis [45]. Understanding the mechanism involved in mammalian cells is an area of active interest and is particularly relevant to the development of anti-cancer therapeutics. Actin itself is a substrate for caspase-dependent cleavage. In vivo the induction of apoptosis is accompanied by actin proteolysis to a 31 kDa (Fractin) and 15 kDa (tActin) fragment [46]. The ectopic expression of tActin in human embryonic kidney cells was found to induce morphological changes characteristic of apoptotic cells [46]. These morphological changes were not accompanied by caspase activation, suggesting the involvement of actin is limited to terminal apoptotic events. However, a more recent study found that upon caspase-mediated cleavage tActin is subject to N-myristolation, and this post-translational modification promotes targeting of tActin to the mitochondria [47]. This finding adds to the emerging hypothesis that actin may directly stimulate apoptotic events upstream of mitochondria.

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The role of actin in mediating mitochondrial events associated with apoptosis has principally been investigated in yeast, which express a single form of actin and therefore represent a highly transparent system for studying actin function. Yeast strains bearing a mutant allele of actin associated with either suppressed or increased actin dynamics, demonstrate a respective loss and gain in cell viability [48]. Importantly, cell viability correlated inversely with mitochondrial ROS production, supporting a role for the actin cytoskeleton in regulating oxidative stress. In more complex eukaryotic models, drugs which manipulate actin dynamics have served as important tools for investigating the relationship between actin and apoptosis. Treatment of the T-cell line CTLL-20 with jasplakinolide, showed that actin stabilisation both accelerated and amplified the apoptotic response induced by cytokine deprivation [49]. Moreover susceptibility to jasplakinolide-induced cell death has been reported in several transformed murine cell lines in the absence of other apoptotic stimuli [50]. Specifically, an increase in caspase-3-like activity has been shown to precede jasplakinolide-induced apoptosis and could be abrogated by pre-treatment with caspase inhibitors [50] or over-expressing the anti-apoptotic protein Bcl-xL [49]. Studies with jasplakinolide therefore support the notion that actin plays a role in the initial phase of cell death, where cells are committed to the apoptotic programme. Inhibiting actin polymerisation with latrunculin-A was shown to promote a rounded cell morphology and concomitant apoptosis in non-tumourigenic mammary epithelial cells. Interestingly, while changes to cell shape were also observed in MDA-MB-231 metastatic breast tumour cells treated with latrunculin-A, cell survival was not compromised. In a cytoskeleton-based genetic screen of MDA-MB-231 cDNA, Bcl-xL was identified as the anti-apoptotic factor responsible for conferring resistance to latrunculin A-induced cell death [51]. While there is evidence to support both actin filament stabilisation and destabilisation in stimulating apoptosis, it is likely that these differences reflect variability in the ratio of monomeric to filamentous actin between cell types [52]. It is therefore not surprising that several actin-binding proteins which function to regulate this dynamic equilibrium between G-actin and F-actin states have also been implicated in apoptosis (Fig. 10.2).

10.6 The Role of Actin-Binding Proteins in Apoptosis Translocation of cofilin to mitochondria was shown to be an early response to apoptosis-inducing stimuli [53]. While localisation of cofilin to mitochondria occurs independently of actin-binding, a functional actin-binding domain is necessary to trigger apoptosis [53]. Thus, it appears the actin-regulatory activity of cofilin is pivotal in the progression of apoptosis. Specifically, oxidant-induced apoptosis has been shown to mediate mitochondrial damage by cofilin oxidation. Here, it was found that only the active, oxidised form of cofilin was able to open the mitochondrial permeability transition pore to induce cytochrome c release [54]. A pro-apoptotic

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Cytotoxic Stress

? F-actin

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Pro-apoptotic factors e.g. Bax Bim Bid Bmf

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cofilin

Cytochrome-c release Caspase activation Substrate Proteolysis e.g. actin, gelsolin

Fig. 10.2 Relationship between actin remodelling and the intrinsic apoptotic pathway. In response to an apoptotic cue, proteins of the BH3-only family (such as Bid, Bim and Bmf) stimulate the translocation of Bax family members to the mitochondrial outer membrane. This leads to membrane permeabilisation and subsequent cytochrome c release which can be antagonised by the Bcl-2 family of anti-apoptotic factors. The actin remodelling proteins gelsolin and cofilin have also been shown to directly regulate mitochondrial integrity to inhibit and promote apoptosis, respectively. Caspase activation, downstream of cytochrome c release, results in the cleavage of a number of substrates leading to controlled demolition of the cell. A heightened sensitivity to apoptotic stimuli has been observed in a number of cell types where the integrity of the actin cytoskeleton is compromised. In some instances cytoskeletal stress alone is sufficient to induce apoptosis. While it is known that disruption of the actin cytoskeleton leads to the release of Bmf, its contribution to other signalling pathways which converge on mitochondria is yet to be defined

role for cofilin is further supported by the observation made by Bamburg et al. [55], that cofilin is sequestered into cofilin-actin bundles in neurites under conditions of transient stress. The short-term appearance of these rod-like structures was beneficial to cell survival by slowing the loss of mitochondrial membrane potential, possibly reflecting the lack of free cofilin available for translocation to the mitochondria [55]. The actin filament severing protein gelsolin is a known substrate for caspase-3 activity in vitro. Expression of the N-terminal cleavage product in mouse embryonic fibroblasts was shown to accelerate morphological changes associated with apoptosis by severing actin filaments in an unregulated manner [56]. Upstream of mitochondria, however, a pro-survival role for gelsolin has been described. Jurkat cells over-expressing full length gelsolin demonstrated resistance to a broad range of apoptosis-inducing agents [57]. Gelsolin was shown to effectively stabilise the mitochondrial membrane potential and block cytochrome c release by inhibiting the voltage-dependent anion channel (VDAC) of the mitochondrial permeability transition pore [57, 58]. How the function of gelsolin, which extends beyond apoptosis, correlates with its expression in cancer is yet to be reconciled. From

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a clinical perspective gelsolin has been characterised as a putative tumour suppressor. Gelsolin is down-regulated in a number of non-invasive cancer cell types [59] and transfection of gelsolin cDNA in human bladder cancer cells markedly reduced tumourigenicity in vivo [60].

10.7 Anoikis In epithelial cells, signals required for survival are transduced through their interaction with the ECM. A loss of integrin-mediated adhesion signals results in a specific form of apoptosis termed anoikis. Tumour cells acquire a resistance to anoikis, enabling them to survive and proliferate independently of ECM attachment. Anchorage independence in tumour cells is also an essential feature for metastasis [61]. There is evidence to suggest the integrity of the actin cytoskeleton is important for mediating the anchorage-independent phenotype. Restoring expression of the tropomyosin Tm1 in MDA-MB231 breast cancer cells, led to a significant reorganisation of the cytoskeleton accompanied by resensitisation to anoikis. Moreover, treatment with latrunculin-A effectively rescued MDA-MB231 cells from Tm1-induced anoikis, supporting a role for the actin cytoskeleton in adhesion-dependent signalling [62]. Understanding the mechanisms which trigger anoikis, may foster new therapeutic approaches aimed at restoring anoikis sensitivity in tumour cells. While it is clear that manipulation of the actin cytoskeleton by several factors is necessary for apoptosis to proceed, the exact function of actin in apoptosis remains elusive. The actin cytoskeleton has been shown to directly regulate the activity of pro-apoptotic factor Bmf, which is sequestered by the myosin V motor protein and released in response to apoptotic stimuli including anoikis [63]. One hypothesis is that the actin cytoskeleton may serve as a physical linker between the cytosol and mitochondria to facilitate the recruitment of pro-apoptotic proteins [64]. In support of this, apoptosis-induced accumulation of β-actin in the mitochondria was shown to occur before mitochondrial insertion of Bax and cytochrome c release [64].

10.8 Actin Cytoskeleton and Motility Aberrant cellular movement is a hallmark feature of tumourigenesis, which facilitates the propagation of tumour cells to distant sites in the body. The formation of secondary metastatic tumours, which are refractory to conventional therapies, account for the majority of cancer mortalities. This reflects an obvious deficit in chemotherapeutic agents designed to target specific steps in the metastatic pathway. Tumour cells with an elongated morphology adopt a mesenchymal mode of migration which requires dynamic remodelling of the actin cytoskeleton. Here, cells acquire a polarised morphology in response to extracellular signals characterised by the development of leading edge protrusions. The actin cytoskeleton is anchored to

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Fig. 10.3 Key events in the motility cycle driven by actin remodelling. The mesenchymal mode of cell motility in a three-dimensional environment is characterised by a five-step model. In response to a directional cue, cells extend leading edge protrusions (1) which weakly adhere to the underlying substratum (2). Ventral protrusions termed invadopodia remodel the extracellular matrix by focalised proteolysis (3). The contraction of stress fibres tethered to stable adhesions results in the translocation of the cell body (4) followed by disassembly of adhesions at the trailing edge (5). Each stage is associated with a unique profile of actin-binding proteins to yield actin filament structures which fulfil distinct functional outcomes. Actin-binding proteins which serve to integrate several stages in the motility cycle may represent robust targets for the design of anti-metastatic agents (see text)

the ECM by adhesions which stabilise protrusions and serve as traction points. Actomyosin contraction generates the tensile force against adhesions to promote forward translocation of the cell body and tail retraction (Fig. 10.3). Amoeboid migration is an alternative mode of migration used by some tumour cell types which describes cell translocation by alternating cycles of morphological expansion and contraction, driven by cortical actin reorganisation [65].

10.9

Leading Edge Protrusions—Lamellipodia

The leading edge of motile cells is characterised by flat membranous lamellipodial protrusions, behind which extends the lamellae. Protrusive force is driven by localised actin polymerisation at the plasma membrane. Two key actin-binding proteins, the Arp2/3 complex and cofilin, are implicated in the treadmilling of F-actin within the lamellipodia. The Arp2/3 complex generates a branched actin network by initiating filament formation on the sides of pre-existing actin filaments. The barbed end of the new filament is orientated towards the plasma membrane and is available for filament elongation [66–68]. The potent severing activity of cofilin functions synergistically with the Arp2/3 complex by increasing the availability of polymerised actin filaments for dendritic nucleation [69, 70].

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Arp2/3-complex function is a basic requirement for lamellipodia extension [71, 72] and is regulated by WAVE2 [73, 74], a nucleation-promoting factor of the Wiskott-Aldrich syndrome protein (WASP) family acting downstream of Rac. The expression of both Arp2 and WAVE2 has been shown to correlate with a greater metastatic risk in several cancer types [75–77] and may therefore represent viable therapeutic targets. In support of this, RNAi-mediated knockdown of WAVE2 significantly impaired invasion and pulmonary metastasis of B16F10 melanoma cells [78]. Genes coding for the p41 subunit of human Arp2/3 complex have also been identified within a locus recurrently amplified in pancreatic cancer. The silencing of ARPC1A in the AsPC-1 pancreatic cell line led to a considerable reduction in cell migration and invasion [79]. Highly localised activation of cofilin near the plasma membrane is required for lamellipodial protrusion and directed cell motility [80, 81]. This is exemplified in mammary tumour cells stimulated with epidermal growth factor, where the local release of active cofilin following phosphatidylinositol 4, 5-bisphosphate (PIP2 ) hydrolysis is accompanied by the global activation of LIM Kinase 1 (LIMK1). LIMK1 inactivates cofilin that has diffused away from the membrane to spatially restrict its activity [82, 83]. Therefore, the synchronised activation and inactivation of cofilin results in a spatially localised increase in the number of actin filament barbed ends. In line with this, components from both the stimulatory and inhibitory branch of the cofilin activity cycle are found to be simultaneously over-expressed in a number of invasive carcinomas [84, 85]. Therefore, it appears that the invasive and metastatic behaviour of tumour cells requires an optimal balance between the expression of cofilin and its regulatory components [84]. Therapeutic intervention should therefore aim to disrupt this balance and ultimately reduce the output of the cofilin activity cycle, that is to say, the formation of free barbed ends primed for actin polymerisation.

10.10 Leading Edge Protrusions—Filopodia Probing the local micro-environment for directional cues is thought to be mediated by filopodia—finger-like projections containing long parallel bundles of actin filaments. The convergent elongation model of filament formation suggests that filopodial actin filaments are derived from reorganisation of the lamellipodial meshwork. Here, Arp2/3-generated actin filaments gain protection by vasodilator-stimulated phosphoprotein (VASP) against capping allowing them to elongate persistently at their barbed ends. These actin filaments coalesce at the cell edge and are subsequently reorganised to filopodia by bundling at their vertices [86, 87]. Fascin has been described as the major actin bundler in filopodia. In addition to initiating filopodia formation in mammalian cells, fascin was shown to impart rigidity upon actin filaments, allowing an emerging filopodium to overcome the compressive force of the cell membrane [87]. The role of fascin in promoting long protrusions, as well as a recently described role in stabilising actin in invadopodia [88], was shown to facilitate 3D matrix invasion in melanoma cell types utilising the mesenchymal but not the amoeboid mode of migration [88].

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The role of fascin in cell motility and metastasis has been explored in a number of models. Fascin has been shown to drive a more motile phenotype when overexpressed in colorectal adenoma-derived cell lines [89], while fascin knockdown in glioma cells reduced the formation of filopodia and restricted their migratory and invasive capacity in vitro [90]. Fascin-depleted DU145 prostate cells implanted in a murine prostate model dramatically reduced the formation of lymph node metastases [91]. A noteworthy observation is that fascin levels are low or absent in normal adult epithelia but are dramatically up-regulated in many human carcinomas [92]. Fascin therefore presents an exciting opportunity to develop chemotherapeutics that exclusively target the tumour, thereby alleviating patient morbidity associated with non-specific toxicity. Importantly, fascin expression has been reported to correlate with clinical aggressiveness in a number of cancers [93]. In a recent study into the molecular basis behind the potent anti-metastatic activity of migrastatin analogues, macroketone was found to bind to an actin-binding site of fascin to significantly reduce its F-actin bundling activity [94]. This provides an exciting insight into the value of compounds which target key modulators of the actin cytoskeleton. With the finding that filopodia can form in the absence of the Arp2/3 complex [72], de novo filament nucleation by formins has been proposed as an alternative mechanism for filopodia formation. The mammalian family of formin isoforms are characterised by a highly conserved formin homology 2 (FH2) domain. The FH2 domain facilitates the nucleation of a nascent actin filament and promotes elongation at the barbed end to produce long, parallel filaments which are often bundled in cells [95]. One member of the family of diaphanous-related formins, mDia2, is recruited to the cortical actin network where it promotes filopodia formation downstream of the Rho GTPases Cdc42 and Rif [96, 97]. Accordingly, siRNA knockdown of mDia2 impaired filopodia formation in the B16-F1 melanoma cell line [98]. In the context of cellular motility, recent data implicate mDia2 within a signalling pathway which regulates the amoeboid phenotype [99]. Non-apoptotic membrane blebbing induced in the DU145 prostate cancer cell line with epidermal growth factor stimulation could be amplified by targeted knockdown of human mDia2. In support of this, human prostate tumours profiled for chromosomal alterations at the locus encoding mDia2 revealed a higher frequency of deletions in metastases compared to the primary tumour [99]. In another study, systematic analysis of all 15 human formins using siRNA knockdown revealed Formin-Like 2 (FMNL2), acting downstream of Rho C, was specifically involved in promoting the amoeboid invasive phenotype of MDA-MB-435 melanoma cells [100]. This supports a positive correlation previously established between the expression of FMNL2 in primary colorectal tumours and lymphatic metastasis [101]. A recently identified small molecule inhibitor of formin FH2 domain-mediated actin assembly (SMIFH2) was found to significantly impair migration rates in NIH 3T3 fibroblasts accompanied by a reduction in the number of cells extending a lamellipodia. This morphological phenotype was not observed in the lung carcinoma cell line A549, where dynamic bleb-like protrusions were observed near the cell periphery with SMIFH2 treatment [102]. These data indicate that general inhibition of all formin isoforms is likely not a suitable therapeutic strategy, as it cannot

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account for the relative contribution of different isoforms to migration or changes in the expression of formin isoforms which occur in malignancy.

10.11 ECM Remodelling In order to infiltrate surrounding tissue, invasive carcinoma subtypes utilise actinrich protrusions known as invadopodia to proteolytically degrade the ECM. In the highly metastatic carcinoma cell line, MTLn3, formation of invadopodia was shown to depend on the N-WASP-Arp2/3 pathway but not on cofilin [103, 104]. Rather, suppression of cofilin by siRNA reduced invadopodia lifetime and matrix degradation activity [104]. Recent work suggests the involvement of Arp2/3 and cofilin at distinct stages of invadopodia assembly and maturation is orchestrated by the cytoskeletal protein cortactin [105]. This finding points to a scaffold function for cortactin where it is known to bind and activate an array of proteins localised to invadopodia including Arp2/3 [106, 107] and the Arp2/3 nucleator N-WASP [108]. In addition to regulating actin dynamics, cortactin is essential for trafficking key invadopodial proteases involved in matrix degradation [109]. Targeting cortactin for knockdown, as reported in several transformed cell types, impairs both invadopodia formation and function [109–111] and therefore may translate to an effective therapeutic strategy. Importantly, the expression of cortactin has been shown in pre-clinical models to correlate with enhanced metastasis in breast [112], hepatocellular [113] and esophageal squamous cell [114] carcinomas.

10.12 Cell Traction and Translocation Following the extension of polarised protrusions at the leading edge, the intra-cellular actin cytoskeleton is coupled to the ECM by integrin-rich adhesions. In the lamella, focal adhesions tether contractile actin filament bundles termed stress fibres to the substratum to facilitate traction of the cell body. Stress fibre contraction is mediated by the motor protein myosin II, which slides anti-parallel actin filaments past each other to generate force. Tension exerted on focal adhesions promotes forward translocation of the cell body and retraction of the cell’s trailing end [115]. Furthermore, myosin-II driven contractility appears to act as a restraint on lamellipodial protrusion. Although myosin II is absent from the lamellipodium, siRNA knockdown or pharmacological inhibition of myosin II is reported to increase the rate of cellular protrusion at the leading edge [116, 117]. One hypothesis is that myosin II-mediated retrograde flow of actin in the lamellum counters actin polymerisation at the leading edge thereby reducing the net protrusion rate. Adhesions might act to modulate these opposing forces by serving as traction points, diverting the forces created by the retrograde flow of actin to the substratum [118]. Therefore, myosin-II contractility integrates spatially segregated activities within the motility cycle and may represent a robust therapeutic target. The challenge remains in understanding

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how to manipulate myosin II activity with the outcome of impairing cell migration in vivo. In a similar way to myosin, tropomyosin has been identified as a key regulator of the spatiotemporal interplay between the lamellipodium and lamella [119, 120]. In epithelial cells injected with skeletal muscle tropomyosin, the lamellipodium was replaced by a lamellar-like actin array, characterised by the absence of a dendritic network, depleted Arp2/3 and cofilin from the leading edge and gained myosin-II-dependent retrograde F-actin flow. Changes to morphology and F-actin dynamics were accompanied by increased protrusion persistence and migration [120]. Tropomyosin has been shown to account for these changes by inhibiting the rate of Arp2/3-dependent branch formation and antagonising the depolymerising effect of ADF to slow the rate of filament turnover [119]. Finally, the reliance of cancers on a restricted repertoire of tropomyosin isoforms may impact the plasticity of the actin cytoskeleton. Interest in the relationship between tropomyosin expression and metastasis is only starting to gain momentum. A recent study found that reinstating the HMW tropomyosin Tm3 in MDA-MB-231 breast cancer cells greatly reduced the formation of lung metastases [121]. Understanding the mechanisms which drive the down-regulation of HMW tropomyosins may help forge new therapeutic strategies for the prevention of metastatic disease [42].

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97. Peng J, Wallar BJ, Flanders A, Swiatek PJ, Alberts AS (2003) Disruption of the Diaphanousrelated formin Drf1 gene encoding mDia1 reveals a role for Drf3 as an effector for Cdc42. Curr Biol 13:534–545 98. Yang C, Czech L, Gerboth S, Kojima S, Scita G, Svitkina T (2007) Novel roles of formin mDia2 in lamellipodia and filopodia formation in motile cells. PLoS Biol 5:e317 99. Di Vizio D, Kim J, Hager MH, Morello M, Yang W, Lafargue CJ, True LD, Rubin MA, Adam RM, Beroukhim R, Demichelis F, Freeman MR (2009) Oncosome formation in prostate cancer: association with a region of frequent chromosomal deletion in metastatic disease. Cancer Res 69:5601–5609 100. Kitzing TM, Wang Y, Pertz O, Copeland JW, Grosse R (2010) Formin-like 2 drives amoeboid invasive cell motility downstream of RhoC. Oncogene 29:2441–2448 101. Zhu XL, Liang L, Ding YQ (2008) Overexpression of FMNL2 is closely related to metastasis of colorectal cancer. Int J Colorectal Dis 23:1041–1047 102. Rizvi SA, Neidt EM, Cui J, Feiger Z, Skau CT, Gardel ML, Kozmin SA, Kovar DR (2009) Identification and characterization of a small molecule inhibitor of formin-mediated actin assembly. Chem Biol 16:1158–1168 103. Lorenz M, Yamaguchi H, Wang Y, Singer RH, Condeelis J (2004) Imaging sites of N-wasp activity in lamellipodia and invadopodia of carcinoma cells. Curr Biol 14:697–703 104. Yamaguchi H, Lorenz M, Kempiak S, Sarmiento C, Coniglio S, Symons M, Segall J, Eddy R, Miki H, Takenawa T, Condeelis J (2005) Molecular mechanisms of invadopodium formation: the role of the N-WASP-Arp2/3 complex pathway and cofilin. J Cell Biol 168:441–452 105. Oser M, Yamaguchi H, Mader CC, Bravo-Cordero JJ, Arias M, Chen X, Desmarais V, van Rheenen J, Koleske AJ, Condeelis J (2009) Cortactin regulates cofilin and N-WASp activities to control the stages of invadopodium assembly and maturation. J Cell Biol 186:571–587 106. Uruno T, Liu J, Zhang P, Fan Y, Egile C, Li R, Mueller SC, Zhan X (2001) Activation of Arp2/3 complex-mediated actin polymerization by cortactin. Nat Cell Biol 3:259–266 107. Weaver AM, Karginov AV, Kinley AW, Weed SA, LiY, Parsons JT, Cooper JA (2001) Cortactin promotes and stabilizes Arp2/3-induced actin filament network formation. Curr Biol 11:370– 374 108. Martinez-Quiles N, Ho HY, Kirschner MW, Ramesh N, Geha RS (2004) Erk/Src phosphorylation of cortactin acts as a switch on-switch off mechanism that controls its ability to activate N-WASP. Mol Cell Biol 24:5269–5280 109. Clark ES, Whigham AS,Yarbrough WG, Weaver AM (2007) Cortactin is an essential regulator of matrix metalloproteinase secretion and extracellular matrix degradation in invadopodia. Cancer Res 67:4227–4235 110. Artym VV, ZhangY, Seillier-Moiseiwitsch F,Yamada KM, Mueller SC (2006) Dynamic interactions of cortactin and membrane type 1 matrix metalloproteinase at invadopodia: defining the stages of invadopodia formation and function. Cancer Res 66:3034–3043 111. Webb BA, Jia L, Eves R, Mak AS (2007) Dissecting the functional domain requirements of cortactin in invadopodia formation. Eur J Cell Biol 86:189–206 112. Li Y, Tondravi M, Liu J, Smith E, Haudenschild CC, Kaczmarek M, Zhan X (2001) Cortactin potentiates bone metastasis of breast cancer cells. Cancer Res 61:6906–6911 113. Chuma M, Sakamoto M, Yasuda J, Fujii G, Nakanishi K, Tsuchiya A, Ohta T, Asaka M, Hirohashi S (2004) Overexpression of cortactin is involved in motility and metastasis of hepatocellular carcinoma. J Hepatol 41:629–636 114. Luo ML, Shen XM, Zhang Y, Wei F, Xu X, Cai Y, Zhang X, Sun YT, Zhan QM, Wu M, Wang MR (2006) Amplification and overexpression of CTTN (EMS1) contribute to the metastasis of esophageal squamous cell carcinoma by promoting cell migration and anoikis resistance. Cancer Res 66:11690–11699 115. Vicente-Manzanares M, Ma X, Adelstein RS, Horwitz AR (2009) Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat Rev Mol Cell Biol 10:778–790 116. Ponti A, Machacek M, Gupton SL, Waterman-Storer CM, Danuser G (2004) Two distinct actin networks drive the protrusion of migrating cells. Science 305:1782–1786

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Part III

Microtubules and Disease

Chapter 11

Microtubules as a Target in Cancer Therapy April L. Risinger and Susan L. Mooberry

Abstract Drugs that target cellular microtubules are some of the most important agents used in the treatment of adult and pediatric cancers. This chapter aims to provide an overview of diverse microtubule targeting agents, including microtubule stabilizers and destabilizers, and describes how these agents bind to tubulin/ microtubules and suppress microtubule dynamics leading to defects in mitotic spindle function, mitotic arrest, and initiation of apoptosis. Although the microtubule and antimitotic effects of these agents are the focus of this chapter, there is evidence to suggest that the anticancer actions of these agents are multifactorial, which may contribute to their pleiotropic biological activities. The emphasis of this chapter is on microtubule targeting drugs that are currently used clinically for cancer therapy as well as agents that are being evaluated in clinical trials. A few agents in preclinical development that have a unique binding site or mechanism of action are also described.

11.1

Introduction

Microtubules are dynamic cytoskeletal structures that play essential and diverse roles in cellular homeostasis, from motor-driven intracellular transport to maintenance of cell shape and motility. Interphase microtubules nucleate from the microtubule organizing centers in animal cells and form a radial network of dynamic structures (Fig. 11.1a). Arguably, one of the most important roles for microtubules occurs during mitosis when they form the specialized structure of the mitotic spindle. The mitotic spindle first aligns chromosomes in the metaphase plate and then separates and evenly distributes the sister-chromatids into daughter cells. The process of mitosis is complex and finely tuned, involving the temporal and spatial coordination of hundreds of proteins. The normal bipolar mitotic spindle (Fig. 11.1b) consists of microtubules nucleated from both centrosomes and chromatin/kinetochores. Mitotic spindles generate tension through dynamic instability and treadmilling in a tightly regulated manner in cooperation with microtubule motors and the spindle matrix S. L. Mooberry () · A. L. Risinger Departments of Pharmacology and Medicine, University of Texas Health Science Center at San Antonio, 7703 Floyd Curl Drive, San Antonio, TX 78229, USA e-mail: [email protected]

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Fig. 11.1 Antimitotic mechanisms of action of microtubule targeting agents. At clinically relevant concentrations, microtubule stabilizers and destabilizers suppress microtubule dynamics. As drug-treated interphase cells a enter mitosis, b the microtubule dynamics of the mitotic spindle are inhibited, precluding formation of a normal metaphase plate. As a result, the cells are unable to progress through a normal anaphase c and instead arrest in mitosis with multiple aberrant mitotic spindles d. Drug-treated cells are unable to complete normal mitosis and ultimately undergo apoptosis

to move each sister-chromatid to the appropriate pole (Fig. 11.1c). The fidelity of mitosis is essential for life of multicellular organisms and cell cycle checkpoints control various stages of mitosis to insure accuracy. Lack of mitotic accuracy has been implicated in the pathogenesis of cancer and failure at any stage of mitosis can lead to initiation of apoptosis to prevent loss of genetic fidelity. Cancer is a disease of uncontrolled proliferation. The ability to interfere and prevent mitosis, thus halting cell proliferation, is the basis for the development of antimitotic anticancer drugs. A number of antimitotic targets have been evaluated, however, to date; only agents that bind to tubulin have achieved clinical utility. Many structurally diverse compounds, mostly derived from nature, bind to cellular tubulin/microtubules and inhibit normal microtubule dynamics. Inhibition of microtubule dynamics during mitosis causes severe mitotic defects including metaphase arrest and the formation of aberrant multipolar mitotic spindles (Fig. 11.1d) that preclude normal mitotic progression. Mitotic failure eventually leads to cell death. Multiple binding sites for microtubule targeting agents have been identified on the αβ-tubulin heterodimer, and on the interior and the exterior of the formed microtubule. A schematic of the interaction of the microtubule targeting agents discussed in this chapter with tubulin/microtubules is shown in Fig. 11.2. Microtubule targeting agents have played a significant role in the treatment of cancer since the early 1960s when the vinca alkaloids were approved for cancer therapy. Our goal in this chapter is to provide an overview of the mechanisms of action of this chemically diverse

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Fig. 11.2 Drug-binding sites on tubulin/microtubules a Microtubule-destabilizing agents. Colchicine-site agents bind to the tubulin heterodimer at the αβ, dimer interface to form a complex that is incorporated into the growing microtubule. Vinca-site agents bind to the β-tubulin subunit at the exposed plus end of the microtubule. b Microtubule-stabilizing agents. Taxane-site agents bind to the β-tubulin subunit on the interior of the intact microtubule. Recent data suggest that laulimalide-site agents bind to the β-tubulin subunit on the exterior of the intact microtubule

class of drugs and their utility in treating cancer. Agents that have failed to advance through clinical testing and some new mechanistically diverse classes of compounds that have potential for clinical development are also described.

11.2 Vinca Alkaloids The vinca alkaloids were the first class of microtubule targeting agents used for the treatment of cancer. Vincristine and vinblastine, the first vinca alkaloids identified, were isolated from the periwinkle plant, Catharanthus roseus, in the late 1950s. Soon after the serendipitous observation that these compounds caused bone marrow suppression in mice, intense efforts were undertaken to understand their mechanism of action and to develop vinblastine and vincristine as anticancer agents [1]. Vincristine (Oncovin) and vinblastine (Velban) were approved by the FDA in the early 1960s for the treatment of multiple hematological malignancies, germ cell cancers, and a few types of solid tumors. Since then, two second-generation semisynthetic vinca alkaloids, vinorelbine (Navelbine) and vindesine (Eldisine), were approved for the treatment of non-small-cell lung cancer and melanoma, respectively. A thirdgeneration semisynthetic bifluorinated alkaloid, vinflunine, was approved for bladder cancer in the EU and remains in clinical development in the United States [2, 3]. While the vinca alkaloids are useful in curative combination therapy in certain tumors, they are not a mainstay of therapy for most common adult solid tumors. Biochemical experiments show that the vinca alkaloids bind directly to the αβtubulin heterodimer in solution or in microtubules. Each αβ-tubulin heterodimer

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contains a vinca-binding site near the hydrolyzable GTP site on the β-subunit [4]. However, on the intact microtubule there are two distinct locations where the vinca alkaloids can bind, at the plus end of the microtubule or along the microtubule protofilaments. The binding of vinca alkaloids to β-tubulin at each of these locations on the microtubule causes distinct effects on microtubule dynamics and structure. The vincas bind with a high affinity to the exposed β-tubulin subunit at the plus end of the microtubule [5]. Vinca alkaloid binding to the end of microtubules inhibits microtubule dynamics, resulting in this class of drugs being referred to as “end poisons.” When vinblastine is added at substoichiometric concentrations to tubulin, approximately one molecule of drug binds to the plus end of each microtubule [6]. This high-affinity binding to the most dynamic portion of the microtubule suppresses microtubule dynamics by decreasing the rate of microtubule growth and shortening and can inhibit treadmilling up to 50% without producing significant changes in tubulin polymer [5, 7]. However, when the vinca alkaloids are added at near-stoichiometric concentrations to free tubulin, the drug also binds to β-tubulin subunits along the exterior of the intact microtubule with a ratio of approximately 16 molecules of drug per microtubule [8]. Vinca binding along microtubule protofilaments directly destabilizes microtubules, resulting in net loss of polymerized tubulin. Studies measuring microtubule dynamics in intact cells demonstrate that the vinca alkaloid-induced inhibition of microtubule dynamics observed in biochemical experiments is mirrored in cellular microtubules [9]. The vinca alkaloids are classified as microtubule destabilizers because they cause dramatic loss of cellular microtubules at concentrations above 10 nM. Not surprisingly, at this concentration mitotic spindles fail to form normally and cells are unable to complete a normal mitosis [10]. However, concentrations of vinblastine as low as 0.8 nM can cause mitotic arrest without any observable changes in microtubule mass [10]. Therefore, at low, clinically relevant concentrations, the vinca alkaloids lead to mitotic arrest and apoptosis of rapidly dividing cancer cells as a direct result of their suppression of microtubule dynamics in a manner that is independent of changes in gross microtubule structure [7]. The vinca alkaloids present interesting structure-activity relationships. Vincristine differs from vinblastine only by the replacement of a methyl group with a formyl group on the vindoline moiety, however, the clinical indications and dose-limiting toxicities of these two drugs are different. Vinblastine is approved for use in solid tumors with a dose-limiting toxicity of neutropenia, whereas vincristine is primarily used in childhood hematological cancers with a dose-limiting toxicity of peripheral neuropathy. Second- and third-generation semisynthetic vinca alkaloids also demonstrate efficacy and toxicity profiles that are distinct from their natural precursors or each other. Why such structurally similar drugs have different clinical indications and limiting toxicities is not known. Preclinical studies indicate that vinflunine and vinorelbine have slightly different effects on the suppression of microtubule dynamics as compared to vinblastine [11]. Vinflunine binds tubulin with a lower affinity than other vinca alkaloids, however, is concentrated and retained in cells to a greater extent than other family members [12]. Vinflunine is also the first vinca alkaloid to demonstrate vascular disrupting activity as evidenced by rapid destruction of tumor vasculature within 2 hours of drug administration at a submaximally tolerated dose,

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a property that has previously only been demonstrated with a subset of microtubule targeted agents that bind to the colchicine site on tubulin [13]. Studies to elucidate the complicated structure-activity relationships for this multifunctional class of molecules could provide valuable insight about the drug properties that are optimal for treating specific cancers.

11.3

Other Vinca-Site Binding Agents

In addition to the third-generation vinca alkaloid, vinflunine, other chemically diverse agents that bind to the vinca site on tubulin are also in clinical development. Eribulin (E7389, eribulin mesylate) is a highly simplified synthetic analog of themarine natural product halichondrin B, which was originally isolated from the sponge Halichondria okadai. Eribulin is a microtubule destabilizer that noncompetitively inhibits the binding of vinblastine to the plus end of microtubules [14, 15]. The effects of eribulin on microtubule dynamics vary slightly in comparison to vinblastine in that eribulin inhibits microtubule growth, however, has little effect on microtubule shortening [16]. Vinflunine also has minimal effects on microtubule shortening when compared with other vinca alkaloids [7]. In spite of these minor differences on microtubule growth and shortening, eribulin and each of the vinca alkaloids share a common mechanism of binding to the microtubule plus end, inhibiting microtubule dynamics and causing mitotic arrest and apoptosis. At higher concentrations each of these compounds causes loss of cellular microtubules. Eribulin showed anticancer activity in heavily pretreated patients accompanied by a low incidence of neuropathy and was approved for the treatment of drug-resistant, metastatic breast cancer in late 2010 [3, 17]. Other vinca-binding agents in clinical development include maytansinoidantibody conjugates [18]. As a single agent, maytansine failed in clinical trials in the 1970s due to dose-limiting gastrointestinal and central nervous system toxicities [19]. The maytansine derivative DM1 has been conjugated to various antibodies to allow tumor cell targeting of this potent cytotoxin. Trastuzumab-DM1(t-DM1) is currently undergoing clinical trials against HER2-expressing breast cancers [3, 20]. MLN2704A, a fusion of DM1 to a prostate-specific membrane antigen (PSMA) antibody, is also in clinical development against hormone refractory prostate cancer [3, 21]. The idea of using antibodies to target cytotoxic agents specifically to cancer cells is a logical next step in the more effective use of microtubule targeted agents. However, it is unclear what the most effective mechanism of tumor targeting will be and whether a cytotoxic agent such as DM1, that has not proved its efficacy as a single agent, is the best compound to test this strategy. A number of other vinca site-binding agents have entered clinical trials and failed to advance to approval for clinical use due to lack of anticancer efficacy or for undisclosed reasons. The dolastatins 10 and 15 are small peptides first identified from the marine mollusk Dolabella auricularia [22]. The biosynthetic source of these peptides was later found to be cyanobacteria that are presumably ingested by the sea hares [23]. Dolastatin 10, the dolastatin 10 analog TZT-1027, and the dolastatin 15

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analogs cemadotin and tasadotin hydrochloride were all evaluated in a number of clinical trials and either failed to advance or did not produce objective anticancer responses [24, 25]. The cryptophycins and hemiasterlins are two additional classes of potent microtubule inhibitors that bind within the vinca site on tubulin [26]. Cryptophycin 52 and two hemiasterlin derivatives, E7974 and HTI-286, were evaluated in early phase clinical trials, however, failed to advance [27, 28]. It is interesting to note that the dolastatins, cryptophycin 52 and the hemiasterlins all bind within what has been referred to as the peptide groove within the vinca-binding site [29, 30]. To date all of the agents that bind within this region have failed in clinical development. Whether this relates to occupancy of this region as compared to the overlapping site occupied by the vinca alkaloids, or the physiochemical properties of the agents evaluated is not known. It is interesting to speculate that there may be undiscovered downstream differences caused by occupancy of these two regions within the vinca-binding site that translate into differential anticancer effects. A full understanding of the nature of these differences may streamline the development of effective anticancer pharmaceuticals in the future.

11.4 Taxanes Taxol was first isolated in 1967 from the bark of the Pacific Yew, Taxus brevifolia, as part of an initiative by the National Cancer Institute (United States) to identify novel antitumor agents from plants [31]. The intense interest in taxol as a potential anticancer agent did not occur until Susan Band Horwitz and coworkers discovered that taxol promotes microtubule polymerization, a unique property distinct from the microtubule depolymerizing activity of all other tubulin-binding natural products that had been identified at that time [32]. Today, the natural product, which was renamed paclitaxel by Bristol-Myers Squibb (Taxol is now a registered trademark), and its semisynthetic analog docetaxel (Taxotere) are two of the most effective drugs used for the treatment of common adult malignancies. Docetaxel is a second-generation taxane that differs from paclitaxel at the carbon 13 side chain. These two drugs are used singly and in combination with other cytotoxic or targeted agents to treat nonsmall-cell lung, breast, ovarian, prostate, and head and neck cancers. The continued clinical value of these drugs is evidenced by the ongoing development of multiple new microtubule stabilizers. Initial studies showed that paclitaxel stimulates the polymerization of purified tubulin [32]. At stoichiometric concentrations, one molecule of paclitaxel is associated with each tubulin heterodimer causing robust tubulin polymerization [33, 34]. The 1:1 binding of paclitaxel to tubulin in microtubules essentially eliminates the lag time associated with the assembly of purified tubulin into microtubules and these paclitaxel-stabilized microtubules are somewhat resistant to depolymerization by either CaCl2 or cold treatment [32]. At much lower, substoichiometric concentrations, paclitaxel binds to 1 out of every 688 tubulin heterodimers in a single microtubule, which results in a dramatic decrease in the dynamicity of microtubules without significantly affecting polymer mass [34]. The taxane-binding site was mapped to

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β-tubulin within formed microtubules at a site unique from the vinca- and colchicinebinding sites (Fig. 11.2). Paclitaxel binds reversibly and with a high affinity to the interior of the intact microtubule lumen and enhances protofilament-protofilament interactions by inducing a conformational shift that effectively straightens GDPbound tubulin into a form that more closely resembles the more stable GTP-bound structure [35, 36]. Interestingly, it is thought that paclitaxel gains access to the interior of the microtubule by diffusing through pores in the microtubule by way of low-affinity binding sites on the external surface of the microtubule [37, 38]. In cells paclitaxel is an antimitotic, causing metaphase arrest and mitotic spindle defects due to inhibition of microtubule dynamics. The IC50 for the inhibition of proliferation of most drug-sensitive cancer cell lines is between 2 and 10 nM for paclitaxel and slightly lower for docetaxel. At these concentrations, the dynamic instability of cellular microtubules is suppressed roughly 50%, which correlates with decreased tension at kinetochores in the absence of changes in microtubule mass [7, 39]. The suppression of microtubule dynamics in the mitotic spindle initiates mitotic arrest. In addition, paclitaxel causes the formation of highly abnormal spindles that consist of multiple microtubule asters [40, 41] (Fig. 11.1d). How these aberrant mitotic spindles are related to inhibition of microtubule dynamics and the signaling pathways leading from suppression of microtubule dynamics to apoptosis are not yet known. In contrast to the effects of low nanomolar concentrations of paclitaxel that suppress microtubule dynamics, much higher, micromolar concentrations of paclitaxel shift the cellular tubulin equilibrium from soluble tubulin heterodimers into microtubule polymer. This increase in cellular microtubule mass results in the appearance of stable, heavily acetylated microtubule bundles [42, 43]. Although the concentration-dependent effects of the taxanes on the dynamicity and structure of microtubules has been firmly established, it is important to consider that paclitaxel accumulates several hundred fold intracellularly as compared to the surrounding media, making exact comparisons between media and intracellular concentrations difficult [44]. The clinical development of paclitaxel was not a straightforward process; it took 21 years after the initial identification of its antitumor properties to achieve FDA approval for use in treating cancer [45]. Multiple challenges were overcome, including supply and formulation problems. The yield of paclitaxel from the bark of the Pacific Yew was abysmal, requiring the destruction of approximately one and a half 100-year-old trees to produce 1 g of the drug [46]. Although the complete synthesis of paclitaxel was eventually successful [47], the process is not amenable to economical commercial production. To circumvent the supply problem, plant tissue culture was eventually employed and the entire supply of paclitaxel is now manufactured by semisynthesis from10-desacetylbaccatin obtained from large-scale plant tissue culture [48]. The aqueous insolubility of paclitaxel required significant formulation efforts that culminated in the use of 50% ethanol and 50% polyethoxylated castor oil (Cremophor EL). Similarly, docetaxel requires formulation in polysorbate-80. These vehicles elicit hypersensitivity reactions and require pretreatment of patients with steroids and antihistamines [49]. Consistent with the effects of the vinca alkaloids, the dose-limiting toxicities of the taxanes include neutropenia, peripheral

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neuropathy, and mucositis [49]. The fact that the taxanes are one of the most widely used class of chemotherapeutic agents in spite of these limitations has prompted intense efforts to develop new drugs with the efficacy of paclitaxel that overcome its limitations of low aqueous solubility as well as innate and acquired drug resistance. Third-generation taxanes have been approved for clinical use. Abraxane, an albumin-conjugated paclitaxel, dramatically increases the aqueous solubility of paclitaxel, thus eliminating the need for administration with vehicles that can illicit hypersensitivity reactions [50]. The ability to administer the drug in a simple saline solution has the added benefit of decreasing the infusion time from 3 hours to 30 minutes [51]. Abraxane has been approved for the treatment of multidrug refractory breast cancer. Cabazitaxel (Jevtana) was approved in June 2010 for second line use in advanced hormone refractory prostate cancer in docetaxel-pretreated men and is the first clinically approved taxane that can circumvent resistance caused by the overexpression of the P-glycoprotein drug efflux pump [52]. Additional taxanes in clinical development include: tesetaxel (DJ927), BMS188797, TPI287, and milataxel (TL139, MAC-321) [3]. The advantages of these agents include oral dosing, improved bioavailability, and the ability to circumvent P-glycoprotein-mediated multidrug resistance [53, 54].

11.5

Epothilones

The 16-membered ring macrocyclic lactones epothilone A and B, isolated from the myxobacterium Sorangium cellulosum, have microtubule-stabilizing properties [55]. Although structurally diverse from the taxanes, the epothilones have similar biological effects, including the ability to promote polymerization of purified tubulin, cause cellular microtubule bundling, and form aberrant mitotic spindles, leading to mitotic arrest and apoptosis [55, 56]. The epothilones have advantages over paclitaxel and docetaxel in that they are not substrates of the P-glycoprotein efflux pump [55] and are synthetically tractable, facilitating the total synthesis of the natural products and dozens of analogs that have been used to perform detailed structure-activity relationship studies [57]. Although the epothilones competitively displace radiolabeled paclitaxel [56] and initial studies suggested a common binding pharmacophore, analysis of cell lines with mutations in the taxane-binding site suggested a common, however, not identical binding site [58, 59]. Additional studies detected differences in the ability of the epothilones and paclitaxel to promote the assembly of yeast tubulin suggesting that they did not share an identical binding mode [60]. Nettles and colleagues identified a unique epothilone pharmacophore within the expansive taxane-binding pocket [61], which explains the discrepancies noted above. However, the binding of paclitaxel and the epothilones is mutually exclusive and the two drugs do not synergistically stimulate tubulin polymerization [62] or inhibit cell proliferation [63]. The lactam analog of epothilone B, ixabepilone (Ixempra), was approved in late 2007 for the treatment of metastatic or locally advanced taxane and anthracyclineresistant breast cancer [64, 65]. Ixabepilone retains the potent antimitotic and

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microtubule-stabilizing properties of epothilone B and was engineered to increase metabolic stability [63]. However, ixabepilone still requires formulation in Cremophor EL, requiring the use of premedication and unlike epothilone B it has been shown to be a substrate for P-glycoprotein mediated transport [66]. A water-soluble epothilone B analog, BMS-310705, was developed and tested in phase-I clinical trials, however it failed to advance [67]. Other epothilones undergoing clinical development include the natural product epothilone B (Patupilone) and the epothilone B analog ZK-EPO (Sagopilone), which has the ability to cross the blood brain barrier [3]. Several epothilone D analogs were evaluated in phase-I clinical trials, however they did not progress further (Table 11.1).

11.6

Other Taxane-Site Binding Agents

Additional compounds that bind within the taxane-binding site on microtubules have been identified. Discodermolide, which was first isolated from the marine sponge Discodermia dissoluta [68], is a potent microtubule stabilizer that causes extensive microtubule bundling in interphase cells and strongly promotes the polymerization of purified tubulin [69]. Discodermolide competes with paclitaxel for binding to microtubules, indicating that they share a common binding site [70]. Mechanistic studies show that discodermolide suppresses microtubule dynamics in a manner that is subtly distinct from paclitaxel [71]. The combination of discodermolide and paclitaxel provides synergistic inhibition of microtubule dynamics [72] that functionally translates into synergistic cytotoxic effects in vitro [73]. These results suggest that discodermolide and paclitaxel share overlapping, however, not identical binding sites. Discodermolide was developed clinically and evaluated in phase-I clinical trials, however, it failed to advance presumably due to lung toxicity encountered in that trial [74]. Other microtubule-stabilizing agents that bind to the taxane site on tubulin include eleutherobin, sarcodictyins A and B, dictyostatin, and cyclostreptin [75].

11.7

Laulimalide and Peloruside A

The laulimalides are polyketide macrolides that were first isolated in 1988 from marine sponges by two independent groups [76, 77]. A decade later the mechanism of action of these compounds as novel microtubule stabilizers was discovered. Laulimalide increases the density of interphase microtubules and causes the formation of highly aberrant mitotic spindles that lead to mitotic accumulation and apoptosis [78]. In biochemical assays with purified tubulin, laulimalide stimulates tubulin polymer assembly in a manner essentially identical to paclitaxel [62, 78, 79]. Laulimalide is a potent cytotoxin with activity against a wide range of cancer cell lines and can overcome drug resistance mediated by P-glycoprotein [78–80] or mutations in β-tubulin within the taxane-binding site [79]. Laulimalide is synthetically accessible and numerous analogs have been synthesized and structure-activity relationships defined [81, 82].

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Table 11.1 Microtubule targeting drugs described in this chapter. The binding site on tubulin, class of agent, and stage of clinical development as determined by searches of active clinical trials are indicated for each drug Class

Drug

Stage of development

References

Vincristine (oncovin), vinblastine (velban), vinorelbine (navelbine), vindesine (eldisine) Vinflunine Eribulin (E7389) Maytansine DM1 antibody conjugates Dolastatin 15 Cryptophycin 52 E7974, HTI-286

FDA approved

[116]

In clinical trials FDA approved Did not progress in clinical trials In clinical trials

[2, 3] [3, 17, 117] [19] [3, 118]

Never entered clinical trials Did not progress in clinical trials Did not progress in clinical trials

[22] [27] [28]

FDA approved

[49, 50, 52]

In clinical trials

[3, 119–121]

FDA approved In clinical trials

[65] [3, 122]

Did not progress in clinical trials

[122]

Preclinical development Did not progress in clinical trials

[123] [74]

Laulimalide-site agents Laulimalide Laulimalide Peloruside A Peloruside A

Preclinical development Preclinical development

[80, 85] [87]

Other stabilizers Taccalonolide

Preclinical development

[91]

Did not progress in clinical trials In clinical trials

[99] [3]

Did not progress in clinical trials

[112]

Preclinical development

[113]

Vinca-site agents Vinca alkaloid

Vinca alkaloid Halichondrin Maytansinoid Maytansinoid Dolastatin Cryptophycin Hemiasterlin Taxane-site agents Taxane

Taxane Epothilone Epothilone Epothilone Dictyostatin Discodermolide

Paclitaxel (taxol), docetaxel (taxotere), abraxane (ABI-007), cabazitaxel (jevtana) Tesetaxel (DJ929), BMS188797, TPI287, milataxel (MAC-321) Ixabepilone (Ixempra) Epothilone B (patupilone), ZK-EPO (sagopilone) Epothilone D, BMS-310705 Dictyostatin Discodermolide

Taccalonolides

Colchicine-site agents Colchicine Colchicine Combretastatin Combretastatin A4P (zybrestat), combretastatin A1P (Oxi4503), NPI-2358, MPC-6827, ABT-751, CYT997, AVE8062 2-Methoxyestradiol 2-Methoxyestradiol (panzem) 2-Methoxyestradiol ENMD-1198

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Surprisingly, laulimalide does not inhibit the binding of radiolabeled paclitaxel to microtubules and equimolar amounts of laulimalide and paclitaxel can bind alone or in combination, consistent with laulimalide having a distinct, nonoverlapping binding site with the taxanes [79]. This represents the first new nontaxane microtubule stabilizer-binding site on tubulin. Peloruside A, another sponge-derived macrolide with microtubule-stabilizing activity (see below) binds within same site on tubulin. The nature of the laulimalide-binding site was recently uncovered. Mass spectrometry-based mass shift perturbation analyses of the laulimalide-binding site predict that it is localized on the external surface of the microtubule [83]. Refinement of the binding mode predicts that laulimalide assumes a widened conformation in solution and this explains the structure-activity results obtained with numerous analogs [83]. Functionally, laulimalide acts synergistically with taxane-site agents to stimulate tubulin polymer assembly [62, 84]. The antitumor actions of the natural and synthetic laulimalide have been evaluated and synthetic laulimalide showed minimal antitumor activity with unacceptable toxicity [80]. In contrast, a second report indicated that the naturally derived compound had antitumor effects with no associated toxicity [85]. Additional murine antitumor trials will be needed to determine the clinical potential of laulimalide and simplified, synthetically accessible analogs may provide opportunities for improvements. Peloruside A is a sponge-derived macrolide with potent cytotoxic activity that was first identified in 2000 [86]. Peloruside A causes bundling of interphase microtubules and a shift in cellular tubulin equilibrium toward the polymerized form, identical to the effects of paclitaxel [87]. Peloruside A was shown to compete for laulimalide binding, suggesting a common binding site [88]. As predicted by occupancy of a common binding site, peloruside A and laulimalide are not synergistic cytotoxins, however, peloruside A can act synergistically with other taxane-site binding microtubule stabilizers [89]. Evaluation of the clinical potential of peloruside A awaits publication of murine antitumor efficacy studies. It is interesting to speculate whether combinations of agents that bind to the taxane and laulimalide/peloruside A-binding sites will provide synergistic antitumor actions.

11.8 Taccalonolides The taccalonolides are highly acetylated hexacyclic steroids isolated from the roots and rhizomes of plants of the genus Tacca [90]. The ability of taccalonolides A and E to cause cellular microtubule stabilization was identified in a mechanism-based screen of plant extracts [91]. Although the cellular effects of taccalonolides A and E are similar to the effects of paclitaxel, initial studies suggested that they do not bind directly to tubulin/microtubules or enhance the polymerization of purified tubulin [75, 91, 92]. However, a taccalonolide with low nanomolar potency, comparable to other classes of stabilizers, was recently isolated that demonstrated the ability to directly interact with tubulin [93]. In vitro and in vivo studies have found that the taccalonolides can overcome drug resistance mediated by P-glycoprotein and that

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they retain efficacy in cell lines expressing the βIII isotype of tubulin [94]. Therefore, the taccalonolides appear to have a unique mechanism of action that is different from all other microtubule targeting agents isolated to date. Potential clinical development of the taccalonolides will require solving supply and formulation challenges similar to the issues that were faced in the development of paclitaxel [95].

11.9

Colchicine and Combretastatin

Colchicine is a microtubule-destabilizing agent that was originally extracted from the crocus plant, Colchicum autumnale. Colchicine is not used for cancer therapy because of its toxicity, however, is used to treat gout and familial Mediterranean fever [96]. Colchicine binds to free tubulin heterodimers in solution at the interface between α and β to form a tubulin-colchicine complex that is then incorporated into the growing microtubule plus end [97, 98]. Like the vinca alkaloids, colchicine inhibits microtubule dynamics and causes net microtubule depolymerization in cells. Although colchicine itself is not useful as an anticancer drug, the colchicine-binding site on tubulin is generating significant interest as a drug target [99]. One valuable property of colchicine-site agents is their relative structural simplicity. The synthetic tractability of colchicine-site agents has prompted the synthesis and study of hundreds of these agents and facilitated extensive structure-activity relationship studies and medicinal chemistry optimization [100, 101]. The combretastatins are colchicine site-binding agents that were originally isolated from the bark of the tree Combretum caffrum [102]. Combretastatins A1 and A4 have been derivatized as phosphate prodrugs that have entered clinical trials as OXi4503 (CA-1P) and Zybrestat (CA-4P) [101, 103]. In addition to CA-1P and CA4P, at least half a dozen other colchicine-site agents are currently undergoing clinical evaluations, including NPI-2358, MPC-6827, ABT-751, CYT997, and AVE8062 [3]. Various agents from this class have been in clinical development for over a decade, however, none have achieved regulatory approval for use in cancer.

11.10 Vascular Disrupting and Antiangiogenic Activities of Microtubule Targeting Agents The most interesting property of the combretastatins and the combretastatin-like small molecules undergoing clinical development is their ability to cause rapid disruption of tumor vasculature, which classifies them as vascular disrupting agents (VDA). VDA activity is characterized by a rapid loss of tumor blood vessel integrity resulting in a drop in tumor blood flow that is initiated within minutes after the administration of the drug at submaximally tolerated doses [104]. This loss of perfusion leads to central necrosis of the tumor within hours of drug administration. However, a viable rim of tumor cells remains at the tumor periphery suggesting that VDAs will

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be most effective when used in combination with cytotoxic or antiangiogenic agents that target these remaining tumor cells [105]. The ability of various microtubule targeting agents to disrupt tumor vasculature has been recognized since the 1930s when hemorrhaging was reported within hours of colchicine administration to mice [106]. However, the vascular perturbations that occur with classical microtubule depolymerizing agents such as colchicine, vincristine, and vinblastine require treatment at their maximum tolerated dose, eliminating their therapeutic window for VDA activity. In contrast, the VDA activity of the combretastatins and other colchicine-site agents in clinical development occurs at much lower, well-tolerated doses. The nature of the discrepancies in the doses required to observe VDA activity between these various agents is not known. VDA activity is hypothesized to involve cytoskeletal-mediated shape changes of tumor endothelial cells leading to loss of endothelial layer integrity, and an increase in vascular permeability. Rapid alterations in endothelial cell morphology can be readily observed with the addition of a VDA to endothelial cells in culture [107]. The increased sensitivity of tumor endothelial cells to these effects may be due to the fact that the tumor vasculature is not as well developed or as mature as normal vasculature [108]. Some microtubule targeting agents have also demonstrated antiangiogenic activity. In contrast to VDA activity, which disrupts existing tumor vasculature, the inhibition of angiogenesis prevents the formation of new blood vessels [104]. Microtubule targeting agents appear to inhibit angiogenesis by targeting rapidly dividing tumor endothelial cells, decreasing endothelial cell motility, and reducing the expression of HIF-1α, a proangiogenic transcription factor [104]. Frequent low dose or “metronomic” dosing of these drugs has been proposed for optimal antiangiogenic activity [109]. Microtubule stabilizers and destabilizers, including paclitaxel, docetaxel, and vinblastine, inhibit proliferation of tumor vasculature at concentrations near their maximum tolerated doses [110, 111]. 2-Methoxyestradiol (2ME2), an endogenous metabolite of 17β-estradiol that binds to the colchicine site, showed effective antiangiogenic activity in preclinical studies at submaximum tolerated doses, however it failed during clinical development primarily due to rapid metabolism [112]. Intriguingly, it was recently reported that ENMD-1198, an orally active 2 ME2 analog with improved bioavailability, demonstrates both antiangiogenic and VDA properties in vitro [113]. It will be of interest to see whether these properties translate to increased clinical efficacy as ENMD-1198 enters clinical development.

11.11 Conclusions Microtubule targeting drugs remain important in the palliative and curative treatment of cancer. New microtubule-active agents with the ability to circumvent major drug resistance mechanisms are continually advancing through clinical trials and are being approved for use against cancer. Currently approved agents are also finding new indications including use in combination therapy to increase the efficacy of targeted therapies and novel formulations may provide additional new indications for microtubule targeting drugs. Further optimization of known microtubule targeted agents

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may increase their antitumor efficacy or expand their therapeutic window, thereby decreasing dose-limiting toxicities. In addition, there is reasonable expectation that microtubule targeting agents with novel molecular scaffolds will be identified in the future. There is much work yet to be done to truly understand the mechanism of antitumor action of microtubule targeting agents. The nature of the biological differences of structurally similar compounds for tumor indications and limiting toxicities remains a mystery. The mechanisms of the multifactoral actions of new generation microtubule targeting agents, including their antiangiogenic and vascular disrupting activities also requires further study. A more complete understanding of the mechanism of these drugs will likely include further testing of the intriguing hypothesis that the antitumor actions of microtubule targeting agents may be due at least in part to their effects on interphase microtubules [114, 115]. Finally, exploration into the signaling pathways that lead from disruption of microtubule dynamics to initiation of mitotic arrest and apoptosis are warranted to fully understand the antimitotic mechanisms of microtubule targeting agents. Identification of these pathways and knowledge of the genetic defects in certain tumors that impact mitotic checkpoints may lead to a more rational use of specific microtubule targeting drugs. These studies have the potential to identify new therapeutic targets and drugs directed against these novel targets could achieve the efficacy of tubulin-binding agents without the major dose-limiting tubulin-related side effects. Acknowledgments We apologize to the many investigators who have made important contributions to the identification, mechanisms of action, and chemistry of microtubule targeted agents and whose work was not cited due to space considerations. Grant support from the National Cancer Institute of the National Institutes of Health (United States), CA121138 (SLM), DOD-CDMRP Postdoctoral Award BC087466 (ALR), and the Cancer Therapy & Research Center are gratefully acknowledged.

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Chapter 12

Microtubules, Drug Resistance, and Tumorigenesis Joshua A. McCarroll and Maria Kavallaris

Abstract Microtubules are highly dynamic structures that comprise α- and β-tubulin heterodimers which are essential in mitosis. These features make microtubules an important target for many natural and synthetic anticancer drugs. Mutations in β-tubulin that affect microtubule polymer levels or drug binding are associated with resistance to tubulin-binding agents such as paclitaxel. Moreover, aberrant expression of specific β-tubulin isotypes, namely βIII-tubulin as well as microtubule-binding proteins are now recognized as clinically important determinants in tumor aggressiveness and resistance to chemotherapy. More recently, it has been suggested that β-tubulins may also be linked to the tumorigenic phenotype of certain cancers such as non-small-cell lung cancer. Understanding the mechanisms whereby β-tubulins exert their effect on drug resistance and tumorigenesis are critical to the identification of novel drug targets and improvements in current therapies to increase the long-term survival of cancer patients.

12.1

Introduction

Microtubules are cytoskeletal structures which play an important role in key cellular events such as cell division. During mitosis, microtubules form the mitotic spindle that transports daughter chromosomes to separate poles of the dividing cell [1]. The critical role of microtubules in this cellular process makes them an attractive drug target for anticancer therapies. Chemotherapy agents including the microtubulestabilizing (e.g., Taxanes) and destabilizing drugs (e.g., vinca alkaloids) bind to the β-tubulin subunit of microtubules to exert their toxic effect by inducing a potent mitotic block [2]. These agents are an important component in the treatment of many adult and childhood cancers. However, the potential toxic side effects such as neutropenia and neurotoxicity and the development of resistance can limit the clinical usefulness of these antimitotic drugs. M. Kavallaris () · J. A. McCarroll Children’s Cancer Institute Australia, Lowy Cancer Research Centre, University of New South Wales, 2052 Sydney, NSW, Australia e-mail: [email protected] Australian Centre for Nanomedicine, Faculty of Engineering, University of New South Wales, Sydney, NSW, Australia

M. Kavallaris (ed.), Cytoskeleton and Human Disease, DOI 10.1007/978-1-61779-788-0_12, © Springer Science+Business Media, LLC 2012

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Acquired and/or intrinsic resistance to tubulin-binding agents (TBAs; also referred to as antimicrotubule agents or microtubule targeting drugs) has been the focus of intense research for the past several decades. Resistance to this class of drugs has largely been attributed to the development of multidrug resistance [2]. This process involves the increased expression of ATP-binding cassette (ABC) proteins such as p-glycoprotein (P-gp) and multidrug-resistance associated protein (MRP) [3]. These proteins actively extrude drugs out of cells demonstrating a preference for many of the lipophilic, hydrophobic antimitotic agents including the taxanes and vinca alkaloids [3]. However, it is now evident that other cellular processes play a role in the development of chemoresistance. For example, significant advances have been made to the understanding of the role of microtubule proteins in regulating drug resistance. Numerous studies have demonstrated that mutations in the tubulin protein, namely βI-tubulin, can significantly impact on the ability of TBAs to bind to their target [2, 4]. In addition, aberrant expression of microtubule-associated proteins including stathmin, MAP2, Tau and MAP4 have also been shown to be involved in drug resistance [2]. More recently, an increasing number of studies have reported that altered expression of β-tubulin isotypes play an important role in modulating sensitivity to TBAs [2]. Moreover, there is now evidence to suggest that specific βtubulin isotypes may also be involved in modulating tumor growth and development as well as acting as survival factors in response to cytotoxic stress [2]. This chapter will focus on the tubulin/microtubule network and its importance as a drug target for anticancer therapies. In addition, functional as well as clinical studies which examine the differential expression of β-tubulin isotypes and their role in regulating drug resistance in epithelial and non-epithelial cancers will be discussed. Finally, recent evidence which demonstrates a broader role for β-tubulins in tumorigenesis will also be described.

12.2 Tubulin and Microtubules Microtubules comprise heterodimers of α and β-tubulin. Both the α- and β-tubulin proteins are 50 kDa in size and are approximately 50% identical to one another [2]. The tubulin heterodimers assemble head-to-tail with the α-subunit of one dimer in contact with the β-subunit of the next dimer. The resulting linear protofilaments comprise the backbone that gives rise to their characteristic hollow tube-shaped structure (refer to Chap. 2). The microtubule consists of a parallel arrangement of 13 protofilaments in an imperfect helix [2, 5]. The head-to-tail order gives polarity to the microtubule with a negative (α-tubulin) and positive (β-tubulin) end. The negative end of the microtubule is attached to the microtubule-organizing center (MTOC) located at the centrosome in the cytoplasm, while the positive end is distal [2, 5]. An important property of microtubules is their highly dynamic nature which is characterized by rapid phases of lengthening and shortening. This process is critical for many biological processes including proper attachment and segregation of chromosomes during mitosis to local stabilization of microtubules toward the front of migrating cells. Microtubule instability involves the cooperative assembly of

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α- and β-tubulin heterodimers followed by GTP/GDP exchange. The α-tubulin subunit binds to GTP in an irreversible manner, while the β-tubulin subunit can be bound to GTP or GDP, favorable for microtubule polymerization or depolymerization, respectively [2, 6]. This property allows for the lengthening or shortening of α/β heterodimers at both ends of the microtubule. Two types of microtubule dynamic behaviors have been well characterized, treadmilling and dynamic instability [2, 5, 6]. Treadmilling involves the net addition of tubulin dimers at the plus (β-tubulin) end coupled with the net dissociation at the minus (α-tubulin) end. This produces a flow of subunits from one end of the microtubule to the other without significantly altering the length of the microtubules [2, 5, 6]. Dynamic instability involves microtubules alternating between growing and shortening. A transition from a growing to a shortening phase is referred to a catastrophe, while a transition from a shortening to a growing phase is known as rescue [2, 5, 6]. Collectively, these processes are essential in mitosis, particularly for the proper function of the mitotic spindle.

12.3 Targets for Anticancer Therapy Given the importance of the microtubule network in cell division, it is not surprising that an intense research effort has been made into the development of chemical agents which target microtubules for the treatment of cancer. TBAs are mitotic inhibitors which are classified as microtubule-stabilizing (e.g., taxanes, epothilones) or microtubule-destabilizing (e.g., vincristine, vinblastine, and vinorelbine; [2, 5, 7]). Microtubule-stabilizing compounds exert their effect by promoting polymerization and increasing the microtubule polymer mass within the cell. In contrast, microtubule-destabilizing compounds inhibit polymerization and decrease polymer mass [2, 5, 7]. However, it is to be noted that at low concentrations both stabilizing and destabilizing drugs suppress microtubule dynamics without altering polymer mass [2, 5, 7]. The synthesis of the crystal structure for the α- and β-tubulin heterodimer has allowed researchers to identify the binding site of clinically important TBAs [8, 9]. This class of drugs exert their biological effect by binding to the β-tubulin subunit in the α- β-tubulin heterodimer [8, 9]. Once bound to this site, TBAs potently suppress microtubule dynamics by blocking the transition from metaphase to anaphase during mitosis [2, 5, 7]. This in turn, leads to the disruption of the mitotic spindle and an extended mitotic arrest and ultimately cell death [2, 5, 7]. However, the mechanism(s) as to how mitotic arrest leads to cell death still remains to be fully determined.

12.4

Resistance to TBAs

Intrinsic or acquired resistance to TBAs is a common occurrence in many tumors and severely limits their clinical effectiveness. To date, the mechanisms mediating resistance to TBAs are not well known. However, evidence is mounting to suggest that

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Fig. 12.1 Tubulin-binding agent resistance. Drug resistance is multifactorial and this schematic diagram depicts mechanisms of resistance identified in cell lines and human tumors. Resistance mechanisms are associated with reduced intracellular accumulation of drug due to increased drug efflux mediated by multidrug transporters; mutations in tubulin that affect drug binding and/or microtubule polymer levels; tubulin/microtubule alterations such as increased expression of βIIItubulin; alterations in the actin cytoskeleton that affect the action of TBAs; and defects in apoptotic cell death. Lines in red depict mechanisms that have been identified in clinical resistant tumor samples

resistance to TBAs can occur at multiple stages including cellular efflux of the drug, mutations of the β-tubulin gene, deficient induction of apoptotic signaling cascades, as well as alterations in the expression of tubulin isotypes ([2, 4, 5]; Fig. 12.1). The presence of membrane efflux pumps which belong to the ATP-binding cassette (ABC) family can confer resistance to TBAs in many different types of cancer cells [3, 10]. For example, P-gp, which is the product of the MDR1 gene and is responsible for the classical multidrug resistance phenotype, is known to actively extrude both vinca alkaloids and taxanes from the cell [3]. This in turn, leads to lower intracellular drug concentrations and poor cytotoxic activity [3]. In addition, both multidrug resistance-associated protein 1 (MRP1), MRP2, and MRP7 have also been shown to be substrates for both vinca alkaloids and taxanes [10–12]. However, despite strong in vitro data demonstrating their ability to significantly limit the intracellular accumulation of TBAs into cancer cells, their clinical relevance in cancer patients has been controversial. Several clinical trials using inhibitors to multidrug transporters and their effect on reversing TBA resistance have been disappointing [3, 13–15]. Moreover, studies which have assessed the clinical association of multidrug

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resistance proteins and TBA resistance have also been conflicting [3, 13–15]. Collectively, these studies support the idea that drug resistance is multifactorial and other mechanisms are also responsible for promoting resistance to TBAs. Acquired mutations of the β-tubulin gene namely, βI-tubulin have been reported in a number of different cancer cell lines selected for resistance to TBAs in the laboratory (reviewed in [16]). These mutations alter the endogenous stability of microtubules and thereby compensate for the stabilizing or destabilizing activities of TBAs such as paclitaxel and vincristine [17, 18]. To confirm whether such mutations exist in a clinical setting, several studies have screened patients for the presence of β-tubulin mutations before or after exposure to chemotherapy. However, to date, there has been no strong indication that these mutations are present in patients and if they are present, they are likely to be rare occurrences [16]. Alterations of the apoptotic signaling cascades which act downstream of the microtubule-associated effects of TBAs in cancer cells are also thought to be important in the resistance process. It is well documented that microtubules interact with various cellular organelles and act as scaffolds for regulatory proteins such as Bcl-2, inhibitor of apoptosis (IAP) proteins and p53 all of which are involved in regulating apoptosis [2]. For example, the Bcl-2 protein family protects cells from undergoing apoptosis by preventing mitochondrial dysfunction and the release of proapoptotic molecules from the mitochondria [19]. Interestingly, several studies have demonstrated that increased expression of Bcl-2 in leukemia cells potently suppresses the apoptotic response induced by TBAs without affecting their actions on microtubule dynamics or cell cycle arrest [20–22]. The IAP family of proteins is known to play a major role in suppressing apoptotic cell death as well as regulating cell division [23]. Studies have shown that one member of the IAP family survivin is involved in both the preservation of cell viability and regulation of mitosis in cancer cells [24, 25]. Indeed, a correlation between increased levels of survivin and a poor clinical outcome has been reported in a number of different cancers [26–28]. Survivin is a dual-function protein; it suppresses apoptosis in cells by inhibiting caspase activity as well as promoting cell growth by stabilizing microtubules during mitosis [19]. Several studies have demonstrated that increased levels of survivin help to counteract the therapeutic effect of microtubule-destabilizing agents in cancer cells by stabilizing tubulin polymers [29, 30]. Moreover, there are reports which show that inhibition of survivin restores sensitivity to a range of chemotherapy agents including the TBAs vinca alkaloids [25]. Another important signaling protein which is associated with microtubules is the tumor suppressor p53. This signaling protein is known to be involved in regulating many cellular functions including cell cycle arrest and apoptosis [31]. Several studies have demonstrated that wild-type p53 binds to microtubules in a variety of human cancer cell lines of distinct origin and that it requires an intact microtubule network for its intracellular trafficking into the nucleus [31]. Indeed, it has been shown that disruption of microtubules by TBAs results in impaired p53 accumulation. A potential link between p53 and sensitivity to TBAs was originally reported by Wahl et al. [32]. The authors demonstrated that deletion of p53 in normal human fibroblasts resulted in a significant increase in sensitivity to paclitaxel [32]. However, further

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Table 12.1 β-Tubulin isotype nomenclature, chromosome localization, and tissue distribution Tubulin isotype

HGNC gene symbol

Accession number

Chromosome localization

Tissue distribution

βI

TUBB

NM_178014

6q21.33

βIIA βIIB

TUBB2A TUBB2B

NM_001069 NM_178012

6p25.2 6p25.2

βIII

TUBB3

NM_006086

16q24.3

βIVa βIVb βV

TUBB4 TUBB2C TUBB6

NM_006087 NM_006088 NM_032525

19p13.3 9q34.3 18p11.21

βVI

TUBB1

NM_030773

20q13.32

Ubiquitous; predominant isotype in many cell lines Major isotype of neurons; low levels in lung, kidney, spleen, stomach, and thymus; increased levels in prostate adenocarcinoma Neurons; sertoli cells of testis; increased levels in a number of tumor types compared to normal tissue of tumor origin Brain-specific Ubiquitous with highest levels in testis Ubiquitous, described in adenocarcinoma, can also be decreased in tumors Hematopoietic-specific cell types; megakaryocytes and platelets

studies in a number of different human cancer cell lines failed to show any significant correlation between p53 status and TBA sensitivity [33, 34]. These results have also been observed in the clinic where p53 status has failed to act as a predictive factor for response to TBAs [35, 36].

12.5 Tubulin Alterations in Cancer Alterations in the expression of microtubule proteins in cancer cells are now considered to be a major factor in chemotherapy drug resistance. An increasing number of reports have described differences in the expression pattern of β-tubulin isotypes in drug-refractory and aggressive tumors (reviewed in [2]). Altogether, there are seven different β-tubulin isotypes, encoded by multiple genes that display tissue-specific expression (reviewed in [2]; Table 12.1). For example, βI-tubulin (encoded by the TUBB gene) is constitutively expressed in many tissues; βIII-tubulin (encoded by the TUBB3 gene) is expressed in neurons and Sertoli cells; and βVI (encoded by the TUBB1 gene) restricted to hematopoietic tissues. The sequences are highly conserved across β-tubulin isotypes, however, they differ in the carboxy-terminal region (last 20–25 amino acids) that are thought to impart functional differences to microtubules [37]. βIII-tubulin is abundant in neuronal tissues where it is differentially expressed during fetal and postnatal development [37]. It has also been reported in fetal respiratory epithelium, specifically in Kulchistky neuroendocrine cells. Most recently, the βIII-tubulinisotype has received increased attention due to a large number of clinical and functional studies showing its role in regulating chemotherapy sensitivity in cancer cells.

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Lung Cancer

Lung cancer is the leading cause of cancer death in adults [38]. The World Health Organization has classified lung cancer into two classes; Small-Cell Lung Cancer (SCLC) and Non-Small-Cell Lung Cancer (NSCLC; [39]). NSCLC accounts for more than 85% of all cases with a majority of patients presenting with advanced disease and local and distant metastases. Hence, the 5-year survival rates for this disease have remained at approximately 15% for the past three decades. These dismal statistics have led investigators to search for potential prognostic (factors which have an impact on disease outcome regardless of the treatment strategy), and predictive (factors which predict the activity of a specific agent) markers for this disease. One candidate protein which holds promise as a prognostic and/or predictive marker for NSCLC is the microtubule protein βIII-tubulin. Mounting clinical evidence suggests that patients with high levels of βIII-tubulin have a much poorer survival outcome and are most likely to be resistant to chemotherapy agents. In 2003, Rosell et al. [40] used RT-PCR to measure βIII-tubulin mRNA levels in 75 NSCLC tumor samples. They demonstrated that NSCLC patients with low levels of βIII-tubulin had a better response when treated with carboplatin/paclitaxel and a longer time to disease progression when treated with vinorelbine and gemcitabine compared to patients with high levels of βIII-tubulin mRNA [40]. A study by Seve et al. [41] assessed the prognostic and predictive value of βIII-tubulin expression using immunohistochemistry in 91 NSCLC patients with locally advanced or metastatic NSCLC treated with paclitaxel-based or other regimens that did not include TBAs [41]. Among patients receiving paclitaxel low levels of βIII-tubulin correlated with a better response rate, longer progression-free survival and overall survival. However, this was not found to be the case in patients receiving treatment regimens without TBAs. Moreover, a multivariate analysis taking into account patients sex, age, tumor histology, and stage of disease confirmed that low expression of βIII-tubulin in tumor cells was independently associated with progression-free survival and overall survival [41]. Recently, the results of these earlier studies have been confirmed in larger NSCLC patient cohorts [42–44]. Collectively, these studies highlight the potential of βIIItubulin as a predictive and/or prognostic indicator which could form part of a panel of biomarkers with strong prognostic and predictive power for NSCLC. Functional evidence to support the clinical observation of increased levels of βIII-tubulin with chemoresistance and aggressive tumor growth in NSCLC has been lacking. A study by Kavallaris et al. [45] provided the first laboratory evidence that βIII-tubulin is associated with chemotherapy drug resistance. In this study, the authors demonstrated that A549 NSCLC cells selected for resistance to paclitaxel had significantly increased expression of βIII-tubulin when compared to the parental drug-sensitive cell line. Importantly, when βIII-tubulin was suppressed using antisense oligonucleotides the sensitivity of these cells to paclitaxel was partially restored [46]. The partial restoration to paclitaxel could be explained by the multifactorial nature of drug resistance in these cells, which include tubulin mutations and increased expression of drug transporters. Recently, a direct functional role for

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βIII-tubulin in modulating sensitivity to chemotherapy in NSCLC was provided by Gan et al. [47]. In this study, the authors used highly specific and potent siRNA directed against βIII-tubulin in two independent NSCLC cell lines (H460 and Calu-6) which aberrantly express βIII-tubulin. Suppression of βIII-tubulin sensitized these cells to a broad range of TBAs including taxanes and vinca alkaloids. Interestingly, in H460 NSCLC cells, suppression of βIII-tubulin did not affect microtubule dynamics, however, increased the level of apoptosis in cells treated with low concentrations of vincristine or paclitaxel [48]. The increase in apoptosis appeared to be independent of mitotic arrest, and these low drug concentrations did not affect microtubule dynamics [48]. However, at higher concentrations of vincristine or paclitaxel, both mitotic arrest and apoptotic cell death were observed, suggesting that suppression of βIII-tubulin expression increased the effectiveness of TBAs through two mechanisms: suppression of microtubule dynamics and a mitosis-independent mechanism of cell death [48]. Another important finding was that knockdown of βIII-tubulin expression using RNA interference increased NSCLC cell sensitivity to several DNA-damaging agents that are structurally and functionally unrelated to TBAs [47, 49]. This result suggested that βIII-tubulin has a broad role in chemotherapy drug resistance. Although this result was unanticipated, it is not surprising given that DNA-damaging agents such as cisplatin and carboplatin are often used in combination therapy in NSCLC and treatment failure also means that this class of drugs are not effective against tumor cells. The increased sensitivity of the βIII-tubulin-knockdown cells to TBAs and DNA-damaging agents was associated with an increased propensity of cells to undergo drug-induced apoptosis [47, 49]. Taken together, these findings raise the possibility that βIII-tubulin could be a cellular survival factor that when overexpressed confers resistance to chemotherapy.

12.5.2

Ovarian Cancer

Ovarian cancer is the fifth most common cause of cancer death in women, and accounts for more deaths than any other cancer of the female reproductive system [50]. Despite initial response to chemotherapy most patients will become chemoresistant. First-line treatment often includes the use of a DNA-damaging agent in combination with a TBA such as paclitaxel [51]. One of the earliest studies to demonstrate altered β-tubulin iostype expression in drug-resistant ovarian tumors was reported by Kavallaris et al. [45]. In this study, tumors resistant to paclitaxel were shown to express significantly higher levels of βIII- and βIVa-tubulin when compared to untreated primary tumors. Importantly, this study has been supported by more recent data to show that indeed increased expression of βIII-tubulin is a marker of chemoresistance and poor prognosis in ovarian cancer. For example, Mozzetti et al. [51] used a cohort of 41 ovarian tumor samples (28 patients chemotherapy-sensitive and 13 chemotherapyresistant) and measured the levels of MDR1 as well as examined the presence of point mutations in the α- and β-tubulin gene. They also examined the expression levels of

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several β-tubulin isotypes. No significant difference in MDR1 expression was observed between the drug-sensitive and resistant tumors. The presence of mutations in the α- and β-tubulin gene were also absent between the two groups. However, the authors reported a significant increase in βIII-tubulin at both the gene and protein level in paclitaxel-resistant tumors when compared to drug-sensitive tumors or normal tissue [51]. More recently, several reports have also confirmed that βIII-tubulin is a prognostic indicator of shorter progression-free survival and chemoresistance in advanced ovarian cancer [52–54]. In contrast to most other epithelial cancers, there has been a large body of laboratory evidence to confirm that βIII-tubulin is involved in regulating drug resistance in ovarian cancer cells. For example, studies by Cicchillitti et al. [55] have used a proteomic approach to identify differentially expressed proteins in paclitaxel-resistant and sensitive ovarian cancer cell lines. Significant changes in protein expression between the two cell lines were detected with the greatest differences being observed for proteins involved in stress response, cell cycle/apoptosis, and the cytoskeleton [55]. Interestingly, the authors demonstrated that βIII-tubulin in the drug-resistant cell lines was able to form protein-protein interactions (via disulfide bridges) with a number of proteins known to play a role in the adaptation to cellular stress (oxidant stress and glucose deprivation; [55]). These results suggest that βIII-tubulin may be part of a survival pathway that enables cells to adapt to a stressful microenvironment and appears to be enhanced in drug-resistant cancer cells. This is further supported by evidence which demonstrates that βIII-tubulin is activated by hypoxia in these cells and is under the transcriptional control of HIF-1α [56]. Taken together, it is possible that increased levels of βIII-tubulin allows drug-resistant cancer cells to survive in a hostile tumor microenvironment of the most advanced and aggressive ovarian cancers. However, whether this scenario is recapitulated in other cancers remains to be determined.

12.5.3

Breast Cancer

An important advancement in the treatment of breast cancer has been the introduction of TBAs (docetaxel) as a single agent or in combination with other chemotherapy drugs [57]. Although up to 50% of patients will have a clinical response upon treatment with this class of drug, most will eventually relapse. Studies performed using breast cancer cell lines which are estrogen receptor positive or negative and made resistant to docetaxel were found to contain significant changes in the expression of several β-tubulin isotypes including class I, II, III, IVa, and IVb when compared to the parental drug-sensitive cells lines [58]. These results were later confirmed by Tomamasi et al. [59] who also demonstrated increased levels of βIII-tubulin in breast cancer cells which displayed less sensitivity to paclitaxel. Moreover, when these breast cancer cells were treated with antisense oligo nucleotides against βIII-tubulin, sensitivity to paclitaxel was partially restored. These results were also confirmed at the clinical level where a retrospective immunohistochemical study assessed β-tubulin levels in 92 patients receiving paclitaxel therapy [59]. Importantly, 35% of patients with high

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expression of βIII-tubulin showed disease progression when compared to only 7% of patients with low levels of βIII-tubulin. Collectively, these results suggest that βIII-tubulin could be considered as a predictive marker for TBA resistance in breast cancer patients. Further functional evidence supporting a role for βIII-tubulin in chemoresistance was most recently provided by Stengel et al. [60] In this study, the authors constructed an in vitro model which allowed for subtle changes in the expression of βIII-tubulin in several different breast cancer cell lines. Breast cancer cells transfected with a vector construct containing wild-type βIII-tubulin cDNA had a two- to fourfold increase in βIII-tubulin when compared to cells transfected with the empty vector [60]. This increase in βIII-tubulin did not cause any toxicity to the cells. In contrast, cells treated with siRNA targeting βIII-tubulin showed a 50% decrease in βIII-tubulin levels when compared to controls. Importantly, cells with increased expression of βIII-tubulin showed resistance to the TBAs paclitaxel and vinorelbine. In contrast, cells treated with siRNA against βIII-tubulin displayed increased sensitivity to both drugs [60]. Interestingly, altered levels of βIII-tubulin had no effect on drugs which bind to the colchicine-binding site. It is to be noted that these compounds are structurally different to paclitaxel and vinorelbine. The authors suggest that this structural difference may allow for the formation of a stable complex with βIII-tubulin and that βIII-tubulin resistance in breast cancer may be drug binding-site related [60]. However, this has yet to be fully explored in other cancers.

12.5.4

Prostate Cancer

Prostate cancer is the most common malignancy and the second most common cause of cancer death in men [61]. Resistance to taxane-based therapy has been reported and clinical studies have shown altered expression of β-tubulins in these tumors [62, 63]. An early report by Ranganathan et al. [63] showed an increase in the expression of βIII-tubulin and βIVa-tubulin in prostate cancer cells that were made resistant to the TBA Estramustine [63]. Recently, a study by Terry et al. [64] demonstrated that castration-resistant prostate tumors which are highly refractory to hormone therapy (depletes circulating androgen levels) and most alternate chemotherapies, had significantly increased levels of βIII-tubulin when compared to hormone-na¨ıve or hormone-therapy-treated tumors [64]. Notably, βIII-tubulin expression was also detected in cancer cells of metastatic lesions in the liver of patients with castration-resistant prostate malignancy. These results were confirmed in the laboratory when androgen-sensitive prostate cancer cells displayed a significant increase in βIII-tubulin protein expression when switched to an androgen-depleted growth medium. This increase persisted for up to 3 months under these growth conditions. Moreover, when mice were inoculated with prostate cancer cells and tumors collected after castration (androgen depletion), increased levels of βIII-tubulin were detected in the primary tumors when compared to control (noncastrated) mice. More recently, studies also showed that the protein levels of βIII-tubulin increased in prostate cancer cells when exposed to acute or chronic amounts of docetaxel [62].

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This result was also confirmed in the clinic where βIII-tubulin was shown to be a predicative indicator for overall survival in patients receiving docetaxel [62]. Together, these findings highlight the importance of βIII-tubulin and its potential as not only a predictive marker for disease aggressiveness but also as a means for selecting patients for docetaxel-based treatment regimens.

12.5.5

Pancreatic Cancer

Pancreatic cancer is a devastating disease with a dismal prognosis. It is the fourth leading cause of cancer-related deaths in Western society, with a 5-year survival rate of less than 5%. A major reason for the poor clinical outcome is its well-known resistance to a broad range of chemotherapeutic agents which include the TBAs. To understand the mechanisms of resistance to TBAs in pancreatic cancer, Lui et al. [65] developed pancreatic cancer cell lines which were resistant to taxotere (member of the taxane family). As expected these cell lines had significantly higher levels of the drug efflux pump MRP1 when compared to their drug-sensitive parental cell lines. In addition, there was also a greater than twofold increase in the mRNA expression of βII- and βIII-tubulin [65]. Clinical studies assessing the expression levels of β-tubulins in pancreatic tumors have been lacking. Recently, a study by Lee et al. [66] showed for the first time that βIII-tubulin is absent in nonneoplastic ducts of the pancreas, however, it is highly expressed in the majority of invasive pancreatic ductal adenocarcinoma samples (47/60 tumor samples) and in several pancreatic cancer cell lines [66]. Moreover, βIII-tubulin expression was increased in the progressive stages (1–3) of pancreatic intraepithelial neoplasia (PANINs, precursor lesions of ductal carcinoma). Interestingly, the authors also reported that pancreatic cancer cells which harbored the oncogenic kRAS gene (a promoter of tumorigenesis) had significantly more βIII-tubulin, suggesting that its expression may be associated with activation of kRAS and promotion of tumorigenesis. However, to date, the functional role of βIII-tubulin in chemosensitivity and tumor growth in pancreatic cancer is unknown. Taken together, it is clear from the evidence outlined above that βIII-tubulin is an important indicator of sensitivity to TBAs as well as overall survival and disease progression. Therefore, future studies designed to include βIII-tubulin in a panel of diagnostic markers as well as the design of therapies which target this protein may help refine personalized medicine as well as significantly improve the long-term survival of patients diagnosed with epithelial-derived malignancies.

12.6 β III-Tubulin and Nonepithelial Cancer In contrast to somatic organs and tissues, βIII-tubulin is highly expressed in both the central and peripheral nervous systems. This is most evident during fetal and postnatal development. In adult tissues the distribution of βIII-tubulin is almost exclusively

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neuron-specific. Importantly, an increasing number of studies have confirmed that alterations in the expression of β-tubulins are involved in chemoresistance and tumor growth in nonepithelial cancers.

12.6.1

Brain Cancer

Glioblastoma is the most common form of brain cancer and the second most common cause of death from neurological disease [67]. Five-year survival rates for this malignancy are less than 15% [67]. A distinctive feature of these tumors is their highly invasive and chemoresistant nature. Hence, there is an urgent need to identify novel drug targets for this disease. Importantly, several clinical studies have identified significant alterations in the microtubule cytoskeleton including increased expression of βIII-tubulin, γ-tubulin and other microtubule-associated proteins in glioblastoma [68–70]. In the central nervous system (CNS), βIII-tubulin displays a different pattern of expression in embryonal/neuronal neoplasms as compared to neoplasms of glial origin (reviewed in [71]). For example, in embryonal tumors such as medulloblastomas, cerebral neuroblastomas and supratentorial primitive neuroectodermal tumors, βIII-tubulin is constitutively expressed in tumor cells undergoing neuritogenesis and ganglionic maturation concomitant with decreased cell proliferation. In contrast, βIII-tubulin expression in glial tumors is considered to be ectopic and aberrant, with its levels increasing according to an ascending scale of histological malignancy (reviewed in [71]). Indeed, immunohistochemical studies have identified βIII-tubulin as one of six marker proteins to show significant differences in expression between low-grade and high-grade gliomas [72]. Interestingly, βIII-tubulin has been shown to colocalize with γ-tubulin suggesting a possible interaction between the two proteins [70]. Moreover, increased expression of βIII-tubulin has been reported in glioblastoma tumor cells that are adjacent to areas of ischemic necrosis [69]. These cells are thought to be inherently resistant to irradiation and chemotherapy. However, whether βIII-tubulin plays a role in regulating these processes or in the tumor growth and aggressiveness of glioblastoma has yet to be determined.

12.6.2

Melanoma

Malignant melanoma comprises only 10% of all skin cancers, however, it accounts for 80% of all skin cancer deaths [73]. Taxane-based regimes are considered as secondline chemotherapy for metastatic melanoma. However, clinical data are mixed for taxane therapy with one study reporting no difference in median survival time or median time to disease progression for patients treated with paclitaxel alone or in combination with the DNA-damaging agent carboplatin [74]. Therefore, the efficacy of TBAs remains controversial for malignant melanomas and at best is considered beneficial for only a small subgroup of patients. Recently, there have been several in vitro studies which have reported increased levels of βIII-tubulin in docetaxelresistant melanoma cell lines [75]. Moreover, docetaxel was shown to activate several

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important signaling proteins in these cells [76]. One of the proteins known as PKCε has been reported to be antiapoptotic. Interestingly, this protein was found to be physically associated with εβIII-tubulin in the resistant cell lines [76]. In the absence of drug the distribution of βIII-tubulin with PKCε was mainly punctate. However, after exposure to docetaxel a small proportion of the PKCε shifted to the peripheral region of the cell. This suggests that βIII-tubulin may be part of an antiapoptotic pathway which aids in the translocation of signaling proteins from the cell cytoplasm to the periphery in order for them to become activated. To confirm whether βIII-tubulin was present in tumor tissue, Akasaka et al. [77] immunohistochemically examined 49 malignant melanoma tumor tissue sections and 10 nonneoplastic skin samples. βIII-tubulin was found to be present in normal epidermal melanocytes and nerve cells in the nonneoplastic lesions [77]. Expression of βIII-tubulin in the malignant tumors was varied and intratumoral heterogeneity was commonly observed. βIII-tubulin was present in approximately 80% of the malignant tumors. However, there was a trend for high-stage tumors to exhibit a loss of βIII-tubulin expression. Notably, βIII-tubulin protein expression disappeared in one case of malignant melanoma in situ. This suggests that loss of βIII-tubulin may occur in a subset of patients with malignant melanoma and that this loss is more frequent as the tumor progresses [77]. Kaplan-Meier curves also showed a trend (did not reach significance, p = 0.06) toward a worse prognosis for patients with βIII-tubulin-negative tumors [77]. Functional studies have also confirmed that the expression levels of βIII-tubulin in melanoma cancer cells correlate with paclitaxel resistance [77]. Treatment of cells with siRNA against βIII-tubulin was able to restore sensitivity to paclitaxel in a number of different melanoma cancer cell lines. Interestingly, the authors also demonstrated for the first time that epigenetic modifications may be involved in regulating βIII-tubulin protein expression. Treatment of a βIII-tubulin-negative melanoma cell line with a histone deacetylase (HDAC) inhibitor markedly induced βIII-tubulin mRNA and protein expression in these cells. A similar effect was also observed in cells which expressed βIII-tubulin. This suggests that loss of βIII-tubulin in these cells may involve modulation of histone acetylation [77].

12.7

Functional Role of β -Tubulin in Tumorigenesis

In addition to their role in chemoresistance, there is an accumulating body of evidence (both clinical and laboratory) to suggest that β-tubulins are also involved in regulating tumor growth and development. Given the importance of microtubules and their associated proteins in regulating cellular processes, such as microtubule dynamics and protein trafficking, it is not surprising that altered expression and/or activity of these proteins could have a major influence on tumorigenesis. As described previously in this chapter, high levels of βIII-tubulin correlate with a poorly differentiated and aggressive tumor phenotype in both epithelial and nonepithelial cancers such as NSCLC, ovarian, and glioblastoma. For example, βIII-tubulin has

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been strongly associated with decreased overall survival in ovarian and NSCLC patients regardless of the response to chemotherapy. However, despite clinical studies linking βIII-tubulin to tumorigenesis, laboratory evidence has been lacking. Recently, our laboratory [49] showed for the first time that long-term suppression of βIII-tubulin directly influenced anchorage-independent growth (a measure of tumorigenic potential) in NSCLC cells. Importantly, stable knockdown of βIII-tubulin also significantly delayed tumor growth and reduced tumor incidence in a mouse subcutaneous xenografted tumor model [49]. Interestingly, this effect was unlikely to be due to reduced cell proliferation, as stable knockdown of βIII-tubulin did not affect cell proliferation in vitro or in vivo, suggesting that other factors associated with βIII-tubulin were at play. The mechanisms whereby βIII-tubulin exerts its effect on tumorigenesis are currently unknown. However, several recent studies have provided some clues as to how increased levels of this microtubule protein may provide cancer cells with a growth advantage. Indeed, it is established that cancer cells survive and proliferate in a harsh tumor microenvironment which includes, hypoxia, glucose deprivation, and oxidant stress. Studies by Raspaglio and Cicchillitti [55, 56] support the idea that βIII-tubulin may act as a survival factor or form part of a network of proteins which actively promote tumor growth under these stressful conditions. In 2008, Raspaglio et al. [56] showed for the first time that βIII-tubulin is controlled at both the gene and protein level by the hypoxia-induced transcription factor HIF-1α. Moreover, Cicchillitti et al. [55] recently demonstrated that βIII-tubulin is able to form proteinprotein complexes with a number of proteins which are involved in promoting cell survival in glucose-deprived and/or high oxidant stress environment [55]. Most recently, De Donato et al. [78] further delineated this survival pathway by identifying a small GTPase (a molecular switch in signaling cascades) which directly interacts with βIII-tubulin in ovarian cancer cells [78]. Importantly, this interaction was shown to be necessary for the recruitment of the oncogene, PIM1 to the cell cytoskeleton to initiate a signaling cascade of proteins which ultimately leads to the enhancement of prosurvival and increased proliferative capabilities of cancer cells. Collectively, these data suggest that βIII-tubulin has a broad role in tumor biology and therapeutic strategies aimed at specifically reducing the expression of βIII-tubulin may have the dual advantage of suppressing cancer growth while enhancing the chemosensitivity of the tumor cells.

12.8

Conclusions and Perspectives

Identifying and understanding the complex interactions of tubulin and microtubule proteins, and their regulatory pathways in cancer is slowly being unraveled. It is clear that differential expression of specific β-tubulin isotypes in tumor cells influence patient response to TBAs and in some cases, broader therapy. Identifying patients that are likely to respond or fail therapy may lead to biomarkers to tailor treatment to optimize response and survival. The full physiological and pathological functions

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of tubulin and microtubule proteins still need to be determined. Limited knowledge on how expression of tubulin–microtubule proteins are differentially regulated in normal cells and cancer cells highlights an area that requires greater research effort. New TBAs that have recently been approved for clinical use such as the epothilones and eribulin (reviewed in [7]) may offer some hope to bypass resistance mechanisms, at least in a subset of patients. Recent evidence that βIII-tubulin is involved in tumorigenesis highlights the complex and multifunctional role of tubulin/microtubule proteins in cancer. Acknowledgments We would like to thank all the researchers who have contributed to our understanding of microtubules, drug resistance, and cancer, and due to space limits we regret that we were not able to cite all the important contributions to the field. Joshua McCarroll and Maria Kavallaris are supported by grants from the National Health and Medical Research Council (NHMRC), Cancer Council New South Wales (MK), Cure Cancer Australia Foundation Grant (JM), Balnaves Young Researcher Award (JM), Cancer Institute New South Wales Early Career Development Fellowship (JM), and an NHMRC Senior Research Fellowship (MK).

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15. Meisel C et al (2000) How to manage individualized drug therapy: application of pharmacogenetic knowledge of drug metabolism and transport. Clin Chem Lab Med 38(9):869–876 16. Berrieman HK, Lind MJ, Cawkwell L (2004) Do beta-tubulin mutations have a role in resistance to chemotherapy? Lancet Oncol 5(3):158–164 17. Giannakakou P et al (1997) Paclitaxel-resistant human ovarian cancer cells have mutant betatubulins that exhibit impaired paclitaxel-driven polymerization. J Biol Chem 272(27):17118– 17125 18. Kavallaris M et al (2001) Multiple microtubule alterations are associated with vinca alkaloid resistance in human leukemia cells. Cancer Res 61(15):5803–5809 19. Green DR, Reed JC (1998) Mitochondria and apoptosis. Science 281(5381):1309–1312 20. Gajate C et al (2000) Induction of apoptosis in leukemic cells by the reversible microtubuledisrupting agent 2-methoxy-5-(2′ ,3′ ,4′ -trimethoxyphenyl)-2,4,6-cycloheptatrien-1-one: protection by Bcl-2 and Bcl-X(L) and cell cycle arrest. Cancer Res 60(10):2651–2659 21. Ibrado AM et al (1996) Bcl-xL overexpression inhibits taxol-induced Yama protease activity and apoptosis. Cell Growth Differ 7(8):1087–1094 22. Tang C et al (1994) High levels of p26BCL-2 oncoprotein retard taxol-induced apoptosis in human pre-B leukemia cells. Leukemia 8(11):1960–1969 23. Deveraux QL, Reed JC (1999) IAP family proteins-suppressors of apoptosis. Genes Dev 13(3):239–252 24. Altieri DC (2001) The molecular basis and potential role of survivin in cancer diagnosis and therapy. Trends Mol Med 7(12):542–547 25. Cheung CH et al (2009) Survivin counteracts the therapeutic effect of microtubule de-stabilizers by stabilizing tubulin polymers. Mol Cancer 8:43 26. Ryan BM et al (2006) Survivin expression in breast cancer predicts clinical outcome and is associated with HER2, VEGF, urokinase plasminogen activator and PAI-1. Ann Oncol 17(4):597–604 27. Schlette EJ et al (2004) Survivin expression predicts poorer prognosis in anaplastic large-cell lymphoma. J Clin Oncol 22(9):1682–1688 28. Sui L et al (2002) Survivin expression and its correlation with cell proliferation and prognosis in epithelial ovarian tumors. Int J Oncol 21(2):315–320 29. Zaffaroni N et al (2002) Expression of the anti-apoptotic gene survivin correlates with taxol resistance in human ovarian cancer. Cell Mol Life Sci 59(8):1406–1412 30. Zhang M et al (2005) Adenovirus-mediated inhibition of survivin expression sensitizes human prostate cancer cells to paclitaxel in vitro and in vivo. Prostate 64(3):293–302 31. O’Brate A, Giannakakou P (2003) The importance of p53 location: nuclear or cytoplasmic zip code? Drug Resist Updat 6(6):313–322 32. Wahl AF et al (1996) Loss of normal p53 function confers sensitization to Taxol by increasing G2/M arrest and apoptosis. Nat Med 2(1):72–79 33. Debernardis D et al (1997) p53 status does not affect sensitivity of human ovarian cancer cell lines to paclitaxel. Cancer Res 57(5):870–874 34. Fan S et al (1998) Disruption of p53 function in immortalized human cells does not affect survival or apoptosis after taxol or vincristine treatment. Clin Cancer Res 4(4):1047–1054 35. King TC et al (2000) p53 mutations do not predict response to paclitaxel in metastatic nonsmall cell lung carcinoma. Cancer 89(4):769–773 36. Malamou-Mitsi V et al (2006) Evaluation of the prognostic and predictive value of p53 and Bcl-2 in breast cancer patients participating in a randomized study with dose-dense sequential adjuvant chemotherapy. Ann Oncol 17(10):1504–1511 37. Luduena RF (1998) Multiple forms of tubulin: different gene products and covalent modifications. Int Rev Cytol 178;207–275 38. Ferlay J et al (2010) Estimates of worldwide burden of cancer in 2008: GLOBOCAN 2008. Int J Cancer 127(12):2893–2917 39. Travis WD et al (2011) International association for the study of lung cancer/american thoracic society/european respiratory society international multidisciplinary classification of lung adenocarcinoma. J Thorac Oncol 6(2):244–285

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40. Rosell R et al (2003) Transcripts in pretreatment biopsies from a three-arm randomized trial in metastatic non-small-cell lung cancer. Oncogene 22(23):3548–3553 41. Seve P et al (2005) Class III beta-tubulin expression in tumor cells predicts response and outcome in patients with non-small cell lung cancer receiving paclitaxel. Mol Cancer Ther 4(12):2001–2007 42. Koh Y et al (2010) Expression of class III beta-tubulin correlates with unfavorable survival outcome in patients with resected non-small cell lung cancer. J Thorac Oncol 5(3):320–325 43. Reiman T et al (2011) Cross-validation study of class III beta-tubulin as a predictive marker for benefit from adjuvant chemotherapy in resected non-small-cell lung cancer: analysis of four randomized trials. Ann Oncol 23(1):86–93 44. Vilmar AC, Santoni-Rugiu E, Sorensen JB (2011) Class III {beta}-tubulin in advanced NSCLC of adenocarcinoma subtype predicts superior outcome in a Randomized Trial. Clin Cancer Res 17(15):5205–5214 45. Kavallaris M et al (1997) Taxol-resistant epithelial ovarian tumors are associated with altered expression of specific beta-tubulin isotypes. J Clin Invest 100(5):1282–1293 46. Kavallaris M, Burkhart CA, Horwitz SB (1999) Antisense oligonucleotides to class III betatubulin sensitize drug-resistant cells to Taxol. Br J Cancer 80(7):1020–1025 47. Gan PP, Pasquier E, Kavallaris M (2007) Class III beta-tubulin mediates sensitivity to chemotherapeutic drugs in non small cell lung cancer. Cancer Res 67(19):9356–9363 48. Gan PP et al (2010) Microtubule dynamics, mitotic arrest, and apoptosis: drug-induced differential effects of betaIII-tubulin. Mol Cancer Ther 9(5):1339–1348 49. McCarroll JA et al (2011) betaIII-tubulin is a multifunctional protein involved in drug sensitivity and tumorigenesis in non-small cell lung cancer. Cancer Res 70(12):4995–5003 50. Sankaranarayanan R, Ferlay J (2006) Worldwide burden of gynaecological cancer: the size of the problem. Best Pract Res Clin Obstet Gynaecol 20(2):207–225 51. Mozzetti S et al (2005) Class III beta-tubulin overexpression is a prominent mechanism of paclitaxel resistance in ovarian cancer patients. Clin Cancer Res 11(1):298–305 52. Ferrandina G et al (2006) Class III beta-tubulin overexpression is a marker of poor clinical outcome in advanced ovarian cancer patients. Clin Cancer Res 12(9):2774–2779 53. Hetland TE et al (2011) Class III beta-tubulin expression in advanced-stage serous ovarian carcinoma effusions is associated with poor survival and primary chemoresistance. Hum Pathol 42(7):1019–1026 54. Ohishi Y et al (2007) Expression of beta-tubulin isotypes in human primary ovarian carcinoma. Gynecol Oncol 105(3):586–592 55. Cicchillitti L et al (2009) Comparative proteomic analysis of paclitaxel sensitive A2780 epithelial ovarian cancer cell line and its resistant counterpart A2780TC1 by 2D-DIGE: the role of ERp57. J Proteome Res 8(4):1902–1912 56. Raspaglio G et al (2008) Hypoxia induces class III beta-tubulin gene expression by HIF-1alpha binding to its 3’ flanking region. Gene 409(1–2):100–108 57. Cortes J, Baselga J (2007) Targeting the microtubules in breast cancer beyond taxanes: the epothilones. Oncologist 12(3):271–280 58. Shalli K et al (2005) Alterations of beta-tubulin isotypes in breast cancer cells resistant to docetaxel. FASEB J 19(10):1299–1301 59. Tommasi S et al (2007) Cytoskeleton and paclitaxel sensitivity in breast cancer: the role of beta-tubulins. Int J Cancer 120(10):2078–2085 60. Stengel C et al (2010) Class III beta-tubulin expression and in vitro resistance to microtubule targeting agents. Br J Cancer 102(2):316–324 61. Jemal A et al (2009) Cancer statistics, 2009. CA Cancer J Clin 59(4):225–249 62. Ploussard G et al (2010) Class III beta-tubulin expression predicts prostate tumor aggressiveness and patient response to docetaxel-based chemotherapy. Cancer Res 70(22):9253–9264 63. Ranganathan S et al (1996) Increase of beta(III)- and beta(IVa)-tubulin isotopes in human prostate carcinoma cells as a result of estramustine resistance. Cancer Res 56(11):2584–2589 64. Terry S et al (2009) Increased expression of class III beta-tubulin in castration-resistant human prostate cancer. Br J Cancer 101(6):951–956

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65. Liu B et al (2001) Taxotere resistance in SUIT Taxotere resistance in pancreatic carcinoma cell line SUIT 2 and its sublines. World J Gastroenterol 7(6):855–859 66. Lee KM et al (2007) Class III beta-tubulin, a marker of resistance to paclitaxel, is overexpressed in pancreatic ductal adenocarcinoma and intraepithelial neoplasia. Histopathology 51(4):539– 546 67. DeAngelis LM (2001) Brain tumors. N Engl J Med 344(2):114–123 68. Katsetos CD et al (2009b) Tubulin targets in the pathobiology and therapy of glioblastoma multiforme. II. gamma-Tubulin. J Cell Physiol 221(3):514–520 69. Katsetos CD et al (2009a) Tubulin targets in the pathobiology and therapy of glioblastoma multiforme. I. Class III beta-tubulin. J Cell Physiol 221(3):505–513 70. Katsetos CD et al (2007) Class III beta-tubulin and gamma-tubulin are co-expressed and form complexes in human glioblastoma cells. Neurochem Res 32(8):1387–1398 71. Katsetos CD, Draber P, Kavallaris M (2011) Targeting III-Tubulin in glioblastoma multiforme: from cell biology and histopathology to cancer therapeutics. Anticancer Agents Med Chem 11:719–728 72. Ikota H et al (2006) Systematic immunohistochemical profiling of 378 brain tumors with 37 antibodies using tissue microarray technology. Acta Neuropathol 111(5):475–482 73. Ives NJ et al (2007) Chemotherapy compared with biochemotherapy for the treatment of metastatic melanoma: a meta-analysis of 18 trials involving 2,621 patients. J Clin Oncol 25(34):5426–5434 74. Zimpfer-Rechner C et al (2003) Randomized phase II study of weekly paclitaxel versus paclitaxel and carboplatin as second-line therapy in disseminated melanoma: a multicentre trial of the Dermatologic Co-operative Oncology Group (DeCOG). Melanoma Res 13(5):531–536 75. Mhaidat NM et al (2008) Melanoma cell sensitivity to Docetaxel-induced apoptosis is determined by class III beta-tubulin levels. FEBS Lett 582(2):267–272 76. Mhaidat NM et al (2007) Regulation of docetaxel-induced apoptosis of human melanoma cells by different isoforms of protein kinase C. Mol Cancer Res 5(10):1073–1081 77. Akasaka K et al (2009) Loss of class III beta-tubulin induced by histone deacetylation is associated with chemosensitivity to paclitaxel in malignant melanoma cells. J Invest Dermatol 129(6):1516–1526 78. De Donato M et al (2011) Class III beta-tubulin and the cytoskeletal gateway for drug resistance in ovarian cancer. J Cell Physiol Apr 25 (Epub ahead of print)

Chapter 13

Posttranslational Modifications of Tubulin Suzan K. Chao, Chia-Ping H. Yang and Susan Band Horwitz

Abstract Recent studies have highlighted the potential importance of posttranslational modifications of tubulin in dictating response to antitumor drugs and disease progression. These modifications include glutamylation, glycylation, phosphorylation, acetylation, and tyrosination. Some of the tubulin-modifying enzymes have been identified but the functional consequences of the posttranslational modifications remain largely unknown. In this chapter, we review the posttranslational modifications of tubulin and current knowledge of the role these alterations may play in human disease.

13.1

Introduction

The cytoskeleton of eukaryotic cells has an amazing ability to respond rapidly to changes in its external and internal environment. A major component of the cytoskeleton is the tubulin/microtubule system, which is highly dynamic and capable of participating in a large number of cellular functions. Microtubules (MTs) play a role in the movement of organelles in the cytoplasm, cell motility, cell shape, and polarity, and they are essential in cell division. The mitotic spindle, which is responsible for normal cell division in which two daughter cells receive equal amounts of parental DNA, is composed of MTs and their interacting proteins. In this capacity, the tubulin/MT system is the target for a number of important antitumor drugs such as the vinca alkaloids and Taxol. Tubulin exists as a heterodimer composed of one α- and one β-tubulin subunit that can be assembled head to tail into MT protofilaments. There are eight isotypes of α-tubulin and seven isotypes of β-tubulin in mammalian cells ([1, 2], Table 13.1). It has been proposed that there is functional significance to this variation, and several in vitro polymerization studies have shown that heterodimers composed of specific isotypes have different dynamic behaviors [3, 4]. While amino acid alterations such as cysteine variation [5] exist throughout the entire sequence of α- and β-tubulin S. B. Horwitz () · S. K. Chao · C.-P. H. Yang Department of Molecular Pharmacology, Albert Einstein College of Medicine, 1300 Morris Park Avenue, Bronx, NY 10461, USA e-mail: [email protected]

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Table 13.1 C-terminal peptides of tubulin isotypes Tubulin isotype

Gene

Accession number

C-terminal peptide

α 1A α 1B α 1C (α6) α 4A α 3C α 3D α 3E α8 βI βII βIII βIVa βIVb βV βVI

TUBA1A TUBA1B TUBA1C TUBA4A TUBA3C TUBA8 TUBAL3 TUBA8 TUBB TUBB2B TUBB3 TUBB4 TUBB2C TUBB6 TUBB1

NP_006000 NP_006073 NP_116093 NP_005991 NP_005992 NP_525125 NP_997195 NP_061816 NP_821133 NP_821080 NP_006077 NP_006078 NP_006079 NP_115914 NP_110400

AALEKDYEEVGVDSVEGEGEEEGEEY AALEKDYEEVGVDSVEGEGEEEGEEY AALEKDYEEVGADSADGEDEGEEY AALEKDYEEVGIDSYEDEDEGEE AALEKDYEEVGVDSVEAEAEEGEEY AALEKDYEEVGVDSVEAEAEEGEEY AALEKDCEEVGVDSVEAEAEEGEEY AALEKDYEEVGTDSFEEENEGEEF YQDATAEEEEDFGEEAEEEA YQDATADEQGEFEEEEGEDEA YEDDEEESEAQGPK YQDATAEEGEFEEEAEEEVA YQDATAEEEGEFEEEAEEEVA YQDATANDGEEAFEDEEEEIDG FQDAKAVLEEDEEVTEEAEMEPEDKGH

isotypes, the majority of the differences are clustered at the C-terminus and this is known as the isotype-defining region [6]. The C-terminal tails are highly acidic and point away from the MT polymer, and are therefore more accessible to microtubuleassociated proteins (MAPs) and other endogenous proteins [6]. In addition to isotype distribution, MTs undergo an unusually large number of posttranslational modifications (PTMs), adding to the structural and functional diversity of these proteins. The expression of tubulin isotypes and MAPs as well as PTMs of tubulin all contribute to the regulation of microtubule dynamics and function. The PTMs include tyrosination, detyrosination, deglutamylation, acetylation, polyglutamylation, polyglycylation, phosphorylation, palmitoylation, and other less well-characterized PTMs. Most of these modifications, with the exception of phosphorylation, occur on the polymerized MT [7]. Interestingly, most of the PTMs are reversible and it has been suggested that there is coordination between PTMs that occur on the microtubule [8]. Most PTMs are found on the carboxyl terminus of tubulin, with the exception of acetylation, which occurs at the amino terminus in the lumen of the microtubule on K40 of α-tubulin. PTMs also have specificity, with some PTMs occurring exclusively on either α- or β-tubulin, and others on both tubulins (Table 13.2). Since enzymes involved in the addition or removal of PTMs show some specificity for the MT substrates they modify, they clearly have a role in regulating the dynamicity and cellular functions of MTs. However, much is still unknown about the regulation and significance of these modifications on tubulin function in vivo, and therefore this area of research is currently of interest to many scientists. Other reviews on the topic of tubulin PTMs have been published [7, 9–14]. This chapter will focus on current knowledge of tubulin PTMs and their role in human disease and its treatment.

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Table 13.2 Posttranslational modifications of tubulin Posttranslational Enzymes, forward Site on α- or modification and reverse β-tubulin

Expression or potential functions

References

Tyrosination Detyrosination

Polyglutamylation

Polyglycylation

Acetylation

Palmitoylation Ubiquitination

Phosphorylation

Removal of alanine and valine

Tubulin tyrosine ligase (TTL) Tubulin carboxypeptidase (CCP1)

α, C-terminus

Many cancers

[28]

α, C-terminus

[16, 19, 28]

Branching: TTLL1, TTLL4, TTLL7, TTLL3 elongation: TTLL6, TTLL9, TTLL10 deglutamylase: CCP5 TTLL3, TTLL8, TTLL10 unknown deglycylase Mec-17 HDAC6, SIRT2

α and β, C-terminal glutamates

Many cancers stabilization of MTs, mitosis, differentiation, migration Mitosis, recruiting motor proteins and MAPs

α and β, C-terminal glutamates

Axonemes cilia and flagella, cytokinesis

[35, 43]

α, K40 in lumen

[57, 61, 62, 65]

Unknown palmitoylase and depalmitoylase BRCA1, parkin, UCHL1 ubiquitin C-terminal hydrolase

α, C376

Stabilization of MTs, cancer drug resistance, cell motility, autophagy Positioning of MTs for bud neck in yeast Cancer tumorigenicity of transformed cells, normal centrosome function and tubulin nucleation Mitosis, cell motility

Casein kinase, PKCα, cdk1, Syk unknown phosphatase Unknown protease

Methylation

Unknown methylase

Arginylation Glycosylation

Unknown arginylase Unknown glycosylase Unknown sumoylase

Sumoylation

α ,β , γ multiple lysines

α and β, β S444, S172; α S165 β IVb-tubulin C-terminal alanine α E434, and β α and β β α

[40, 42, 45]

[72, 73, 74] [80, 89, 91, 92]

[99, 105]

Liver cancer

[113]

Toxoplasma gondii host cell invasion Tubulin degradation Chemotherapy resistance Tubulin degradation

Xiao et al. 2010 [117] [115] [116]

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Fig. 13.1 Tubulin tyrosine ligase cycle. The cycle begins with Tyr-tubulin being detyrosinated by the enzyme TCP. The resulting tubulin, Glu-tubulin can have its C-terminal glutamate removed by the enzyme TDG. The resulting 2 tubulin can no longer be retyrosinated. Glu-tubulin can be retyrosinated by the enzyme TTL and resume the cycle again

13.2 Tyrosination and Detyrosination The most well-studied tubulin PTMs are tyrosination and detyrosination. It has been established that α-tubulin cycles between being C-terminally tyrosinated and detyrosinated. Six out of the eight mammalian α-tubulin isotypes have a tyrosine residue as the terminal amino acid in the polypeptide. Although α4-tubulin has a glutamate as the terminal residue (Table 13.1), it is also subject to the tyrosination cycle, because it enters at the tyrosination stage of the cycle. The terminal tyrosine of α-tubulin is removed by a tubulin-specific carboxypeptidase (TTCP, [15]), leaving glutamate as the terminal amino acid [16]. Detyrosination takes place exclusively on α-tubulin at its carboxyl terminus and occurs on the nascent α-tubulin peptide almost immediately after its translation [17]. A tubulin that has been detyrosinated is also known as “Glu-tubulin” because glutamate becomes the terminal amino acid following detyrosination [18]. It was discovered in 1993 that Glu-tubulin can be retyrosinated by the enzyme tubulin tyrosine ligase (TTL) in an ATP-dependent reaction and the resulting tyrosinated tubulin is referred to as Tyr-tubulin [19]. This cycle of detyrosinating and tyrosinating is known as the TTL cycle (Fig. 13.1).

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There is an increase in abundance of Glu-tubulin in differentiated cells, which have a larger population of stabilized MTs, compared to dividing cells [20–22]. Stabilized MTs have a slower turnover rate and are exposed to TTCP for a longer period of time, resulting in an increased population of Glu-tubulin. Therefore, Glu-tubulin has been associated with MT stability [20, 21, 23]. Many cancers have decreased expression of TTL resulting in increased levels of Glu-tubulin. Its importance in cancer progression was discerned when NIH3T3 cells deficient for TTL were able to induce the growth of tumors when injected into nude mice. Inhibition of the TTL cycle is a known occurrence during tumor progression [24]. By immunohistochemical staining, it has been shown that the levels of Glu-tubulin are increased in breast cancer [25], prostate cancer [26], and neuroblastomas [27]. It has recently been shown in breast cancer that there is a strong correlation between Glu-tubulin levels and Scarf-BloomRichardson (SBR) grade, a known marker of tumor aggressiveness used for breast cancer prognosis. In particular, there was a correlation of Glu-tubulin with mitotic score [25]. This is in agreement with studies that have shown that Glu-tubulin levels fluctuate in accordance with the cell cycle and it has been proposed that suppression of TTL confers some growth advantage to tumor cells, although the precise mechanism is unknown [24, 25]. In some instances, the penultimate glutamate or amino acid preceding the terminal tyrosine of α-tubulin, is removed by deglutamylation and the resulting tubulin is referred to as “2 tubulin” [28]. 2 Tubulin can no longer enter the TTL cycle because TTL does not recognize 2 tubulin as a substrate [29]. 2 tubulin was first discovered in tubulin isolated from the brain, where its levels are elevated. It is usually not detectable in normal tissues except the brain [30]. However, some cancers including breast [25] and prostate [26] have increased levels of 2 tubulin, though not to the same extent as Glu-tubulin levels are increased in these cancers. In spite of this, 2 tubulin is considered a better marker of decreased TTL activity than Glu-tubulin since its levels do not fluctuate with the cell cycle [20, 31, 32]. Importantly, 2 tubulin provided the first example of coordination between tubulin PTMs. Banerjee et al. discovered in 1985, using purified bovine brain tubulin, that tyrosinated α-tubulin was polyglutamylated, while 2 tubulin was preferentially polyglycylated [8]. This selectivity and complexity of regulation occurring on the C-terminal tails of MTs is reminiscent of histone modifications, which also occur sequentially on the exposed tail region of histones and regulate the availability of chromatin for subsequent transcription. This analogy has brought about the concept of a “tubulin code” that has similarities to the “histone code.” PTMs on MTs may determine when cells carry out specific functions that require restructuring of MTs for crucial cellular events such as cell division and intracellular trafficking [13, 14, 33].

13.3

Polyglutamylation and Polyglycylation

Polyglutamylation and polyglycylation are rare PTMs once thought only to occur on MTs, but have since been found on other proteins [34]. While polyglycylation has been limited in mammalian cells to axonemes of cilia and flagella that are composed

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Fig. 13.2 Polyglutamylation of tubulin. Addition of glutamates occurs in two steps. The first glutamate is added by an isopeptide bond, this is the branching step. The subsequent glutamates are added by peptide bonds on the alpha carbons of each addition, this is the elongation step

of MTs [35, 36], polyglutamylation has been found in the MTs of centrioles [37], neurons [38], and the spindle of dividing cells [39]. Both of these PTMs occur on conserved glutamate residues at the C-terminus of α- or β-tubulin [35, 40, 41], and create covalently bonded side chains of either glycines or glutamates that can be up to 30 amino acids long. Attachment of these long side chains happens in two steps; first branching and then elongation. In the branching step, an isopeptide bond is formed on the gamma carboxylate group of a conserved C-terminal glutamate residue. In elongation, another isopeptide bond or α peptide bond is formed with the newly attached glutamate on its gamma carboxylate group (Fig. 13.2). The enzymes responsible for polyglutamylation and polyglycylation are TTL-like (TTLL) proteins (Table 13.2, [42–44]). It is worth noting that one TTLL is responsible for the branching step, while a different TTLL is responsible for elongation [34]. Recently, a family of deglutamylases has been identified, they include CCP1, CCP4, CCP5, and CCP6 (Table 13.2, [45, 46]). These enzymes are also involved in the detyrosination of tubulin, further indicating that there is coordination among tubulin PTMs [46]. Glutamylated tubulin is increased during the G2 phase of the cell cycle, and it is believed that this increase may reflect the need for rapid regulation of mitotic MTs during cell division to coordinate the binding of motor proteins such as kinesins [47, 48]. Therefore, it seems likely that there is spatial and temporal regulation of PTMs on tubulin to enable tight regulation of the cell cycle. Glutamylation increases the net negative charge of an already negatively charged tubulin, and this may enhance the interaction of MAPs with the acidic C-terminus

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of tubulin. Spastin and katanin are MAPs that function in MT severing and are regulated by polyglutamylation of tubulin [49]. These proteins are positively charged and are attracted to the negatively charged polyglutamylated tubulin tail. Long glutamate side chains are able to induce MT severing in HeLa cells and are localized to the mitotic spindle, thereby indicating that polyglutamylated tubulin regulates spindle dynamics and chromosome movement during cell division [49]. Furthermore, disruption of spastin has been seen in neurodegeneration and spastic paraplesia, suggesting that the level of MT polyglutamylation may play a role in neurodegenerative diseases [49]. Another protein that is regulated by polyglutamination is tau [50], a microtubule-stabilizing protein, which has been associated with altering MT dynamics and possibly contributing to taxane and ixabepilone drug resistance [51–53]. Several groups have shown that patients with low expression of tau respond better to treatment with Taxol, a microtubule-stabilizing agent [51, 54]. Additional MAPs that are regulated by polyglutamylation are MAP1a and the motor protein, kinesin. Intriguingly, MAP1a prefers tubulin glutamate side chains that consist of approximately six glutamates [55], while kinesin1 prefers side chains of only three [56]. This suggests that the number of PTMs can also influence microtubule dynamics, further supporting the idea of the tubulin code.

13.4 Acetylation Perhaps the most well-known posttranslational modification of tubulin is the acetylation of α-tubulin. Acetylation of α-tubulin is most often associated with stabilized MTs. A monoclonal antibody specific for this modification has been widely used as a marker of stabilized MTs [57, 58]. As has been discussed in other parts of this book, many compounds used to treat malignant diseases target tubulin. It has been established that MTs have increased levels of K40 acetylation after treatment of cells with Taxol, a microtubule-stabilizing agent. Cellular MTs that are resistant to depolymerizing agents—such as the vinca alkaloids, nocodazole, and colchicine—contain more acetylated α-tubulin than the rest of the cytoplasmic MTs [59]. In a recent study, it was discovered that treatment with MT-stabilizing agents such as Taxol, epothilone B, or discodermolide was sufficient to induce tubulin acetylation in a human ovarian cancer cell line, Hey. Vinblastine, on the other hand, had essentially no effect on K40 acetylation [60]. Only recently has MEC17—a K40-specific tubulin acetyl transferase—been identified in tetrahymena [61]. A human tubulin acetyl trasferase is yet to be identified. Thus far, only two human tubulin deacetylases have been characterized, histone deacetylase 6 (HDAC6, [58, 62–64]) and sirtuin 2 (SIRT2 [65]). HDAC6 is a tubulin deacetylase that is constitutively active [65], ubiquitously expressed and conserved among species. Inhibition of HDAC6 increases acetylation of α-tubulin and may result in increased stability of MTs [63]. In addition, HDAC6 overexpression and silencing have been shown to influence cell motility [58, 64]. SIRT2 is an NAD+ dependent deacetylase that has high specificity for acetylated tubulin peptides in vitro

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and has been shown to deacetylate tubulin in vivo [65]. SIRT2 is phosphorylated by cyclinE-cdk2 on S331. Upon phosphorylation, SIRT2 deacetylase activity toward α-tubulin is reduced and it has been proposed that this is a way in which cell cycle progression can be controlled [66]. Unlike other HDACs, HDAC6 and SIRT2 are largely cytoplasmic with less localization in the nucleus. Both deacetylases coimmunoprecipitate and colocalize together with MTs in 293 T cells and are able to acetylate tubulin in addition to histones, although the preferred substrate is tubulin [65, 67]. In particular, HDAC6 expression has been correlated with clinical prognosis in breast cancer patient samples [68, 69]. HDAC inhibitors have entered clinical development in recent years for treatment of cancer [70]. Not only the HDAC inhibitors are promising anticancer agents because of their effects on gene expression through histone acetylation but also due to their effects on nonhistone substrates such as α-tubulin. In particular, tubucin, a small molecule that specifically inhibits HDAC6, has been shown to decrease cell motility, which plays an important role in metastasis. HDAC inhibitors in combination with a variety of chemotherapeutics have enhanced efficacy in cancer cells [70]. Identification of markers to determine the efficacy of HDAC inhibitors for use in patients is lacking. However, recent studies have identified reduced acetylation of α-tubulin in ductal carcinoma in situ and in invasive ductal carcinoma of the breast relative to normal mammary epithelium, suggesting that α-tubulin acetylation status may be useful as a marker of breast cancer progression [71].

13.5

Palmitoylation

Tubulin is posttranslationally modified in vivo by palmitoylation, the covalent attachment of the long-chain fatty acid palmitate to cysteine residues. Tubulin palmitoylation has been found in vivo in studies using human platelets and PC-12 cells [72, 73]. In these cells, both α- and β-tubulin were labeled with [3 H] palmitate, however, α-tubulin was labeled to a greater extent than β-tubulin [72, 73]. This PTM contributes to the association of tubulin with intracellular membranes. There are 20 cysteine residues in an α-β-tubulin heterodimer, although the major site for palmitoylation was reported to be on C376 of α-tubulin [74]. Recent studies using acyl-biotinyl exchange chemistry and mass spectrometry have determined that 11 cysteine residues of the α-β tubulin heterodimer from rat brain tubulin were palmitoylated in vitro [75]. In a cell-free system, the substrates for palmitoylation were either nonpolymerized tubulin or MTs assembled with slowly hydrolyzable GTP analogs. Taxol assembled MTs were not a substrate for palmitoylation, probably because the SH-groups of cysteine were not accessible for palmitoylation in Taxol-polymerized MTs [72]. It has also been demonstrated that palmitoylation of tubulin in vitro is inhibited by stoichiometric concentrations of MT-depolymerizing drugs, such as colcemid, nocodazole, and vinblastine, although the mechanism of inhibition is not yet elucidated [72].

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It is not clear whether tubulin palmitoylation takes place enzymatically or through spontaneous S-acylation (autopalmitoylation). It has been reported that pure α- and β-tubulin were readily palmitoylated in vitro using [3 H] palmitoyl CoA without added enzyme. A maximum of approximately six palmitic acids were added per dimer in a few hours. Palmitoylation of tubulin resulted in a marked reduction in polymerization competence [76, 77]. A possible role for α-tubulin palmitoylation has been studied using a strain of Saccharomyces cerevisiae that harbors a Cys to Ser mutation at C377 of α-tubulin in tub1, the putative site of palmitoylation in α-tubulin in the wild-type yeast strain [78]. The astral MTs in these mutant cells appeared to be excessive in number, abnormally long, and in some cases misoriented, compared with the wild-type cells. In addition, translocation of spindles through the bud neck was altered. As the level of palmitoylated α-tubulin on C377 S tub1 cells is markedly reduced compared with the control cells, it was suggested that palmitoylation of C377 may play a role in some astral MT functions, such as nuclear migration during metaphase and nuclear positioning during anaphase in this yeast strain [78]. The functional role of palmitoylation was later studied in human CEM leukemic lymphocytes. A clinically relevant dose of vinblastine was able to inhibit tubulin palmitoylation, MT disassembly, and apoptosis in CEM cells [79]. The effect of vinbastine appeared to be specific for tubulin, since palmitoylation of nontubulin proteins was not influenced by this drug. Therefore, it has been suggested that depalmitoylation of tubulin may be a target for new chemotherapeutic drugs [79].

13.6

Ubiquitination

Misfolded tubulin monomers are highly toxic to the cell and must therefore be degraded quickly [80]. One mechanism for marking proteins for degradation is ubiquitination. In recent years, ubiquitination of α-, β-, and γ-tubulin has been described as a means for regulating tubulin turnover. For example, α-β-tubulin heterodimers are ubiquitinated by parkin, an E3 ubiquitin ligase that has been associated with Parkinson’s disease [80]. Parkin coimmunoprecipitates with tubulin heterodimers and can be seen colocalizing with MTs by immunofluorescence in neurons and glial cells [80]. Parkin associates with MTs mostly through hydrophobic interactions and binds to them more tightly than even MAP1A or MAP2, which bind to MTs via electrostatic interactions [80, 81]. In addition, parkin overexpression in HEK293 cells increases tubulin ubiquitination and degradation demonstrating that parkin has a role in regulating microtubule turnover. UCHL1, also known as PGP9.5, is a unique ubiquitin editing enzyme that acts to both ubiquitinate and deubiquitinate substrates [82], and is known to associate with tubulin [83]. Altered expression of UCHL1 has been observed in pancreatic [84], lung [85], and breast cancers [86], as well as in neurodegenerative disorders [87]. Wildtype UCHL1 colocalizes with the mitotic spindle and its level of expression increases during the G2-M phase of the cell cycle. Furthermore, overexpression of UCHL1 was

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shown in vitro to decrease MT polymer formation, indicating a strong association of UCHL1 and tubulin polymerization. This link between tubulin polymerization and UCHL1 was originally identified using a mutant of UCHL1, UCHL1 I93A, which has a strong correlation with dominant inherited Parkinson’s disease [88]. Use of this mutant to co-IP for interacting proteins led to the identification of tubulin as an interacting partner. In this study, UCHL1 I93M associated with MTs and induced tubulin polymerization [89] suggesting an involvement of tubulin ubiquitination in heritable Parkinson’s disease. BRCA1 is a tumor suppressor and loss of BRCA1 expression is a common characteristic in breast and ovarian cancers [90]. BRCA1 binds to another protein, BARD1 (BRCA1-associated ring domain protein) and together the BRCA1/BARD1 complex acts as an E3 ubiquitin ligase. BRCA1/BARD1 localizes to centrosomes [91] and one of its known substrates is γ-tubulin [92]. Ubiquitination of γ-tubulin is necessary for MT nucleation and normal centrosomal function [93]. BRCA1/BARD1 regulates centrosomal function by inhibiting MT nucleation [94] and controlling γ-tubulin localization to centrosomes during cell division [95]. Loss of BRCA1 results in aberrant centrosomal duplication, yielding cells with both numerous and hyperactive centrosomes [92]. These studies indicate that ubiquitination of tubulin may play a role in preventing the chromosomal instability that is characteristic of cancer cells.

13.7

Phosphorylation

Phosphorylation, though a well-described PTM for the regulation of various proteins, has not been widely studied in the regulation of tubulin and its functional relevance to MT dynamics is largely unknown. However, the first posttranslational modification of tubulin to be described in 1970 was phosphorylation [96, 97]. Later, using rat brain extracts incubated with 32 PO4 , Eipper et al. concluded that a single serine on tubulin undergoes phosphorylation [98]. The specific tubulin isotype that was modified was unknown until Gard and Kirschner, in 1985, using a mouse neuroblastoma cell line, and 32 P labeling combined with isoelectric focusing were able to identify the modified tubulin isotype as β2 . Treatment with Taxol, increased β2 -tubulin phosphorylation, while treatment with destabilizing agents reduced phosphorylation suggesting that phosphorylation likely reflected the amount of polymerized MTs in the cell [99]. This particular tubulin isotype was extensively studied and is now known as βIIItubulin (encoded by TUBB3 gene, [100]). These studies laid the groundwork for future important discoveries, including the identification of βIII-tubulin as a marker of neuronal differentiation and of Taxol resistance in patients [96, 101, 102]. Several kinases such as cAMP-dependent kinases [103], casein kinase II [104], cdk1 [105], PKC [106], and Syk [107–109] that may be responsible for phosphorylating tubulin have been identified. However, the ability of many kinases to phosphorylate tubulin in vitro does not indicate that tubulin is the preferred substrate in vivo. Several studies in recent years have substantiated the effects of tubulin phosphorylation on MT dynamics by utilizing phosphorylation-site-specific antibodies.

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For example, β-tubulin is a substrate of Cdk1, a cyclin-dependent kinase important for entry into mitosis. In this study, extracts of mitotic HeLa cells were probed with a phospho-specific antibody and shown to have increased phosphorylation of tubulin at S172, indicating a role in regulating cell division [105]. Both α- and β-tubulins undergo phosphorylation. α-Tubulin is phosphorylated by the nonreceptor tyrosine-kinase, Syk, and by PKCα, a serine-/threonine-specific protein kinase [106, 109]. Syk was shown to phosphorylate MTs in activated B-lymphocytes [107]. This was confirmed by the use of a specific inhibitor of Syk kinase, piceatannol, which was able to block phosphorylation of tubulin. In another study, it was demonstrated that in B-cell lines that did not express Syk, α-tubulin was unphosphorylated [109]. These studies demonstrate that MTs provide important scaffolds for signaling [110] and indicate that PTMs could act to recruit various kinases to specific subcellular compartments. PKCα expression is known to contribute to the progression of particular cancers and the level of overexpression or underexpression is cancer type-specific [111]. PKCα phosphorylates α6-tubulin on S165 both in vitro and in vivo [106, 112] and overexpression in the human breast cell line MCF-10A increased cell motility, indicating a potential role in cancer metastasis [106]. It has been suggested that phosphorylation of α6-tubulin on S165 by PKCα and β-tubulin on S172 by Syk may be involved in regulating MT polymerization due to their effects on mitosis and cell motility. This is perhaps due to their position at the α-β heterodimer interface. In particular, S165 is located on α-tubulin near the plus end of the growing MT and could possibly enhance the GTP hydrolysis necessary for elongating MT protofilaments [106].

13.8 Additional PTMs of Tubulin (Demethylation and C-Terminal Truncation) New PTMs of tubulin are currently being identified. Recently, a novel PTM of tubulin was discovered in rat liver cancer, a truncated form of βIVb-tubulin, βIVb∗ , was identified by mass spectrometry. βIVb∗ lacks the C-terminal valine and alanine, exposing glutamate as the final residue, and is expressed approximately threefold more than βIVb in tumor tissue relative to control tissue [113]. The levels of βIVb∗ were also elevated in human lung tissue containing liver metastases [113]. The first evidence of tubulin methylation was observed in Toxoplamsagondii, an intracellular parasite and human pathogen. In this study, both α- and β-tubulin were methylated. Importantly, toxoplamsagondii has specialized MTs that play an important role in host invasion [114], suggesting that this PTM may be important in pathogenesis. Additional PTMs that have been identified in proteomics screens—but have not been extensively studied—are sumoylation, arginylation, and glycosylation [115– 118]. Both sumoylation and arginylation are PTMs that are involved in protein

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stability. Glycosylation was found to be specific to βIII-tubulin in a study involving chemotherapy drug-resistant and drug-sensitive cancer cell lines. In this study, glycosylation and phosphorylation of βIII-tubulin correlated with drug resistance, indicating that glycosylation could be a potential marker for resistance to MT interacting agents [115].

13.9

Future Directions

Proteomic approaches have enabled the identification of several new PTMs of tubulin, however, many of the modifying enzymes have yet to be discovered. This provides an exciting new area of research since it has been suggested that there likely is subcellular compartmentalization of these modifying enzymes in order to rapidly reorganize the cytoskeleton during cell division or cellular stress [13]. For example, the localization of Rho-GTPases has a role in cell motility by modulating the actin cytoskeleton [119]. In addition, the exact role of these modifications on MT dynamicity is largely unknown. For example, treatment with Taxol is known to stabilize MTs and increase levels of tubulin acetylation and detyrosination [59, 60], but it is unknown whether these PTMs influence the dynamicity of MTs or if they are a consequence of MT stability. Furthermore, the accumulation of PTMs could be due to stabilized MTs possessing a longer half-life than that of unpolymerized tubulin [23, 120], thereby allowing the accumulation of PTMs. Recent studies, such as those described in drug-resistant cells [115] and liver cancer using mass spectrometry [113], convey the necessity of researching these PTMs to find new therapies and markers of disease progression.

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Chapter 14

Stathmin and Cancer Dominic Chi Hiung Ng and Frances Byrne

Abstract Stathmin (STMN) is an evolutionarily conserved, ubiquitously expressed, tubulin-binding protein that has been associated with cell proliferation and differentiation. The widely reported over-expression of STMN in human malignancies suggests significant function in cancer and may represent a viable target to reduce disease. We discuss the transcriptional and post-transcriptional regulation of STMN expression and review the complex signalling networks that impact STMN activity through post-translational modification and protein interactions. Recent advances have also revealed STMN contributions towards chromosome instability leading to aneuploidy, cell migration/invasion and chemoresistance. These studies highlight that STMN potentially contributes to tumour development and progression through the regulation of multiple cellular processes. Reports of STMN interaction with anti-cancer drugs have also provided new insights on STMN mechanisms in drug-refractory malignancies and highlighted the potential of combining STMN inhibition with established therapies. These advances provide fundamental information necessary to evaluate the therapeutic value of targeting STMN in the treatment of cancer.

14.1

Introduction

Microtubules (MTs) are hollow protein filaments comprising heterodimeric α-/βtubulin sub-units that can change between phases of self-assembly and disassembly with either fast or slow kinetics [1]. The dynamic nature of MTs in cells is exquisitely orchestrated by a network of MT-regulatory proteins which promote polymer assembly (stabilisers) or promote disassembly (destabilisers; [1]). Stathmin (STMN) represents one of the few regulators identified with MT-destabilising properties and is critically involved in the control of MT dynamics in vitro and in vivo. D. C. H. Ng () Department of Biochemistry and Molecular Biology, Bio21 Molecular Science and Biotechnology Institute, University of Melbourne, 30 Flemington Road, Parkville, VIC 3010, Australia e-mail: [email protected] F. Byrne Children’s Cancer Institute Australia, Lowy Cancer Research Centre, University of New South Wales, 2052 Sydney, NSW, Australia Australian Centre for Nanomedicine, Faculty of Engineering, University of New South Wales, Sydney, NSW, Australia M. Kavallaris (ed.), Cytoskeleton and Human Disease, DOI 10.1007/978-1-61779-788-0_14, © Springer Science+Business Media, LLC 2012

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Fig. 14.1 Members of stathmin (STMN) family of microtubule-destabilising proteins. STMN family members consist of high homology C-terminal stathmin-like domains (SLD) which bind tubulin through α-helical regions (grey). The microtubule-destabilising activity of STMN family proteins are regulated by multi-site serine/threonine phosphorylation on highly conserved residues shown (inverted triangle). STMN-like SCG10, SCLIP and RB3 contain an additional N-terminal region (black) which is lipid-modified, anchoring these proteins to vesicular membranes associated with the Golgi apparatus

STMN was discovered over two decades ago as a protein phosphorylated in response to a wide range of extracellular stimuli [2] and thus named after ‘stathmos’, the Greek term for relay [3]. It was later separately identified as a differentially expressed protein in leukemia cells [4] and subsequent studies indicating strong associations with cancer led to STMN being commonly known as oncoprotein 18 (Op18). Perhaps surprisingly the MT-regulatory properties of STMN were only later revealed in a study which sought to identify MT-destabilising factors [5]. Since those pioneering studies, increased STMN levels have been linked to a wide variety of human malignancies and a great deal has been revealed regarding STMN function in cancer cells and the complex mechanisms involved in regulating STMN activity. STMN is the archetypal member of a family of proteins that includes SCG10, SCLIP and RB3 (alternatively spliced into RB3′ and RB3′′ ). STMN is a 17-kDa (149 aa) cytosolic protein with universal tissue distribution but is particularly enriched in the brain [3]. The STMN-like proteins share high sequence homology with STMN at their C-terminus (termed stathmin-like domains, SLDs) but are extended at the N-terminus which is variable in sequence (Fig. 14.1). The N-terminal regions of STMN-like proteins are additionally lipid modified, anchoring these proteins predominantly to vesicular membranes of the Golgi apparatus (Fig. 14.1; [6]). Like

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STMN, the STMN-like proteins are also enriched in neuronal tissue but their expression in other tissues varies from moderate to negligible levels [7]. In this chapter, we review the current views on the contribution of STMN-regulated MT dynamics in cancer cell processes, the regulatory mechanisms controlling STMN activity and their implications in cancer progression and treatment.

14.2

STMN Regulation of MT Dynamics

The MT destabilising properties of STMN were first characterised by Belmont and Mitchison who identified a small heat stable protein that could bind unpolymerised tubulin and promote MT catastrophe (the transition of MTs from growth to shrinking phase; [5]). Indeed, the alteration of STMN levels can have a profound effect on MTs with STMN over-expression in cells reducing MT polymer levels [6]. Conversely, STMN knockdown increases MT polymer levels but does not increase stability of microtubules per se (e.g. resistance to low-dose nocodazole is unchanged; [8, 9]). In addition to MT polymer levels, STMN activity regulates the dynamic instability of MT plus and minus ends and MT treadmilling (simultaneous plus-end growth and minus-end shortening without length change; [10, 11]).

14.2.1

STMN Secondary Structure and the T2S Ternary Complex

Biochemical and biophysical analyses of recombinant STMN incubated with purified tubulin revealed that STMN bound two molecules of heterodimeric α-/β-tubulin in a T2 S ternary complex (Fig. 14.2a; [12]). Investigations into the secondary structure of STMN indicated a relatively unstructured N-terminus and α-helical core domains spanning the C-terminal half of the protein [13]. The helical core domain (aa 42– 126) was shown to be sufficient for tubulin interaction [13–15]. Studies utilising truncation mutants of STMN identified multiple regions within the helical core that were capable of tubulin association but the higher-order T2 S structure required the full-length helix [13, 14]. The conserved SLD of STMN-like proteins likewise bound tubulin heterodimers in a 2:1 molar ratio with varying affinities and interaction kinetics and all display MT-destabilising properties [16]. Of the STMN family members, the SLD of RB3 binds tubulin with the highest stability and slowest dissociation rates [16] which has facilitated structural delineation of RB3 SLD/tubulin complex (Fig. 14.2b; [17, 18]). This has provided valuable insight into the nature of tubulin association by STMN and the STMN-like proteins. The bound tubulin sub-units are arranged in headto-tail fashion and contact RB3 SLD through a 91-residue α-helical C-terminal region [17]. This is consistent with the tubulin interaction region identified in previous biochemical studies and verified that tubulin associates with two distinct sites on the α-helix [14, 15, 17]. The N-terminus of STMN forms a β-hairpin structure which interacts with α1-tubulin, ‘capping’ one end of the tubulin-SLD ternary

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Fig. 14.2 The STMN: tubulin ternary complex. a Schematic representation of STMN bound to two α-/β-tubulin heterodimers in a T2 S complex. Tubulin-interacting helical domains and conserved serine phosphorylation sites are indicated. b X-ray structure of tubulin: RB3-SLD complex drawn with Jmol Java applet using published data deposited with Protein Data Bank (PDB ID: 1SAO)

complex (Fig. 14.2; [18]). Furthermore, the crystal structure of the T2 S complex indicates that the STMN may interact and stabilise two tubulin heterodimers in a curved conformation (Fig. 14.2; [17, 18]). The curved T2 S complex, coupled with N-terminal capping, likely prevents the longitudinal incorporation of bound tubulin heterodimers to the growing ends of MT polymers.

14.2.2

Tubulin Sequestration and Catastrophe Promotion

The observation that tubulin bound in the T2 S complex is unable to participate in MT assembly has led to a tubulin sequestration model for STMN-mediated MT destabilisation. In this model, high-affinity interactions of STMN with tubulin, reduce the pool of tubulin available for MT assembly. This slows MT growth rates and increases MT catastrophe which is dependent on free tubulin concentrations [8]. However, whether tubulin sequestration is the sole mechanism by which STMN regulates MT dynamics remains under question. For example, the low expression levels of STMN relative to tubulin levels in some cell types [19] casts doubt on tubulin sequestration as a universal mechanism. In addition, under physiological conditions (pH, temperature, ionic strength), the stability of the T2 S complex is low and whether this is sufficient to alter tubulin partitioning in cells through a tubulin sequestration model has been challenged [20]. In the in vitro setting the stability of the T2 S complex varies depending on buffer conditions. The STMN-tubulin-binding affinity is optimal at pH 6.5 [8] but decreases as pH rises towards physiological pH [20]. An alternative proposal is that STMN possesses tubulin-directed activities that are independent of its capacity to sequester tubulin. That is, STMN could specifically promote increases in the rate of MT catastrophes through association with tubulin or directly to MTs. Indeed, C-terminal truncation mutants of STMN that are deficient in tubulin sequestration are still able to induce MT polymer loss when expressed in cells [10, 21]. Similarly, expression of

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a catastrophe-promoting N-terminal STMN derivative caused accumulation of abnormal mitotic spindles whereas over-expression of tubulin-sequestering C-terminal STMN derivative did not [20, 22]. An exhaustive analysis of the evidence supporting tubulin sequestration or catastrophe-promoting mechanisms is beyond the scope of this chapter but has been elegantly reviewed previously [23]. In summary, the precise function of catastrophe-promoting or tubulin-sequestering STMN activities in cells is still to be fully resolved and likely depends on cellular context.

14.2.3

Post-Transcriptional Regulation of Tubulin

Emerging evidence suggests that in addition to controlling tubulin monomer-polymer partitioning STMN regulation of MTs may include actions on tubulin synthesis and degradation. STMN has been shown to prevent the loss of unpolymerised α- and β-tubulin sub-units induced by over-expression of proteins (such as the tubulinspecific co-factors, TBCE and E-like) that regulate tubulin turnover and promote tubulin degradation [24]. In addition, STMN may also positively regulate tubulin heterodimer levels through a post-transcriptional mechanism [25]. STMN depletion by siRNA in cultured T cells resulted in a decrease in mature tubulin mRNA levels without changes in tubulin transcription [25]. This indicated post-transcriptional regulation by STMN potentially by promoting tubulin mRNA stability [25]. In support of this, STMN over-expression increases tubulin mRNA levels and could counter-act tubulin mRNA destabilisation induced by an MT-depolymerising drug, colchicine [25]. The effect of STMN over-expression on tubulin mRNA stability also mirrors the effect of MT-targeting drug, taxol [25]. These observations led to the suggestion that STMN expression could control the expression of tubulin via similar auto-regulatory mechanisms that regulate the stability of polysomal tubulin mRNA (i.e. co-translational inhibition by tubulin monomers), initially elucidated with MTtargeting drugs [26]. Whether this involves formation of the ternary STMN-tubulin complex is unclear [26]. Interestingly, these findings hint at potential mechanisms by which increased STMN levels, commonly observed in cancer, could antagonise the actions of some MT-targeting drugs.

14.3 14.3.1

Regulation of STMN Activity Phosphorylation

The activity of STMN is exquisitely regulated by multi-site phosphorylation on four highly conserved serine residues (S16, S25, S38, S63; Fig. 14.1), which negatively impacts its MT-destabilising activities [27]. The phosphorylation of individual serine residues leads to varying degrees of inhibition of STMN activity [27, 28]. STMN S16 phosphorylation near the N-terminal β-hairpin loop structure is thought to impede

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binding to the α1-tubulin sub-unit while STMN S63 phosphorylation has been shown to disrupt the central helix core domain and prevent T2 S complex formation [18, 29]. Reflecting the important role of these serine residues in STMN regulation, the phosphorylation of STMN S16 and S63 in vitro contributes substantially to the inhibition of STMN binding to tubulin and activities in decreasing MT polymerisation and promoting MT catastrophe [11, 28]. In contrast, STMN phosphorylated on S25 and S38 binds tubulin and has marginal effects on the tubulin-sequestering and catastrophepromoting activities [11, 27, 28]. However, complete phosphorylation of all four serine residues, as observed during mitosis, results in the greatest inhibition of STMN activity [27, 28]. During mitosis, stathmin is phosphorylated to high stoichiometry (dubbed STMN hyper-phosphorylation) allowing spindle assembly and ordered progression through to anaphase [30]. The S25 and S38 residue on STMN are located in a disordered linker region that does not interact with tubulin sub-units [17, 18]. Thus, it is unclear how STMN S25 and S38 phosphorylation contributes to inhibition of MT-destabilising activities. However, STMN S25 and S38 phosphorylation appears to be required for subsequent S16 and S63 phosphorylation during mitosis [27]. In support of this, an S25A/S38A double mutant of STMN that cannot be phosphorylated on these critical sites leads to aberrant mitotic spindle formation, transient G2 -M arrest followed by S phase progression without proper separation of chromosomes and thus results in endore duplication [30, 31]. These findings highlight the importance of STMN hyper-phosphorylation in the context of cell division [32]. Thus, in the case of mitosis, a mechanism of sequential phosphorylation triggered by STMN S25/S38 may be important for complete inactivation of STMN MT-destabilising activity. To date, a similar mechanism of hierarchal STMN phosphorylation regulated by STMN S25/S38 residues in the interphase cell has not been reported. Multi-site serine phosphorylation of STMN is regulated by a number of different kinases (Fig. 14.3). During mitosis, STMN is phosphorylated on S25 and S38 by cyclin-dependent protein kinases (CDK; [30]). The mitotic kinases involved in targeting STMN S16 and S63 remain undefined. However, several kinases localised to mitotic chromatin, such as polo-like kinase Plx1 and Aurora-B kinase, are likely to be involved as their inhibition reduces STMN hyper-phosphorylation during mitotic progression [33, 34]. In the later stages of mitosis, STMN dephosphorylation by serine/threonine phosphatases sensitive to okadaic acid (i.e. PP1, PP2A) is required for completion and exit from mitosis [35]. Under non-mitotic conditions, other kinases known to directly target phosphorylation of STMN on S16 and S63 include cAMP-dependent protein kinase (PKA), Ca2+ -/calmodulin-dependent kinases (CaMK) and p65PAK [36–39]. In response to growth factor stimulation, the prolinedirected MAPKs family members, ERK1/2, target the proline-flanked STMN S25 residue predominantly and S38 to a lesser extent [40, 41]. A recent study demonstrated that Kinase interacting with stathmin (KIS) phosphorylates STMN S38 in migrating smooth muscle cells [42]. Collectively, these studies emphasizes that the involvement of particular kinases is dependent on cell context. STMN is also targeted by kinases activated by cellular stress stimuli, proinflammatory cytokines and differentiation agents. STMN S25 is phosphorylated

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Fig. 14.3 Kinases targeting STMN. List of kinases implicated in phosphorylating STMN in cells. Conserved STMN serine residue/s thought to be the preferred target sites for specific kinases are also indicated

by ERK1/2 in response to heat shock and chemical toxicity [43]. Isoforms of p38 MAPK (α, β, δ) target STMN S25 primarily in response to oxidative or osmotic stress [44, 45]. Interestingly, related stress-activated MAPK family members, JNK1/2, target STMN S38 preferentially in response to hyper-osmolarity [46]. In response to TNFα, STMN is phosphorylated on multiple serines (S16, S25 and S38) by unidentified kinases [47]. Depending on the specific stress stimulus, STMN phosphorylation may contribute to or prevent cell death [46–48]. This likely reflects complex functions of STMN activity in regulating apoptosis and the cell stress response. Furthermore, a recent study demonstrated targeting of STMN S16 site by an Epstein-Barr virus encoded kinase, BGLF4, may be involved in modulating changes in MT organisation associated with host-pathogen interactions [49]. Clearly, STMN links many signalling pathways to MT changes associated with various cellular processes in addition to cell cycle regulation.

14.3.2

Protein–Protein Interactions

STMN has been shown to bind proteins other than tubulin. In yeast or bacterial twohybrid assays, STMN interactions with molecular chaperones (e.g. BiP), coil-coil forming proteins (CC1, CC2) and a peptidyl-prolyl isomerase (PPIase) have been demonstrated [50–52]. The interaction between Hsc70 (a constitutively expressed Hsp70 protein) and STMN was further verified in mammalian cells by co-immunoprecipiation studies and was shown to be regulated by STMN phosphorylation and the ATP activation state of Hsc70 [50]. The biological significance of STMN/Hsc70 complex is currently unknown but one possibility is a role in the cell stress response given extensive STMN phosphorylation in response to cell stress [43, 46]. STMN is also co-immuno-precipitated with petidyl-prolyl isomerase-like 1 (PPIL1) when exogenously expressed in cells. Interestingly, it was phosphorylated STMN in particular which co-precipitated with PPIL1 [52]. PPIases regulate the activity, stability and localisation of a number of mitotically phosphorylated proteins and transcription factors (e.g. p53, c-Jun) through cis-trans isomerisation of phosphoserine-proline

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bonds [53]. The STMN S25 (-ILSPRS-) and S38 (-PLSPPK-) residues thus are potential target sites for PPIase activity. The possibility of PPIase regulation of STMN conformation and MT-destabilising activity awaits further characterisation. In other studies, protein interactions between STMN and a cell cycle regulatory protein, p27kip1 and a latent transcription factor, STAT3, have been reported [54, 55]. Interactions with p27kip1 or STAT3 negatively affect the MT-destabilising and tubulin-binding activities of STMN independently of phosphorylation status [54, 55]. Domain mapping studies indicated that STAT3 bound the C-terminal tubulin-binding domain of STMN which likely interferes with T2 S complex formation. In in vitro tubulin polymerisation assays, STAT3 could interfere with STMN pre-complexed with tubulin which suggested that the affinity of STAT3 interaction with STMN was sufficient to disrupt the T2 S complex [55]. The STMN binding site on p27kip1 was mapped to a conserved 28 amino acid C-terminal region of p27kip1 [54] while the STAT3 domain responsible for binding STMN has not been identified. To date, the function of the STMN-interacting proteins has been evaluated in the regulation of MT dynamics associated with cell migration [54–57]. A role for protein-binding partners and regulators of STMN in other cellular processes remains a possibility as both STAT3 and p27kip1 have well-characterised roles in cell cycle regulation and cell proliferation with STAT3 directly up-regulating cyclin D1 to induce cell transformation, while p27kip1 is a cyclin/cdk inhibitory protein [58, 59]. Thus, further characterisation of p27kip1 or STAT3 regulation of STMN activity in cell cycle regulation may be warranted.

14.3.3

Transcriptional Regulation

The elevated expression of STMN in human malignancies highlights the importance in understanding how the STMN gene is transcriptionally regulated during disease. STMN over-expression in cancer can be brought about by the constitutive activity of oncogenic factors or the loss or inactivation of proteins with tumoursuppressive functions. Several transcription factors have been found to regulate STMN expression. STMN was identified as a gene that was down-regulated following p53 induction, thus indicating a negative regulatory p53 role in the control of STMN expression [60, 61]. In support of this, p53 over-expression inhibited promoter activity of an STMN luciferase reported gene construct in fibrosarcoma cells [61]. STMN over-expression in carcinoma is also positively correlated with loss-of-function p53 mutations [62, 63]. However, this is not reported in all cases as high STMN levels have been shown in hepatocarcinoma with functional p53 [64]. Nevertheless, silencing STMN expression decreases proliferation and increases apoptosis in breast cancer cells with mutant p53 [62]. In addition, STMN over-expression can override p53-mediated G2 /M cell cycle arrest [61]. These studies indicate that enhanced STMN expression is mediated by p53 loss or mutation and may contribute to disease at least in a sub-set of human tumours.

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The mechanism of transcriptional control of STMN expression by p53 appears to be a complex one. p53 is not thought to directly repress STMN mRNA expression as the STMN gene promoter lacks a p53-binding site [65]. Instead, Egr1 and E2F1 transcription factors associated with p53 activity have been proposed as potential mechanisms that repress STMN expression [65, 66]. The STMN promoter contains both Egr1 and E2F regulatory response elements and both proteins have been shown to directly bind the STMN gene promoter in vivo [65, 66]. Egr1 and E2F1 occupy the STMN promoter in response to p53 induction and DNA damage, respectively, which then leads to STMN down-regulation [65, 66]. Other factors that have been shown to repress STMN expression include TGF-β-inducible early gene 1 (TIEG1) which induces growth inhibition in pancreatic cancer cells [67] and JNK-mediated c-Jun (which forms part of the AP-1 transcription factor complex) activation serves to repress basal STMN expression and fibroblast proliferation [68]. Transcriptional regulators that promote STMN expression include FoxM1b (a transcript variant of FoxM1), CCAAT-binding transcription factor, NF-Y, far upstream sequence element-binding protein-1 (FBP-1) and the enhancer of zeste homolog 2 (EZH2; [69–72]). Recent studies have demonstrated that FoxM1b, a proliferation-related transcription factor over-expressed in numerous malignancies, binds directly to the stathmin promoter to stimulate expression [70, 73]. The inhibition of stathmin expression in FoxM1b-expressing hepatocellular carcinoma cells inhibited lung metastasis in vivo [73] highlighting the contribution of FoxM1b in stathmin over-expression in numerous malignancies. NF-Y was shown to directly bind the STMN promoter in embryonic kidney cells while FBP-1 and EZH2 activity promotes STMN expression in non-small-cell lung cancer and liver cancer cells, respectively [69, 71, 72]. Furthermore, while E2F1 in the context of p53 activity in basal cells represses STMN expression, the E2F sites on the STMN promoter may also be required for high STMN expression in prostate carcinoma [74]. This suggests that E2F transcription factors can positively regulate STMN expression in some situations. In agreement with this, over-expression of E2F1 in embryonic fibroblasts increases STMN mRNA levels [75]. Similarly, c-Jun over-expression in fibroblasts induces an increase in STMN expression which is required for anchorageindependent growth, a characteristic of cellular transformation [76, 77]. Thus, the opposing effects of E2F1 and c-Jun transcription factors on STMN expression depend on the specific cellular context. Taken together, these studies indicate that a complex network of repressors/promoters is involved in regulating STMN over-expression in cancer cells.

14.3.4

Post-Transcriptional Regulation

Recent studies have revealed post-transcriptional regulation of STMN in cancer cells by microRNAs (miRNAs). The miRNAs are small (21–25 nucleotides) non-coding RNAs that bind to complementary sequences on target mRNA to induce transcript degradation/silencing and translational down-regulation [78]. Since their discovery,

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important regulatory functions for miRNAs have been characterised in the pathogenesis of human cancers [79]. It was recently revealed that STMN is a target of miR-223 in hepatocellular carcinoma and miR-9 in human neuronal progenitor cells [80, 81]. The down-regulation of miR-223 and a strong inverse correlation with STMN over-expression was identified in tumour biopsies from patients with hepatocellular carcinoma [81]. The miR-223-binding site on the 3′ UTR of the STMN promoter was identified by a luciferase reporter assay demonstrating direct regulation of STMN by miR-223 [81]. Furthermore, the re-expression of miR-223 reduced STMN expression and the proliferation of liver cancer cells implicating a significant role for miR-223-regulated STMN expression in hepatocellular carcinoma [81]. Similarly, STMN expression is inversely correlated with miR-9 expression in neuronal progenitor cells. Cell migration is increased as a result of miR-9 loss and this is in turn attenuated by STMN knockdown implicating a function for mir-9-regulated STMN expression in the motility of neuronal progenitor cells [80]. It is clear that the post-transcriptional control of STMN expression by miRNA serves important functions in pathogenesis of human cancer and it is likely that other miRNA families that target STMN remain unidentified. Furthermore, specific miRNAs are capable of regulating multiple targets simultaneously [78]. Thus, it is likely that other targets of miR-223 and miR-9 may act in concert with the control of STMN expression to regulate processes in cancer cells, although this has yet to be revealed experimentally.

14.4 14.4.1

STMN in Cancer Development and Progression Clinical Studies: Prognostic Value of Elevated STMN Expression

Many clinical studies have provided evidence linking STMN expression with disease progression across a wide range of cancer types (summarised in Table 14.1). Expression profiling on clinical samples has revealed that STMN mRNA and protein is commonly over-expressed and strongly associated with oncogenesis, disease severity and poor prognosis. Importantly, STMN expression may have independent prognostic value over current risk classification systems using standard clinical and pathological markers. In patients with medulloblastoma, STMN over-expression significantly correlated with disease progression and was predictive of tumour dissemination and survival in cases where other risk classification indicators (e.g. age, tumour size, histological grade) were not statistically significant [82]. Similarly, in hepatocellular carcinoma, STMN over-expression identified early-stage cancer patients with worse prognosis that were not distinguished by standard tumour staging and histological grading systems [83]. However, the prognostic value of STMN is not evident in all cancer types. For example, although a recent study found that STMN over-expression in colorectal cancer correlated with tumour stage [84], a separate study with a larger cohort reported that STMN expression independently predicted improved survival in colorectal cancer patients [85].

STMN expression

∗

[111] [112–114] [82, 115–117] [84] [85] [118] [119, 120] [121] [122] [123]

RT-PCR RT-PCR, WB, IH IH, 2D-PAGE IH IH WB, IH IH WB IH 2D-PAGE, WB, IH

[124]

[72, 106, 107] [108] [109, 110]

RT-PCR, WB, 2D-PAGE IH IH, RT-PCR

2D-PAGE

[4] [93] [94, 95] [96–98] [99] [100, 101] [86, 87, 102] [88, 103, 104] [63, 64, 83, 105]

Reference

2D-PAGE 2D-PAGE IH, WB WB, IH, 2D-PAGE RT-PCR IH WB, RT-PCR, IH IH RT-PCR, WB, 2D-PAGE, IH

Detection∗

IH immune-histochemistry, WB western blot, RT-PCR real time quantitative PCR, 2D-PAGE 2-dimensional gel electrophoresis

Increased expression in ALL and AML Increased expression and phosphorylation in ALL. Correlated with white blood cell count Increased expression in malignant samples Increased expression in malignant or advanced carcinoma Increased expression in non-serous histotype and correlated with chemotherapy resistance Increased expression correlating with advanced prostatic carcinoma Increased expression in sub-group of primary tumours Expression predictive of poor prognosis in breast cancer patients Increased expression in hepatocellular carcinoma correlated with tumour progression, recurrence and poor prognosis Lung Increased expression and phosphorylation in non-small-cell lung carcinoma Gastric Expression correlated with tumour metastasis Bladder Increased expression in upper urinary tract urothelial carcinoma and in early recurrent non-muscle- invasive urothelial carcinoma Kidney Increased expression in nephroblastoma and correlated to advanced tumour stage Neuroendocrine Increased expression in pheochromocytomas, paragangliomas and neuroblastomas. Brain and CNS Increased expression in medulloblastoma, anaplastic gliomas and primitive neuroectodermal tumours Colorectal Increased expression correlated with tumour stage and predictive of poor prognosis Colorectal Obesity associated with high mortality in STMN-positive patients Cervical Increased expression in primary tumours and correlated with advanced tumour stage and metastasis Endometrial High expression correlated with poor prognosis and lymph node metastasis Mesothelial Increased expression in malignant pleural mesothelioma Head and Neck Increased expression in primary oral squamous-cell carcinoma. Predictive of poor recurrence-free survival Head and Neck Increased expression in nasopharyngeal carcinoma correlated with tumour differentiation, metastasis and poor prognosis Salivary Increased expression in adenoid cystic carcinoma of the salivary gland

Acute leukemia Acute leukemia Lymphomas Ovarian Ovarian Prostate Breast Breast Hepatic

Cancer type

Table 14.1 Clinical studies of STMN over-expression in different cancer types

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Fig. 14.4 How does STMN contribute to cancer? A role for STMN activity in regulating cellular processes such as proliferation, differentiation, cell survival, mitosis and migration/invasion could contribute to tumourogenesis and cancer progression. Global changes in STMN phosphorylation status or expression levels could impact on cellular proliferation and differentiation. In other cases, such as in cell migration, an STMN activity gradient is involved in directing cellular function

In other cancer types (e.g. breast cancer) STMN over-expression correlated with standard disease parameters to predict poor prognosis and patient survival [86, 87]. STMN over-expression may also reveal tumour sub-types with specific oncogenic pathway signatures. For example, STMN expression is regulated by PTEN loss and PI3K activation in breast cancer cells and was found to be a surrogate marker of the PTEN/PI3K signatures in tumours [88]. Thus, STMN may represent a valuable marker to stratify patients with PTEN/PI3K pathway-specific tumours for subsequent targeted therapy. In general, clinical studies to date highlight the potential value of STMN as an independent prognostic factor and diagnostic biomarker in multiple cancers. Although the precise contribution of STMN over-expression to carcinogenesis is unresolved, it remains likely that multiple mechanisms are involved (Fig. 14.4).

14.4.2

Cell Differentiation and Proliferation: Implication of STMN Levels

The levels of STMN in many cell types, with the exception of neuronal cells, are closely associated with differentiation stage. STMN activity and expression coincide with restructuring of the cytoskeleton during cell differentiation and can be modulated by a range of different factors (e.g. nerve growth factor [NGF] and phorbol esters; [40, 89]). In some cell types, a transient increase in STMN expression is observed in the early stages of cell differentiation [40, 90] after which expression declines in conjunction with growth arrest [91]. Furthermore, STMN depletion represses expression of early-stage markers of differentiation [92]. Interestingly, in

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post-mitotic neuronal cells, STMN expression remains high indicating a distinct functional role for STMN in these cells [90]. STMN over-expression strongly correlates with poorly differentiated tumours [100, 106, 108]. Thus, high STMN levels in cancer cells directly correlate with cell proliferation (reviewed in [125] as seen in lung adenocarcinomas and breast carcinomas where STMN over-expression was found to correlate with poorly differentiated and highly proliferative tumours [87, 106]). In lung cancer and acute leukemia, the phosphorylated STMN sub-population was elevated in line with higher protein expression [93, 106]. More significantly, phosphorylated STMN as a percentage of total STMN protein was also increased [93, 106]. Given the requirement that STMN is hyper-phosphorylated during G2 /M transition, this indicates a greater number of mitotic cells and is consistent with STMN associations with tumours with a high mitotic index (e.g. [87]). In human cervical cancer and human osteosarcoma cell lines, STMN siRNA expression triggered growth inhibition, cell cycle arrest and an increase in apoptosis [126]. Similarly, virally delivered STMN mRNA-targeting ribozymes, leading to down-regulated STMN expression, led to inhibition of proliferation and clonogenecity and increased apoptosis in prostate cancer cells [127]. A recent study also demonstrated that STMN inhibition in gastric cancer-derived cells had anti-tumour effects in a xenograft model in nude mice [108]. These studies provide evidence supporting the beneficial effects of targeting STMN in the treatment of cancer.

14.4.3 Aneugenic Activity: STMN Regulation of Chromosome Stability Somatic chromosomal aneuploidy (abnormal chromosome number) is a common feature in cancer and is indicative of increased chromosomal and genetic instability that is understood to drive tumour progression [128]. The discovery of a novel somatic missense mutation of the STMN gene [STMNQ18E] has provided insight into how STMN can promote tumourgenesis [129]. Tumour xenograft studies indicated that the Q18E mutation potentiated STMN-mediated cellular transformation and tumour development in immuno-compromised mice [129]. The oncogenic properties of mutant STMN are likely linked to MT-destabilising activity as the Q18E mutation was found to antagonise the inactivating effects of multi-site phosphorylation and/or prevent the efficient phosphorylation of STMN under growth conditions [129, 130]. Expression of STMN Q18E at levels similar to endogenous STMN was subsequently found to induce aneuploidy in cultured myelogenous leukemic cells and immortalised T lymphocytes [28, 131]. The precise STMN-regulated mechanism requires further characterisation to evaluate its temporal association with events occurring subsequent to metaphase to anaphase transition that suggest a relationship with chromosomal attachment and/or segregation but being independent of mitotic spindle assembly [131].

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Although the clinical significance of STMN Q18E mutation remains unclear, it remains identified in a single tumour from a single individual, the altered biological properties of this mutant has revealed a novel capacity for excessive STMN activity to promote aneuploidy and chromosomal instability in cancer cells. Subsequent to the discovery of the STMN Q18E mutant, it was found that strong over-expression of normal STMN in cultured cells, which also increases STMN activity, was similarly aneugenic [130]. When highly expressed in a human leukemic cell line, wild-type STMN was found to similarly increase the incidence of micronuclei and frequency of polyploidy [130]. This effect is augmented by the Q18E mutation. A recent clinical study also found a significant association between STMN over-expression and polyploidy in hepatocellular carcinoma [83]. However, further detailed investigations will be required to assess the significance of STMN-mediated aneugenic activity in tumour development and disease progression in vivo.

14.4.4

Migration and Metastasis: Role for STMN Regulation of Interphase MTs

It has been widely reported that STMN over-expression correlates with highly metastatic disease in various malignancies (e.g. [82, 83, 108]). A key step in the metastatic process involves the migration of cells away from the primary tumour site. STMN is implicated in the migration of many different cell types including Drosophila germ cells, embryonic fibroblasts (MEFs), human endothelial cells and trophoblasts [54, 55, 132–134]. Furthermore, STMN may prove a valuable target to treat metastatic disease as silencing of STMN expression reduced cell migration in a range of cancer cell lines [54, 108, 135], while over-expression or increased activity significantly enhanced sarcoma cell migration [136]. However, the exact mechanisms underlying STMN-mediated cell migration are complex. STMN is a key regulator of the interphase microtubule network which is critical for cell migration. Interphase microtubules transport membrane vesicles and actinregulating proteins to different sub-cellular compartments required for cytoskeletal restructuring during cell migration. In interphase cells, STMN activity is spatially and temporally regulated down-stream of various signal transduction events [137]. During migration, inhibition of STMN activity, as judged by its decreased binding to tubulin, is restricted to the leading edge of motile cells, as determined by a fluorescence resonance energy transfer (FRET)-based analyses in migrating Xenopus cells [138]. Using a dual-fluorescently labeled (YPF/CFP) STMN biosensor which displays decreased FRET upon tubulin binding, Niethammer and colleagues demonstrated spatial heterogeneity of STMN: tubulin interactions in live migrating cells which is generated in part by localised differences in tubulin concentration and in part from spatially distinct STMN phosphorylation [138]. In support of this, only a fraction of STMN is phosphorylated during growth-factor-induced cell migration [139]. Thus, differential STMN phosphorylation gradients may mediate local microtubule dynamics and promote ‘pioneer’ microtubule growth at the leading edge of migrating cells [138, 139].

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The shape and structure of cells moving on 2-dimensional surfaces differ greatly from that in three dimensions or in vivo, which begs the question as to how STMN may be contributing to metastasis in vivo. STMN is more abundant in vascular invasive HCC, endometrial carcinomas, gastric cancer and is localised at the invasive front of NSCLC [72, 83, 108, 119]. This suggests that STMN over-expression may promote cell invasion. Indeed, Belletti et al. demonstrated a functional role for STMN in regulating 3-dimensional motility and invasion of sarcoma cells [136]. Specifically, STMN over-expression and expression of the Q18E mutation enhanced sarcoma cell invasion in vitro and metastatic potential in vivo [136]. Further analysis demonstrated that expression of the Q18E mutation reduced extracellular matrix (ECM) contact-mediated STMN phosphorylation, microtubule stabilisation and induced a rounded-shape morphology, stimulating amoeboid-like invasion of sarcoma cells in vitro [136]. Thus, high STMN expression and/or activity in cancer may regulate microtubule-mediated changes in cell morphology upon contact with the ECM which in turn aids cell invasion.

14.5 14.5.1

STMN and Chemotherapy STMN Expression and Drug Resistance

The inherent or acquired resistance to chemotherapy represents a significant obstacle to the successful clinical treatment of cancer. It is clear that multiple mechanisms underlie the development of drug resistance, acting in concert to reduce efficacy. Emerging evidence points to a role for over-expressed STMN in mediating cancer cell resistance to chemotherapy, particularly to tubulin-targeting agents (e.g. taxanes and vinca alkaloids) and various DNA-damaging agents including platinumbased alkylating compounds (e.g. cisplatin; [64, 140–142]). For example, STMN is over-expressed in paclitaxel-resistant ovarian cancer cells [141] and NSCLC [143]. Elevated STMN expression was also observed in human neuroblastoma cells selected in the presence of vincristine [144]. In a clinical study on patients with ovarian cancer, STMN mRNA expression was found to be an independent predictor of poor clinical outcome in patients that had received paclitaxel treatment specifically suggesting that high STMN expression may impact on the efficacy of treatments involving taxanes [99]. However, STMN over-expression is not associated with chemoresistance in all cancers and curiously, in at least one study using a xenograft model of acute lymphoblastic leukemia, acquired vincristine resistance was associated with downregulation of various STMN phosphoisoforms [145]. Similarly, STMN has been shown to increase the affinity of vinblastine for tubulin in vitro, suggesting STMN may enhance sensitivity to microtubule-destabilising agents [146]. In some cell types this may be the case as STMN over-expression enhances lung cancer cell sensitivity to the microtubule-destabilising agents, vincristine and vindesine [147]. Attenuating the levels of over-expressed STMN in cancer cells may also alter responses to tubulin-targeting agents. For example, forced over-expression of

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STMN in breast cancer cells decreased sensitivity to paclitaxel and vinblastine [148]. Conversely, STMN down-regulation with small-interfering RNA (siRNA) increased sensitivity to paclitaxel and vinblastine [140, 149]. These studies indicate that modulating STMN levels is sufficient to influence chemoresistance in some cancer cell types. Interestingly, STMN may also influence resistance to chemotherapeutic agents that do not target tubulin. STMN mRNA expression is increased more than tenfold in squamous cell carcinoma cell lines resistant to the platinum-based drug cisplatin [142]. Furthermore, low STMN expression in malignant gliomas with a loss of heterozygosity of chromosome 1p (which contains the STMN gene) was shown to be responsible for the chemosensitivity of this glioma sub-type to nitrosourea treatment [117]. This study went on to show that siRNA knockdown of STMN in cells heterozygous for chromosome 1p was sufficient to confer sensitivity to nitrosourea, implicating a role for STMN in chemo-resistant malignant gliomas [117]. STMN inhibition also synergistically enhanced the anti-tumour effects of the DNAdamaging agent, etoposide [150].Yet, STMN expression does not affect breast cancer cell sensitivity to a related DNA-targeting agent, doxorubicin [150]. Although there does not appear to be a clear relationship with distinct drug classes, these studies highlight links between over-expressed STMN and tumour resistance to a diverse range of chemotherapeutic agents. It was also demonstrated that a DNA-alkylating agent 1-(2-chloroethyl)-3-cyclohexyl-l-nitrosourea (CCNU), commonly used for the treatment of brain tumours, can covalently bind and promote carbamylation of lysine residues within STMN subsequently inhibiting its microtubule-depolymerising activity [117]. Therefore, agents that can directly inhibit STMN activity may play a key role in treating drug-refractory malignancies.

14.5.2

Combinatorial Therapy

The targeting of STMN in combination with chemotherapy may also improve therapeutic outcome. Significantly, the capacity of STMN inhibition in combination with several chemotherapeutic agents to reduce tumour growth appears to be synergistic and not merely cumulative. STMN inhibition synergised with taxol treatment to inhibit the proliferation and migration of endothelial cells [132] and reduced the clonogenic potential of leukemic cells [149]. STMN inhibition could also synergise with non-MT-targeting chemotherapeutic agents. The combination of STMNtargeting ribozymes with DNA-damaging agent, etoposide, was shown to reduce the malignancy of prostate cancer cells [150]. Furthermore, in combination with STMN down-regulation, the use of taxol and etoposide at sub-therapeutic, normally noninhibitory doses induced profound growth inhibition and reduced clonogenicity in prostate cancer cells [150]. These findings are significant as they suggest the potential for improved drug efficacy when coupled with STMN targeting. However, the growth inhibitory effects of STMN inhibition were merely additive when used in combination with other therapeutic agents such as Adriamycin or 5-fluorouracil [150]. The reasons behind this selectivity are unclear while our lack of understanding of the mechanisms governing the synergistic actions of STMN inhibition is problematic.

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Mechanism of STMN-Mediated Chemoresistance

The precise mechanism involved in STMN-mediated chemoresistance is as yet undetermined. The synergistic anti-cancer effects of targeting STMN and taxol treatment may be related to their shared capacity in modulating MT assembly although direct evidence of this is currently lacking. Another explanation may involve effects of combinatorial therapy on cell cycle progression. Indeed, Iancu et al. proposed that STMN inhibition combined with anti-mitotic drug treatment is likely to increase cell sensitivity due to disruption of distinct steps in the same (mitotic) pathway [149]. As such, in many instances whereby STMN inhibition increases sensitivity to paclitaxel, STMN inhibition alone disrupted mitotic progression [64, 149, 151]. Furthermore, ribozyme-mediated STMN down-regulation in combination with chemotherapeutic drug treatment substantially increased mitotic arrest at G2 /M in several cancer cell types [127, 150]. However, the impact of STMN inhibition on the cell cycle remains controversial as others have not reported mitotic arrest from STMN down-regulation in similar cancer cells [130]. It appears reasonable to propose that the enhanced expression of STMN, an MTdestabilising factor, could directly antagonie the effects of cancer drugs, such as taxol, that specifically bind and stabilise tubulin polymers to induce mitotic block and subsequently cell death [148]. Consistent with this notion is evidence that MT dynamics are elevated and associated with taxol resistance in lung cancer cells [152]. However, it is more difficult to rationalise how the over-expression of active STMN could contribute to resistance against drugs that inhibit the assembly of MTs (i.e. vinca alkaloids). Similarly, it is unclear how STMN-mediated effects on MT dynamics would influence the efficacy of drugs that do not target tubulin (e.g. nitrosurea, etoposide). That STMN inhibition could synergise with chemotherapeutic agents with diverse modes of action [150] to inhibit tumour growth suggests a more general mechanism behind STMN-mediated chemoresistance. Another potential mechanism links stathmin to cell survival and apoptosis. STMN knock-out mice display elevated apoptosis and delayed recovery from renal injury [153], and genetic silencing of STMN in cancer cell lines is shown to enhance apoptotic cell death [62, 126, 154]. Although the exact mechanisms linking STMN to apoptotic pathways are unclear, it has been speculated that STMN-mediated regulation of microtubule dynamics may alter the cellular activity and translocation of apoptotic B-cell lymphoma (BCL) family members [155]. Interestingly, recent studies have shown that susceptibility to cell death pathway activation is a primary determinant of cancer cell sensitivity to anti-mitotic chemotherapeutic agents [156]. Specifically, apoptotic cell death is initiated in response to anti-mitotic drug treatment with the culmination of multiple factors resulting in caspase activation [156]. Thus, tumours with over-expressed STMN may be more resistant to the initiation of apoptosis leading to a generally improved tolerance to anti-mitotic drugs. Alternatively, STMN over-expression prolongs G2 phase of cell cycle in breast cancer cells and this delay in mitotic entry has been proposed to be involved in STMN-mediated resistance to vinblastine [140, 148]. This delay in mitotic entry may also confer resistance to other anti-mitotic agents.

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Concluding Remarks

Research over the past 20 years has revealed complex regulation of STMN activity and diverse STMN contributions towards cellular functions under physiological and pathological conditions. However, the STMN link in most cancer types is predominantly correlative, while the therapeutic benefits of targeting STMN are primarily supported, in principle, by in vitro cellular studies in most cases. Although this highlights the value of elevated STMN levels as a diagnostic marker in cancer, the precise in vivo contributions of STMN towards tumour formation and progression remain to be fully determined. Nevertheless, studies to date highlight the potential of STMN as a target to reduce disease. Thus, the development of strategies to inhibit STMN in vivo represents an important goal. A recent study has demonstrated in vivo safety and efficacy of an STMN shRNA in immune-compromised rodents [157]. In addition to targeting STMN mRNA, with ribozymes and siRNA, the discovery that a DNAalkylating agent could covalently bind, modify and reduce STMN activity point to the possibilities of chemical inhibitors in directly targeting the STMN protein in malignancies [117]. While co-crystallisation of SLD with tubulin has facilitated studies of drug-tubulin interactions (e.g. [158]), recent studies also highlight the complex interactions between STMN activity and chemotherapeutic agents in cancer cells. In particular, the improved effectiveness of sub-therapeutic doses of anti-mitotics, when used in combination with STMN inhibition, suggests that combinatorial therapy may be useful in treating drug-refractory cancers, improving drug efficacy and the management of toxic side-effects from chemotherapy. Clearly, before this can be achieved, further investigation into the mechanism underlying STMN-mediated chemotherapy resistance is required. Thus, despite intensive investigation significant challenges remain in evaluating contributions of STMN in cancer. Acknowledgments We would like to thank Marie Bogoyevitch for helpful suggestions in editing. Research by the authors on STMN function and regulation has been supported by the Australian National Health and Medical Research Council (628335). D.C.H. Ng is also supported by the MDHS CR Roper Fellowship and Frances Byrne by an Anthony Rothe Postgraduate Scholarship.

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Chapter 15

The Biology and Pathobiology of Tau Protein Garth F. Hall

Abstract The cytoskeleton-associated protein tau has a remarkably varied repertory of cellular functions in neurons in both health and disease. Although tau is classically thought of as a neuronal phosphoprotein that stabilizes axonal microtubules (MTs), it interacts with diverse signaling pathways via proteins in the cortical cytoskeleton as well and plays important roles in several aspects of axonal development and function. Abnormalities in non-MT-associated functions of tau are increasingly thought to be involved in the pathogenesis of neurodegenerative disease in humans, which is the focus of this monograph. I will first summarize normal tau interactions with other proteins and cellular functions as we currently understand them in the context of tau structure and expression patterns and discuss how these are altered in the context of neurodegenerative disease at the molecular, cellular, and organismal levels. Particular attention is given to recent studies of the cellular and intercellular aspects of tau misprocessing and their implications for the pathogenesis of human neurodegenerative disease.

15.1

Introduction

Although tau was originally characterized as an axonal microtubule-associated protein (MAP; [1–3]) that regulates MT dynamics and spacing [4, 5], it is now clear that it participates in diverse aspects of neuronal biology and pathobiology, some of which are only tangentially related to MT stability. Tau modulates the structural stability of both MTs and the actin-rich cortical cytoskeleton in response to diverse outside signals via dynamic regulation of its phosphorylation state in both developing and fully differentiated neurons, and plays well-defined roles in axonal outgrowth, the development of axonal identity, and in axon myelination. Specific abnormalities associated with certain of these functions combined with the well-studied tendency of tau to form fibrillar aggregates now appear to play central roles in the pathobiology of Alzheimer’s Disease (AD) and the related conditions called “tauopathies” as well. G. F. Hall () Center for Cellular Neuroscience and Neurodegeneration Research, Department of Biological Sciences, University of Massachusetts Lowell, 198 Riverside St. Lowell, 01854 Lowell, MA, USA e-mail: [email protected]

M. Kavallaris (ed.), Cytoskeleton and Human Disease, DOI 10.1007/978-1-61779-788-0_15, © Springer Science+Business Media, LLC 2012

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Tau is a member of a family of MAPs that are characterized by imperfect multiple tandem repeat (microtubule-binding repeat or MTBR) sequences (31 residues each) via which they bind to and stabilize MTs [6, 7] via a phosphorylation-regulated mechanism [8, 9]. This family includes MAP2 and MAP4 in mammals as well as tau, and has homologues among protostome phyla as well as chordate and nonchordate deuterostomes [10–12]. MAP2 and tau are most strongly expressed in neurons and glia, where they play functionally differentiated roles in axons and dendrites, respectively [13, 14]. While overall tau sequence homology among mammals is relatively high, most of the tau coding sequence outside of the C-terminus and MTBRs is poorly conserved among nonmammalian vertebrates and nonvertebrate phyla [15, 16]. This parallels our level of understanding of tau cellular function; we have always understood the mechanisms associated with the tau C-terminal MTBR domain much better than we have those of the N-terminal domain. This has begun to change in recent years owing to the increased interest in the role played by tau in the pathogenesis of AD, where it is now clear that both the N- and C-terminal domains of tau play critical roles [17, 18].

15.2

Normal Tau Biology

Tau expression, distribution, and alternative splicing Tau expression in humans occurs from a single gene located on chromosome 17 and generates three distinct mRNA species and at least ten distinct protein species in a variety of cell types, with by far the highest expression occurring in neurons. The identity of the tau protein that is eventually produced in a given tissue is tightly regulated by multiple mechanisms that are thoroughly described in the excellent review by Andreadis [19]. The major tau species expressed in humans consist of the six alternatively spliced isoforms (ranging from 352 to 441 residues in length) expressed in the CNS, plus the larger protein (“big tau”) expressed in the PNS and also to some extent in muscle. Additional tau species include 2C-terminal truncated species that lack the MTBR (66p and 66d) produced by inclusion of an alternative reading frame in exon 6 [20] and a nuclear tau species generated from a separate mRNA [21, 22]. Low-level expression of various other splice variants has been reported in the CNS as well as in other tissues [19]. The six splice variants of CNS tau are produced by the inclusion of exon 2, exons 2/3, and/or exon 10 in the tau coding sequence, combined with the constitutive exclusion of Exon 4a (Fig. 15.1).

15.2.1

Tau Structure and Function

Tau is a highly soluble, cytosolic protein that was first identified and characterized in the mid 1970s [1–3] in the context of its association with MTs to which it binds in a phosphorylation-dependent manner [8] via 3 or 4 conserved MT-binding tandem

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Fig. 15.1 Schematic of the longest tau isoform (441 amino acid residues) expressed in the human CNS showing the functional anatomy, major splice variants, and key phosphorylation sites. The binding sites of “normal” alternatives to MT binding (actin, various chaperones, and dynactin—see text) are approximately shown. Alternatively, spliced exons are present in the amino (N-) terminal “projection” domain (2,3) and carboxyl (C-) terminal domain (10), which binds to MTs via basic K/R-rich repeat regions (R). Regulation of tau-MT binding by phosphorylation of the flanking domains is crudely analogous to a “dimmer switch” in which phosphorylation decreases the net charge difference favoring binding in a graded manner based on tau-PO4 stoichiometry [44,100, 125]. By contrast, phosphorylation of residues within the MTBRs (green arrows) block tau-MT association almost completely [56]

repeat motifs (MTBRs; [6, 7]). Cellular studies quickly established that tau expression is largely neuronal and that the protein is selectively accumulated in axons [12], where it is necessary for the initial establishment of both axonal identity [23] and for overall neuronal polarity [24, 25]. More recent studies suggest that tau may be involved in aspects of neuronal plasticity [26] and arousal [27]. Overall, tau protein exhibits well-defined structural motifs which have been assigned moderately wellunderstood functions. The C-terminal domain mediates MT binding via the MTBR which are regulated by phosphorylation of the P- and S-rich flanking domains, while the N-terminal projection domain regulates MT spacing and interactions with perimembranous elements. This is schematized in Fig. 15.1 and discussed in more detail below. MT binding and stabilization via the tau C-terminal repeat domain Tau is generally considered to be an “intrinsically disordered protein” due to its relatively large number of irregularly distributed proline residues, which gives it minimal secondary structure and a “random coil” conformation when free in the cytosol [28, 29]. The MTBRs are the most internally ordered part of tau, each repeat motif having patterns of polar and nonpolar residues and an overall slightly positive charge [28, 30]. Tau appears to bind across the “grain” of tubulin protofilaments the MT at multiple points on the outside of the MT [31, 32] and thus presumably reduces the intrinsic dynamic instability of MTs by preventing the “catastrophes” caused by GTP to GDP conversion at the terminal dimer [4]. Photobleaching experiments suggest that tau MT binding is highly dynamic [33], although some studies have suggested that tau may become stably integrated into MT structure under certain conditions [34]. The strength of tau-MT binding is proportional to the number of MTBRs present, making isoforms with 4 MTBRs (4R isoforms) stronger MT “stabilizers” than 3R isoforms [35, 36].

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The physical properties of the MT are also altered by tau binding, becoming “stiffer” with the presence of even a few tau molecules [37], as does the tau protein itself [38]. While the key posttranslational modifier of tau-MT binding is clearly phosphorylation at serine and threonine residues to regulate the dynamics of tau-MT binding near and within the MTBR, tau-MT association can also be modulated by environmental factors, such as hypothermia, anesthesia, and glucocorticoid-mediated stress, all of which involve changes in the phosphorylation state of tau [26, 39, 40].

15.2.2

Tau Phosphorylation

Regulation of tau-MT binding by serine/threonine phosphorylation in and around the MTBRs The identification of abnormally phosphorylated, filamentous aggregated tau protein as the primary component of neurofibrillary tangles (NFTs) in AD [42, 43] has directed an immense amount of attention to the mechanism by which phosphorylation of serine and threonine residues in the regions flanking the tau MTBR regulates tau-tubulin binding, making this the best understood aspect of tau biology, and the subject of a number of detailed reviews [40, 44, 45]. The overall mechanism can be summarized as consisting of two elements: (a) the proline-directed phosphorylation of one or more of the 12–14 serine and threonine residues in the P- and S-rich regions flanking the MTBR on the N and C sides, respectively, by a variety of serine/threonine kinases (Fig. 15.1), resulting in a graded attenuation of tau-MT affinity and (b) the phosphorylation of 2–4 KXGS motifs within the MTBRs themselves, primarily by MARK kinases, resulting in the near abolition of tau binding to MTs [46]. However, this relatively neat and simple summary becomes much more complicated when examined in detail, which has hindered a generally applicable synthesis of tau functions. Most studies have been directed at testing the ability of specific kinases to phosphorylate tau and tau-derived peptide under defined conditions that may vary somewhat between studies. A wide array of serine/threonine kinases with overlapping substrate and context specificities are now known to phosphorylate tau efficiently under physiological conditions (Fig. 15.2). Some of these kinases, most notably glycogen synthase 3 beta, (GSK3β, sometimes called tau kinase 1 [47–49]), cyclin-derived kinase 5 (cdk5 [50, 51]), the MAP kinase family [52], and the MARK kinases [53] have received more attention because they: (1) are involved in tau hyperphosphorylation/aggregation [54] and (2) participate in other cellular functions that become abnormal in tauopathies, such as neuronal polarity axonal identity and function [55–58], tau isoform splicing [59], mitotic status [60–62], and/or (3) interact with other disease-associated proteins, such as APP [63, 64], presenilin1 [65], or alpha synuclein [66, 67]. Overall, the complex pattern of tau-tubulin affinity regulation by multiple kinases and signal transduction pathways ensures that tau binding to MTs is highly versatile and responsive to signals, especially during complex developmental events. However, it now appears that this arrangement also permits these kinases to become involved in tauopathy pathogenesis via mechanisms that are not yet clear.

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Fig. 15.2 Regulation of tau-MT binding by phosphorylation of S and T residues in and near the MTBRs. The site specificity of serine/threonine phosphorylation by four major tau kinases (protein kinase A [PKA], Calcium-modulated kinase II [CamKII], Glycogen Synthetase Kinase 3 beta [GSK3β], and cyclin-dependent kinase 5 [cdk5]) in aggregated (NFT tau) and normal tau as determined by immunoblots with P-specific mAbs after incubation with each kinase individually and in combination with others is shown to illustrate the range and complexity of NFT-tau sites were identified by first treating NFT-tau preparations with protein phosphatase 2A (PP2A), a broadly acting tau phosphatase. The list is derived from references [41, 45, 100, 128] and is not exhaustive—a variety of other serine/threonine (S/T) kinases, including Casein Kinase 1 and 2, SAD70S6 kinase, and members of the MAP kinase family (ERK1, ERK2) are known to phosphorylate tau and are not shown

Phosphorylation of the projection domain The highly acidic character of the tau projection domain appears to be modulated by phosphorylation of S and T residues by some of the same kinases (PKA, casein kinase I, casein kinase II, CAM Kinase II) responsible for Tau-MT binding [42, 68]. Phosphorylation appears to extend the projection domain away from the MT by exacerbating charge-charge repulsion, possibly modulating MT spacing [5] in a manner analogous to the regulation of neurofilament (NF) spacing at kinesin spindle protein (KSP) repeat motifs [69]. The activity of Casein Kinase II selectively phosphorylates sites within the E2 and E3 amino terminal inserts [68], and may thereby regulate the increased N-terminal extension permitted by these inserts. Tyrosine phosphorylation of tau Human CNS tau has five tyrosine residues, four of which (at 18, 29, 197, and 394) are phosphorylated by one of several nonreceptor tyrosine kinases of the Srk family (i.e., src, fyn syk, abl). The most studied of these is fyn, which binds to tau at 224–236 and phosphorylates tau at tyrosine 18 and is essential for key aspects of both normal and abnormal tau biology [17, 70–72], Abl kinase, which phosphorylates primarily Y394 of tau [73, 74] may mediate signals relevant to growth cone guidance [75]. Both fyn and abl phosphorylation of tau are increased in AD. It appears that both tyrosine and serine/threonine kinase sites have overlapping site specificities [74], consistent with observations that tau interacts with multiple signal transduction pathways and helps to coordinate more complex cellular functions, such as axonal development, myelination and the maintenance of axonal identity [73, 76]. Tau dephosphorylation Phosphorylated tau is the substrate for a broad range of serine/threonine and tyrosine phosphatases in vitro [43–46, 77], and the dynamic nature of tau phosphorylation and dephosphorylation strongly suggests that this is true in vivo as well [32, 33]. PP1, PP2A, PP2B, PP5, and tissue-nonspecific phosphatase (TNAP) have all been identified as tau phosphatases [78–81], although it is

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unclear to what extent and under what circumstances each of these dephosphorylates tau in vivo. PP2A and TNAP have been identified as playing specific roles related to tau, with a reduction of PP2A activity reportedly playing a significant role in intracellular tau hyperphosphorylation in AD [82], while TNAP activity potentiates the neurotoxicity of extracellular tau [83].

15.2.3

Other Site-Specific Protein–Protein Interactions of Tau

Tau interactions with the actin cytoskeleton Tau normally interacts with a variety of proteins other than tubulin which can be divided into those that modulate tau-MT binding (e.g., the serine threonine tau kinases discussed above) and those which are incompatible MT binding and/or occur only in the absence of tau-MT association. The extensive interaction of tau with both serine/threonine and tyrosine kinases that are known to play major roles in signal transduction is consistent with both modulatory and alternative roles of tau with respect to MT binding and stabilization, with the former modulating MT-specific functions of tau and the latter regulating an alternative set of tau functions associated with the actin cytoskeleton and the plasma membrane. Tau–actin interaction has been reported to require the presence of the MTBR domain of tau [84, 85], although it is unclear if the interaction between tau and actin is direct [86]. However, the starkly inverse effects of phosphorylation and/or pseudophosphorylation at KGXS sites in the tau MTBRs (discussed further below) suggest that actin association is largely an alternative function of tau [87]. Actin association and/or perimembranous relocalization of tau away from MTs is typically associated with the phosphorylation of tau atY18 by fyn kinase [70–72, 88], and has been reported to modulate the activation of fyn by external growth factors resulting in changes in the actin cytoskeleton [72], and the modulation of growth cone motility and guidance [76]. Tauopathy mutants-induced abnormalities in taumembrane interactions accompany neurodegenerative changes in fly [89, 90] and lamprey tauopathy models [91] which may involve also cooperative conformational changes between the tau N-terminus and sites in the MTBR domain to produce aggregation-associated epitopes [92] and/or dysfunctional interactions involving other actin-associated proteins. Interestingly, tau-dynactin interaction appears to be mediated via binding at residue 5 on the tau N-terminus, which is the site of the only known N-terminal tauopathy mutations [93]. Tau interactions with chaperone proteins The susceptibility of tau to misfolding when it is dissociated from MTs is consistent with reports of tau interactions with a number of chaperone proteins, including HSP90, HSP70/HSPc70, HSP27, alpha crystallin, and the Prolyl isomerase Pin-1, [94] as well as to disease-associated proteins that probably have chaperone-like functions (i.e., alpha synuclein and prion protein). Of these, HSP70 and HSPc70 are the best characterized, with the latter binding to tau via the same hydrophobic motifs in each MTBR (VQIV) which mediate tau-tau interactions, and that also lead to tau aggregation in tauopathies [95]. Chaperone binding is inversely proportional with tau aggregation and appears specific

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for non-MT-associated tau, since tau-HSP interaction appears to preclude MT binding as well as tau binding via the MTBR [94], but need not prevent indirect tau-MT association via the MT-binding site on HSP70 [96, 97]. The prolyl isomerase Pin 1 appears to act as a chaperone since it specifically binds to phosphorylated tau at the proline-directed Thr231 site and induces a cis to trans conformational change that restores tau-MT-binding capability [98]. However, the demonstration that Pin1 overexpression exacerbates the tauopathic effects of WT tau while protecting against those of P301L mutant tau suggests a more complex relationship between these proteins. Similarly, the binding of the chaperone-like protein alpha synuclein to tau near the MTBR, has received considerable attention over the past few years [66] due to its association of non-MT-bound tau [99] and obvious potential relevance to both tauopathy and Parkinson’s Disease/Lewy Body dementia, where alpha synuclein misfolding plays a central role [100]. However, like Pin-1, alpha synuclein-tau association is inhibited by the presence of the P301L tauopathy mutation, which suggests a possible role for these interactions in normal tau biology as well [101]. Other tau-interacting chaperone proteins that are associated with tauopathy have been shown to be important in mediating the as yet poorly understood role of tau in neuronal plasticity [102], illustrating the complexity of the relationship that tau has with both the regulation and misregulation of protein folding in neurons (discussed further below).

15.2.4

Cellular Functions of Tau

The regulation of tau isoform expression, intracellular distribution, and phosphorylation during embryogenesis suggests that tau has a variety of developmental functions involving the regulation and coordination of MT and actin dynamics as well as its established role in stabilizing the morphology and function of differentiated neurons. Nuclear tau functions The shortest tau isoform (3R0N tau or “fetal tau”) localizes to the nucleus as well as the axon during development [21], where it has been reported to bind to single- and double-stranded DNA [103], nucleoli organizers [22], and to be phosphorylated by cell cycle-regulatory kinases (cdc2, cdk5) at proline-directed sites when expressed in mitotic cells [104]. While the role of nuclear tau is still poorly understood, it is of clinical interest because of the association that abortive reentry of neurons into the cell cycle has in AD [105, 106] as well as in certain malignancies [107]. Specification of neuronal polarity (axonal identity) One of first characteristics of tau to be established was its axonal localization, where it was used together with MAP2 to describe neuronal maturation and polarization both in vivo and in vitro [14, 15, 108]. The mechanisms responsible for the selective targeting and/or retention of tau to the axon relative to MAP2 isoforms were a major focus of early studies of the establishment of neuronal polarity in the 1980s and 1990s, which identified features of both the coding sequence [109] protein phosphorylation state [110]

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and the tau mRNA [111]. Tau expression is sufficient to induce the growth of axonlike neurites from nonneuronal cells [112], and studies by Caceres and coworkers established that tau segregates to the axon early during initial neuronal polarization and played an essential role in the initial establishment of polarity in primary cultures of cerebellar magnocellular neurons [24]. The role of tau was found to be less clearly defined in cultured hippocampal pyramidal neurons [113] and tau knockout studies in situ [114], in which largely normal neuronal polarization and development was accompanied by selective upregulation of other axonal MAPs. A more recent study [115] suggests that several MAPs including tau, MAP2, and MAP1B may be needed at various times for both neuronal polarization and axonal outgrowth [116]. Axonal outgrowth and myelination The localization of tau to the growth cones of neurons by stage 3 of polarization [117, 118] suggested that tau has binding partners in addition to MTs, which are typically absent from GCs. Further studies of neuronal polarization and other developmental roles played by tau (such as its role in myelination) have highlighted the importance of non-MT-associated tau functions. For instance, disruption of F-actin networks prevents this localization and interferes with both neuronal polarity and normal growth cone motility and the guidance of axonal outgrowth [119]. Axonal tau is highly phosphorylated at the time of axonal polarization [110, 120] and phosphorylation of tau at the major MARK kinase sites (262 and 356) is necessary for both actin-tau association [121] and normal axonal outgrowth and pathfinding [56, 57, 76], even though it almost completely abolishes tau-MT interaction via the MTBRs, indicating that normal tau-actin interactions occur in an MT-independent manner. The activity of MARK kinases appears to act as a switch determining whether tau binds MTs or actin via the tau MTBR [55, 87, 121]. The involvement of tau with axonal identity and GC-mediated growth and guidance combined with the ability of tau to bind to, be phosphorylated by, and activate fyn kinase and other Srk family kinases [72–76] is suggestive of a general role for tau in modulating cytoskeletal function in response to external messages [17]. The myelination of CNS axons by oligodendrocytes is a complex event in which tau and fyn are both clearly involved in functional interneuronal coordination [122]. Neurons and oligodendroglia express both fyn and tau at high levels during CNS myelination, and either tau or fyn deletions prevent normal myelination from occurring [123]. It is likely that tau has other as yet unidentified functions that involve fyn-mediated signal transduction, and do not involve MTs, especially in light of the now-established role that both fyn and tau play in mediating the neurotoxicity produced by extracellular beta amyloid [75, 124].

15.3

Pathobiology of Tau

Tau-associated pathobiology is best understood in the context with respect to molecular mechanisms of tau misprocessing (mainly aggregation and hyperphosphorylation) implicated by familial tauopathies and their effects on the interaction of tau with other proteins. The pathological effects of tau misprocessing at the cellular level are much

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less well understood and are best viewed as a dysregulation of normal tau functions: that is, of axonal polarization and outgrowth during development and the maintenance of axonal identity and function in the adult. Although tau pathobiology at the systemic level is almost completely unexplored, recent reports of tau secretion and interneuronal transfer in tauopathy disease models may prompt a reappraisal of the traditional scientific and clinical view of these diseases.

15.3.1

Tauopathy-Associated Abnormalities at the Molecular Level

It is now clear that tau either induces or mediates significant cytopathological changes that play central roles in both AD and several other neurodegenerative conditions (tauopathies) in humans, including Pick’s disease, corticobasal degeneration, and progressive supernuclear palsy in which tau plays a demonstrably pivotal etiological role. In most of these conditions, tau overexpression does not appear to be significantly involved, although the H1 tau haplotype associated with some tauopathies [125] has recently been shown to correlate with high tau expression [126]. Rather, tauopathy mutations induce toxic changes in the folding, phosphorylation state, and intermolecular interactions of tau which largely appear to be pathological consequences of abnormal dissociation of tau from MTs. These consist mainly of the aggregation and/or hyperphosphorylation of the tau C-terminal MTBR domain, which may occur in combination with additional toxicity pathways involving the tau N-terminal projection domain. Familial tauopathy mutations Familial tauopathies offer the most straightforward approach to understanding how abnormalities in tau processing cause human disease. Human tauopathies almost invariably come in both “sporadic” and familial forms, with the latter almost invariably exhibiting autosomal dominant genetics, implying that mutant tau has gained a toxic function [127, 128]. An etiology based on gain of function is also consistent with the lack of conservation of non-MTBR regions of tau and the restriction of tauopathies to primates and a few nonprimate species [15]. Familial human tauopathy can be induced via: (a) dominant missense point mutations in the coding sequence (the majority), (b) intronic mutations that affect the frequency of exon 10 inclusion and disrupt the ratio between 3R and 4R tau isoforms expressed, and (c) deletion mutations in the coding sequence that do not change the reading frame (Fig. 15.3). All but two of the tau exonic point mutations are located within the C-terminal half of tau in or near the MTBR repeat region [100, 125, 128] and all but one of them reduce the affinity of the mutants to microtubules and increase the tendency of the mutant tau to polymerize into filaments [100, 125, 129–132]. Either of these effects might exacerbate aggregation-associated tau toxicity either directly or via effectively increasing cytosolic tau levels via the failure to bind microtubules. The effect of deletion tauopathy mutations (e.g., delta 280) on both tau–tau aggregation and MT binding appears to be similar to that of the missense mutations [128]. Mutations in tau that affect splicing of the fourth MTBR (exon 10) cause a significant

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Fig. 15.3 Characteristics of non-MT-associated tau schematic summarizing current knowledge of events and modifications involving non-MT-associated tau that may be disease-related. This is based on the consensus assumption that tau cytotoxicity and tauopathy results from abnormal processing or distribution of tau that occurs only when it becomes disassociated from MTs. Regions of tau that affect secretion, uptake, and transfer of tau protein [88, 193, 265–268]. Calpain-mediated cleavage of the tau N-terminal releases a 45–230 toxic fragment similar to those produced by caspase activation and exposure to extracellular beta amyloid. Progressive aspase and calpain cleavage of regions flanking the MTBR increase aggregation tendency and eventually produce a minimal sequence (F3) capable of generating PHF tau. Tauopathy mutation sites nearly all of the 40 missense mutations are at sites within or immediately adjoining the MTBR (black arrows—numbering according to the longest tau CNS isoform) between residues 257 and 406 [100]. Only two missense mutations (R5H and R5L) at position 5 on the tau N-terminal (red arrow) are located away from the MTBR at a residue essential for tau-dynactin binding. All nine intronic tauopathy mutations identified to date affect alternative splicing of Exon 10 (red/black arrows), which encodes the fourth MTBR. Similarly, the sites of deletion point mutations that cause human tauopathy are also shown (white down arrows)

proportion of non-AD familial tauopathies by inducing a syndrome-specific overrepresentation (13 mutations) or underrepresentation (2 mutations) of exon 10 in the pathological tau aggregates. Curiously, it applies to both the sporadic as well as the familial forms of diseases such as Pick’s disease (3R-dominant) and frontotemporal dementia (4R-dominant), suggesting that imbalances in the 3R:4R ratio of tau isoforms in either direction lead to toxicity and neurodegeneration, although it is not clear whether this occurs via increasing the toxicity of the aggregates themselves. The morphologies of 3R- and 4R-dominant aggregates do vary characteristically from one another [133–135], and it is no longer implausible to suggest that different toxic conformations taken on by isoforms of a single protein might generate clinically distinct syndromes [136]. Having said this, it remains very unclear how asymmetry between 3R and 4R isoforms exacerbates tau-induced toxicity. “Gain of toxicity” mechanisms—tau aggregation and hyperphosphorylation The accumulation of hyperphosphorylated tau aggregates is the hallmark feature of human tauopathies and has been consistently correlated with neuronal degeneration and death in human patients, suggesting that some event associated with their formation plays a central role in tauopathy pathogenesis [138–140]. It is now well established that tau aggregates form via charge-charge interaction between the MTBRs of tau molecules, resulting in the formation of highly ordered helical, ribbon-like or filamentous polymers including paired helical filaments (PHF) and straight filaments [29, 44, 95, 100, 127–129]. The morphology, distribution, and exon content of tau aggregates varies characteristically among these conditions, suggesting that aggregates

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may be etiologic factors in human tauopathy. However, while the general nature of the alterations that are associated with and which may drive tau aggregate formation and hyperphosphorylation seem relatively well understood, the way in which tau aggregation/phosphorylation states and tau splicing patterns interact with diseasespecific variables to produce tauopathic lesions and syndromes is not yet clear. Almost all tauopathy mutations increase both aggregation tendency and phosphorylation state [128, 131–136], and both aggregation and the balance of kinase/phosphatase activities on tau are profoundly affected by the immediate consequences of tau-MT dissociation [141, 142], particularly in disease [83, 143] making them difficult to distinguish mechanistically under in vivo conditions [44, 45, 100, 125]. It now appears that both hyperphosphorylation of Ser/Thr residues flanking the MTBRs of tau and the oligomerization and/or aggregation associated with it mediate tau toxicity via discrete downstream mechanisms. Tau aggregates obstruct protein turnover Modifications such as the truncation [140, 144–146], phosphorylation [147, 148], ubiquitination [149, 150], enzymatic cross-linking [151, 152], nitration [153], glycation [154, 155], and glycosylation [156] which often accompany hyperphosphorylation of tau aggregates are very likely markers of reduced tau turnover via the ubiquitin/proteasome mechanism [157, 158]. Tau aggregates that cannot be degraded by proteosomes for steric reasons [159, 160] are cleared by alternative turnover mechanisms (i.e., macroautophagy and lysosomal activation; [161–163]) which are themselves characteristically inhibited in tauopathies [164] leading to progressive protein accumulation and eventually to oxidative stress and Ca2+ -mediated toxicity. Obstruction of protein turnover mechanisms is a major feature in AD and other aggregation-associated neurodegenerative diseases [165–167], and it seems likely that sequestration-induced loss of chaperone proteins that interact with tau under normal circumstances may exacerbate both tau aggregate formation and dysregulation of proteosomal and autophagosome-mediated tau turnover [168, 169]. Sequestration-based deficits induced by tau aggregates Since the dominant inheritance pattern of familial tauopathies is typically predictive of gain of toxicity pathogenesis [170] rather than via loss of normal tau function, it remains likely that tau-induced toxicity produces loss-of-function effects indirectly via the failure of tau-MT binding [171], since both tau hyperphosphorylation and MT loss closely accompany disease progression [172]. It now seems clear that tau aggregates can indirectly cause loss-of-function defects by sequestering tau and related MAPs in their immediate vicinity, thereby destabilizing local MTs [173, 77]. A similar mechanism might play a role with respect to actin and actin-associated proteins leading to the formation of actin–rich inclusions such as Hirano bodies [174]. Tauopathy mutant-specific abnormalities in tau-membrane interactions accompany neurodegenerative changes in fly [89, 90] and lamprey tauopathy models [91] which may also involve cooperative conformational changes between the tau N-terminus and sites in the MTBR domain to produce aggregation-associated epitopes [92] and/or dysfunctional interactions involving other actin-associated proteins such as dynactin binding [93]. The binding of the tau MTBR to polyanions such as nucleic acids

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or heparan sulfate proteoglycans favors tau aggregate formation [175–178]. Such aggregates are exacerbated by tauopathy mutations [179] and are a neuropathological feature in human disease [176], but it is not clear whether they are themselves toxic. Aggregates versus oligomers as toxicity agents While the spatiotemporal distribution of NFTs within the brain has long been closely correlated with the spread of clinical deficits in AD and a number of other tauopathies [138, 180], some more recent studies have called into question the common assumption that there is a causal linkage between NFT formation and neurodegeneration [181–186], suggesting that the most toxic moiety of tau is oligomeric rather than aggregated. Since tau is a cytosolic protein, aggregation of tau has generally been assumed to occur in the cytoplasm, causing (or as a direct consequence of) hyperphosphorylation [45]. Oligomer formation directly from MT-associated tau has been demonstrated [185], and granular aggregates have been identified in neurons during what appear to be the earliest stages of neurofibrillary degeneration in AD [187]. However, oligomerization/polymerization of tau can also be potentiated by the presence of fatty acids such as arachidonic acid [188] in vitro. Nascent NFTs in AD brain are associated with plasma membrane [189], and NFTs eventually become fyn positive both in AD [190] and in tg tauopathy mice [191]. Moreover, tau is associated with the plasma membrane and trafficking vesicles in tauopathy models [91, 192–194]. These findings suggest the involvement of trans-Golgi vesicle trafficking in tau misprocessing and also illustrate just how little is really known about the generation of toxic tau oligomers and/or aggregates in human disease. The involvement of vesicleassociated tau with pathological tau secretion [91, 194] is discussed in more detail below. Toxicity mechanisms involving the tau N-terminal The tau N-terminal “projection” domain has two identified roles in tau pathobiology. Tau can take up a “paperclip” conformation upon release from MTs in which the N-terminus folds over part of the MTBR that is identifiable by conformation-specific “disease” mAbs such as Alz50 [92, 195] which label tau in the early stages of pathological hyperphosphorylation and/or aggregation inAD [195–197]. The presence of the N-terminal can also act as a modulator of MTBR domain aggregation [197]. A more important mechanism may involve disease-specific interactions between the tau N-terminal and extracellular beta amyloid peptide [198], which now appear to play an essential role in AD pathogenesis [17, 18, 123, 199] and in the transduction of A beta toxicity in particular in both cell culture [200, 201] and murine transgenic models [202, 203]. This toxicity mechanism appears to be entirely independent of the MTBRs [199] and appears to operate via the generation of toxic N-terminal fragments cleavage by activated calpain and caspases [204–206]. One implication of a separate N-terminalmediated toxicity pathway is that it could provide a self-amplifying mechanism that could account for the existence of the clinically identical sporadic versions that exist for most familial tauopathies [100].

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15.3.2 Abnormalities in Cellular Function in Tauopathy Neuropathological features of tauopathy appear to reflect the pathological alteration of normal tau functions in: (a) the regulation of axonal outgrowth and the establishment of neuronal polarity during development and (b) the maintenance of axonal identity and function in fully differentiated neurons. Axoplasmic transport dysfunction in tauopathy Inhibition of “fast” MT-mediated transport has been consistently but not invariably induced by tau overexpression and/or the expression of tauopathy mutant tau isoforms in model systems [207–214] where it accompanies and in some cases precedes tau-induced degeneration [209]. The actual mechanism by which tau (and especially tauopathy mutant tau) affects fast transport is still under debate, with some studies showing an effect specific to the tau N-terminal domain [213], others showing isoform and motor protein-specific effects and no N-terminal involvement [211, 212], and still others [210] showing no effect at all of exogenous tau overexpression on transport. Abnormalities in fast axoplasmic transport are found in early AD neuropathology [215], but have not yet been specifically linked to tau misprocessing inAD [216]. Nonetheless, these findings raise the possibility that tau misprocessing could exacerbate disease progression via the disruption of growth factor transport to vulnerable neurons in a manner similar to that of other aggregation-prone “disease” proteins [217–220], or possibly by directly disrupting neuronal polarity in a manner similar to proximal axotomy (discussed below). Aberrant axonogenesis and loss of neuronal polarity The “hallmark” cytoskeletal changes seen with the development of AD and other tauopathies consist of the progressive displacement of tau from the axon to the soma and dendrites to form NFTs and the proliferation of axon-like neurites (neuropil threads) in the brain. The possibility that this pathology might reflect an reactivation of axonogenesis and loss of neuronal polarity due to aberrant tau function in AD was first highlighted by the identification of tau as the major component of accumulation in somatodendritic cells [42, 43], the observation that tau is lost from the white matter in AD [221] and the identification of tau as a driver of neuronal polarization [23]. In particular, the uncanny resemblance of neuropil thread sprouting from dendrites in AD and other tauopathies [222–225] to the ectopic axonal sprouts that emerge from the dendrites of CNS neurons in a variety of species after perisomatic axotomy [226–230] suggested that neuronal polarity loss could be a key feature in AD pathogenesis [231, 232]. This possibility has also been supported by more recent studies in cell culture of both the initial establishment of neuronal polarity [223] and axotomy-induced polarity loss [234, 235] showing that these events are regulated by tauopathy-associated proteins [57, 58, 236]. Kinases (e.g., MARK 1–2, GSK3 beta, p706S6K) and phosphatases (PP2A) involved in both axonal outgrowth and the abnormal phosphorylation of tau in AD have been shown to phosphorylate and cause the redistribution of [237–243], or be regulated by [244] proteins that regulate neuronal polarity, many of which are themselves upregulated in AD [243, 245, 246].

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Moreover, tau mislocation suggestive of polarity loss precedes and appears to be prerequisite to neurodegeneration in tauopathy model systems. The first mice transgenic for tau expressed 4R and 3R WT tau isoforms (respectively) without overexpression and showed tau mislocation to the dendrites but not degeneration [247, 248], while an in vivo tauopathy model generated by overexpressing WT tau in specific lamprey CNS neurons, showed that the phosphorylation and routing of exogenous tau to dendritic tips occurred in advance of degeneration [249], a pattern that was later replicated in transgenic mice either with higher expression levels [250, 251] or (more robustly) by expression of tauopathy mutant tau [209, 252] and in transgenic models with inducible tau expression [182, 183]. Taken together, these findings amount to a strong (albeit still circumstantial) case that a primary cellular defect in tauopathies is a failure of neurons to maintain neuronal polarity as well as other axonal functions, and suggest that further study of the cellular consequences of polarity loss might shed light on the cytopathogenesis of tauopathies. Implications for neurodegenerative disease One of the key implications of viewing tauopathy in terms of polarity loss is that the indirect proximal axonal injury (“secondary axotomy”) frequently caused by head trauma may also induce polarity loss and aberrant sprouting [253, 254]. This could account for the well-documented increased risk of developing neurofibrillary degenerative disease posed by repeated traumatic brain injury [255], since dementia could plausibly be triggered or exacerbated by miswiring of normal circuitry induced by injury-associated ectopic axonogenesis, especially in the cortical gray matter—see [256]. It should also be noted that the novel association of hyperphosphorylated somatodendritic tau with membrane trafficking pathways caused by polarity loss in AD [257] could recruit tau into unconventional protein secretory pathways, causing or exacerbating interneuronal spread of tau in tauopathies (discussed below).

15.3.3

Interneuronal Aspects of Tauopathy: Do Toxic Tau Lesions Spread Through the Brain?

Although key systemic features of tauopathies have been well characterized, the mechanisms that generate them have as yet received little attention. A characteristic feature of all tauopathies including AD is that the pattern of neurofibrillary lesion development responsible for generating the specific clinical syndrome in each disease suggests that transsynaptic influences [258, 259] are as significant as intrinsic vulnerability of specific neuron groups [260] to the pattern of disease development in the brain. However, it is not currently known whether such influences are due to: (1) growth factor imbalances associated with defects in axoplasmic transport [214], (2) interneuronal transfer of toxicity without actual tau protein transfer, or (3) an actual prion-like spread of misprocessed tau through the brain via synaptically connected and adjacent neurons [261, 262].

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Evidence for and implications of interneuronal tau transfer in tauopathy Until very recently, it has generally been assumed that tau is an exclusively intracellular protein, since it lacks the coding sequence motifs (i.e., signal sequence, lipidation/GPI anchor consensus sites) which traditionally target proteins for secretion and/or membrane association. This assumption has proved remarkably durable in light of: (1) the recent elucidation of multiple “unconventional” secretion pathways [263] and (2) demonstrations that other key aggregation-prone proteins in neurodegenerative disease (i.e., beta amyloid, alpha synuclein, and prion protein) are secreted via related unconventional mechanisms [264–266] and occasional reports of tau secretion as a by-product of neuroprotection [267] and the toxicity of extracellularly applied tau [50, 268]. However, reports of tau secretion to and uptake from the extracellular space in cell culture [88, 193, 269] and transneuronal tau movement between live neurons in situ [88, 270] have now been published by several laboratories, causing a more widespread reappraisal of this question and raising the possibility that interneuronal tau transfer might contribute to the pathogenesis of tauopathy in the brain as a whole. Initial studies suggest that tau secretion requires the presence of as yet unidentified elements in the tau N-terminal [88], is blocked by the presence of exon 2 of tau [193], and is enhanced by the removal or inactivation of the C-terminal microtubule-binding domain, resulting in the secretion of N-terminal cleavage fragments in a pattern similar to that found in the cerebral spinal fluid (CSF) of human AD patients [88, 193]. Aggregation of the MTBR domain and the presence of tauopathy mutations in the MTBR that affect aggregation [45] also appear to affect secretion [88], uptake [269] and toxicity [268] in cell culture, and tau transfer and/or tau-induced toxic changes in situ [88, 270], suggesting that tauopathy-induced abnormalities in tau aggregation and phosphorylation state may also induce tau secretion. These findings appear to place tau with other aggregation-prone proteins involved in human neurodegenerative disease that are known to be secreted [245] and could have significant clinical implications for the diagnosis and/or treatment of human tauopathy. Examples include: (1) the possibility that some of the increase in CSF-tau in patients with early-stage AD [271] and other tauopathies [272] might be antecedent (rather than consequent) to neuronal loss and (2) that the observed synergy of tauopathy with Parkinson’s Disease, Lewy Body dementia, and prion diseases [273, 274] might be due to pathological interactions between these proteins in a common pathological cellular pathway that results in secretion [275, 276].

15.4

Conclusion

It is now clear that abnormalities in tau biology play a central role in the development and spread of tau-containing neurofibrillary lesions within the human brain, but a great deal about the process by which this happens remains completely unknown. The pathobiology of tau protein in humans is increasingly well understood at the molecular and cellular levels, and therapeutic targets such as tau hyperphosphorylation and aggregation are now being investigated systematically. However, researchers

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are still only beginning to approach tau disorders at the organismal level. While it is as yet unclear whether abnormalities of axoplasmic transport and tau secretion (in particular) are significant factors in the pathogenesis of human tauopathies, their potential significance in providing new targets for the development of new diagnostic markers and therapeutic targets seems clear.

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248. Brion JP, Tremp G, Octave JN (1999) Transgenic expression of the shortest human tau affects its compartmentalization and its phosphorylation as in the pretangle stage of Alzheimer’s disease. Am J Pathol 54:255–270 249. Hall GF, Yao J, Lee G (1997) Tau overexpressed in identified lamprey neurons in situ is spatially segregated by phosphorylation state, forms hyperphosphorylated, dense aggregations and induces neurodegeneration. Proc Nat Acad Sci USA 94:4733–4738 250. Ishihara T, Hong M, Zhang B, Nakagawa Y, Lee MK, Trojanowski JQ, Lee VM (1999) Agedependent emergence and progression of a tauopathy in transgenic mice overexpressing the shortest human tau isoform. Neuron 24:751–762 251. Spittaels K, Van Den Haute C, Van Dorpe J, Bruynseels K, Vandezande K, Laenen I, Geerts H, Mercken M, Sciot R et al (1999) Prominent axonopathy in the brain and spinal cord of transgenic mice overexpressing four-repeat human tau protein. Am J Pathol 155:2153–2165 252. Lewis J, McGowan E, Rockwood J, Melrose H, Nacharaju P, Van Slegtenhorst M, GwinnHardy K, Paul Murphy M et al (2000) Neurofibrillary tangles, amyotrophy and progressive motor disturbance in mice expressing mutant (P301L) tau protein. Nat Genet 25:402–405 253. Uryu K, Chen XH, Martinez D et al (2007) Multiple proteins implicated in neurodegenerative diseases accumulate in axons after brain trauma in humans. Exp Neurol 208:185–192 254. Singleton RH, Zhu J, Stone JR, Povlishock JT (2002) Traumatically induced axotomy adjacent to the soma does not result in acute neuronal death. J Neurosci 22:791–802 255. McKee A, Cantu R, Nowinski C et al (2009) Chronic traumatic encephalopathy in athletes: progressive tauopathy after repetitive head injury. J Neuropathol Exp Neurol 68:709–735 256. Lafont F, Rouget M, Triller A, Prochiantz A, Rousselet A (1992) In vitro control of neuronal polarity by glycosaminoglycans. Development 114:17–29 257. Ittner LM, KeYD, Delerue F, Bi M, Gladbach A, van Eersel J, Wölfing H, Chieng BC, Christie MJ, Napier IA, Eckert A, Staufenbiel M et al (2010) Dendritic function of tau mediates amyloid-beta toxicity in Alzheimer’s disease mouse models. Cell 142:387–397 258. Su JH, Deng G, Cotman CW (1997) Transneuronal degeneration in the spread of Alzheimer’s disease pathology: immunohistochemical evidence for the transmission of tau hyperphosphorylation. Neurobiol Dis 4:365–375 259. Armstrong RA, Cairns NJ, Lantos PL (2001) What does the study of spatial patterns tell us about the pathogenesis of neurodegenerative disorders? Neuropathology 21:1–12 260. Götz J, Schonrock N, Vissel B, Ittner LM (2009) Alzheimer’s disease selective vulnerability and modeling in transgenic mice. J Alz Dis 18:243–251 261. Miller G (2009) Could they all be prion diseases? Science 326:1327–1329 262. SydowA, Mandelkow EM (2010) ‘Prion-like’propagation of mouse and human tau aggregates in an inducible mouse model of tauopathy. Neurodegenerative Dis 7:28–31 263. Nickel W (2005) Unconventional secretory routes: direct protein export across the plasma membrane of mammalian cells. Traffic 6:607–614 264. Emmanouilidou E, Melachroinou K, Roumeliotis T, Garbis SD, Ntzouni M, Margaritis LH, Stefanis L, Vekrellis K (2010) Cell-produced alpha-synuclein is secreted in a calciumdependent manner by exosomes and impacts neuronal survival. J Neurosci 30:6838–6851 265. Fevrier B, Vilette D, Archer F, Loew D, Faigle W, Vidal M, Laude H, Raposo G(2004) Cells release prions in association with exosomes. Proc Natl Acad Sci USA 101:9683–9688 266. Rajendran L, Honsho M, Zahn TR, Keller P, Geiger KD, Verkade P, Simons K (2006) Alzheimer’s disease beta-amyloid peptides are released in association with exosomes. Proc Natl Acad Sci USA 103:11172–11177 267. Hall GF, Lee S, Yao, J (2002) Neurofibrillary degeneration can be arrested in an in vivo cellular model of human tauopathy by application of a compound which inhibits tau filament formation in vitro. J. Mol. Neurosci 19:253–260 268. Gomez-Ramos A, Diaz-Hernandez M, Cuadros R, Hernandez F, Avila, J (2006) Extracellular tau is toxic to neuronal cells. FEBS Lett 580:4842–4850 269. Frost B, Jacks RL, Diamond MI (2009) Propagation of tau misfolding from the outside to the inside of a cell. J Biol Chem 284:12845–12852

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Chapter 16

Tubulin-Related Malformations of Cortical Development Xavier H. Jaglin, Jamel Chelly and Nadia Bahi-Buisson

Abstract The importance of the microtubule cytoskeleton during in utero brain development has emerged from a body of functional and genetic studies and was recently strengthened by the description of tubulin-related malformations of cortical development characterized by the disorganization of the cerebral cortex and the presence of ectopic neurons. Tubulin genes encoding specific isotypes of alpha(TUBA1A, TUBA8) and beta-tubulins (TUBB2B, TUBB3) are associated with a spectrum of neuronal migration disorders ranging from a simplification of the folded aspect of the brain surface to a complete absence of folds (lissencephaly). The spectrum also encompasses forms of polymicrogyria characterized by an excessive number of small brain folds. Major axonal tract disruptions are also observed in combination with the aberrantly located neurons. Biochemical investigations have shown that an important number of the different mutations in TUBA1A, TUBB2B, and TUBB3 lead to folding and heterodimerization impairments, defective incorporation into microtubules, and/or alterations of microtubule dynamics and stability. This abnormal homeostasis of microtubules during neuronal polarization and migration might contribute to the disorganized cortical cytoarchitecture observed in patients as well as to the abnormal development of the major axon tracts connecting the cortex and various subcortical structures.

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Introduction

Progress in deciphering genetic causes of malformations of cortical development was recently achieved through the implication of a variety of tubulin genes [1–6]. Indeed, molecular investigations carried out on human patients with unexplained abnormalities of cortical development led to the description of new syndromes N. Bahi-Buisson () · X. H. Jaglin · J. Chelly Laboratoire de Génétique des Maladies Neurodéveloppementales, CNRS UMR 8104, Institut Cochin, Université Paris Descartes, 75014 Paris, France Institut National de la Santé et de la Recherche Médicale (Inserm), U1016, 75014 Paris, France e-mail: [email protected] N. Bahi-Buisson Inserm U1016, Institut Cochin, Université Paris Descartes, 24 rue du Faubourg Saint Jacques, 75014 Paris, France

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associated with mutations in four tubulin genes: TUBA1A [2], TUBB2B [1], TUBA8 [3] and TUBB3 [4] encoding specific isotypes of α- (TUBA1A, TUBA8) and β-tubulins (TUBB2B, TUBB3). Clinically, the patients demonstrate moderate-to-severe mental retardation and refractory epilepsy, associated with a spectrum of neuroanatomical abnormalities. These malformations encompass gyral abnormalities that range from gyral simplification and lissencephaly to polymicrogyria and major axonal tract disruption. These tubulin-related forms of cortical dysgeneses are thought to be a combination of abnormal neuronal migration (leading to a disorganized cortex with the presence of heterotopic neurons), differentiation, and axonal guidance (leading to projection axonal tract disruption). The interest of the neurogeneticist community was driven a few years ago toward microtubules and tubulin genes from the initial description of type I lissencephaly (LIS) associated with mutations in the Lissencephaly I gene (LIS1)—encoding a protein which interacts with microtubules—as well as in the MicrotubuleAssociated Protein (MAP)-encoding gene DCX (Doublecortin) [7–9]. These forms of lissencephalies encompass a large range of cortical abnormalities, extending from agyria to pachygyria, either posterior- (LIS1-associated pattern) or anteriorpredominant (DCX-associated pattern). Following these discoveries, the importance of the cytoskeleton in neuronal migration further emerged from a body of functional and genetic studies, which contributed to a better understanding of these dramatic diseases. This has been gleaned from the contribution of actin and its associated proteins such as Filamin A, since mutations in the FLNA (Filamin A) gene impairs the migration of neurons out of the germinal zone and leads to nodular periventricular heterotopia [10]. These human diseases offered the investigators—interested in understanding the developmental programs of the cerebral cortex—the possibility to focus on fundamental and critical molecules and processes. In the present chapter, we will discuss how the recent development in human genetics contributed to improve our understanding of neuronal migration disorders and of the critical molecular actors of neuronal migration in the cerebral cortex.

16.2 The Development of the Mammalian Brain Cortex Historically, two main streams of work have converged to better understand the developmental processes underlying the establishment of a properly organized cerebral cortex. First, a fine and thorough description of the developing cortex in primates pioneered by Rakic and Caviness led to the emergence of strong fundamentals and principles, as the existence of both radial and tangential migrations suggested by the observation of a dichotomy between sites of neuronal proliferation and differentiation as well as sites of differentiated functions. The concept of “inside-out” organization was also underlined by their work [11–14]. Further studies from different groups brought up major contributions, which contributed to uncover the lineage of radial glial cells, intermediate progenitor cells, pyramidal neurons, and interneurons [15–19]. The second stream of work consists in a top to bottom approach based

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on data generated from human genetic investigations and analyses of mouse models. For instance, the discovery of the Reelin gene has been allowed by the previous description of the Reeler mice that display obvious neuronal migration defects. Taken together, genetic investigations carried out in the last two decades led to a significant improvement in the knowledge of the molecular basis of cortical development as well as a considerable understanding of congenital forms of malformations of the cerebral cortex. Pyramidal neurons and interneurons are the functional units of the cerebral cortex. Throughout all the developmental period (mainly during the in utero period in most mammals, including rodents and human), these two types of cells undergo a precisely orchestrated succession of events from proliferation and neurogenesis to migration, differentiation, and synaptogenesis that lead to the formation of complex functional networks in the adult. In rodents and humans, the regions containing pyramidal neuron and cortical interneuron progenitors are located away from the cortex. An initial period of proliferation and neurogenesis produces the pool of pyramidal neurons and interneurons in pallial or subpallial ventricular zones, respectively. Human and mouse cerebral cortices are characterized by a six-layered structure. This remarkable organization results from successive waves of radially migrating neurons. Extensive studies have elaborated and documented the model of an “insideout” pattern of migration, which establishes that the younger neurons migrate farther than previously generated neurons. The older neurons settle in the cortical plate forming a homogenous layer of cells, which is further crossed by following pyramidal neuron generations. This pattern of migration tends to place younger neurons in the superficial layers of the cortex. Thus, the joint action of proliferation and neuronal migration leads to changes in size and structure of the cortex. The young postmitotic neuroblasts should orient adequately prior to the initiation of migration toward cortical areas to which they are intended. This involves various morphological modifications and the development of signaling pathways tailored to adapt the cell structure to the migration. Ventrally derived interneurons shall subsequently migrate tangentially to the surface of the telencephalon toward the cortex whereas dorsally derived pyramidal neurons will migrate perpendicularly to the surface of the cortex to reach the developing cortical plate (Fig. 16.1). Thus, structural abnormalities of these networks caused by impaired neurodevelopmental processes are often dramatic, resulting in cognitive deficits, types of psychotic disorders, and/or epilepsy.

16.2.1

Hallmarks of Brain Cortical Development

For a better understanding of this discussion, we describe a few chronological hallmarks of murine cortex development. First, the onset of neurogenesis occurs at about days E9/E10 with the appearance of radial glia cells followed by the first asymmetric divisions. This period ends about 8 days later. Before day E10, the neural tube (which closes at E9) consists of a proliferative neuroepithelium, forming the ventricular zone (VZ). The first wave of postmitotic neurons migrates toward the pial

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Fig. 16.1 Development of the mammalian cerebral cortex. a Schematic representation of rodent embryonic telencephalon. Two main types of neurons are found in the central nervous system: inhibitory interneurons and excitatory projection neurons. They, respectively, originate in the medial ganglionic eminence (MGE) in the ventral telencephalon and in the ventricular zone of the dorsal telencephalon. Postmitotic interneurons have to migrate upward following a path that is tangential to the surface of the cortex. Postmitotic projection neurons migrate radially to reach their final location and further settle within the cortical plate following an inside-out pattern. The younger neurons settle in lower layers whereas the latter neurons integrate in higher layers, further increasing cortical thickness. b The upper inset illustrates neurogenic asymmetrical divisions of radial glial cells (RGC, in gray) in the ventricular zone that subsequently give rise either to one progenitor and one multipolar intermediate progenitor cell (IPC) located within the subventricular zone (SVZ) or directly to postmitotic neurons. Postmitotic projections neurons are also born from symmetrical mitosis of the IPC and further adopt a bipolar shape to migrate radially toward the cortical plate (CP) along the RGC processes. The lower inset shows a tangentially migrating interneuron displaying a characteristic dynamic shape, splitting, extending, and retracting branches at its leading edge

surface of the neural tube along the fibers of radial glia and forms the preplate (PP), which is a transient compartment. It contains different types of early neurons, as the Cajal-Retzius cells [20]. Neurogenic mitoses also generate intermediate progenitor cells, which accumulate in a separate compartment called subventricular zone (SVZ). Subsequent waves of neurons generate the cortical plate (CP) and further split the preplate into two structures: the marginal zone (MZ), under the pial surface, and the subplate (SP) between the CP and the future VZ. The entire area between the SVZ and CP, that is, the space that contains no progenitor cells or postmitotic resident was defined intermediate zone (IZ) by the Boulder Committee (for a comprehensive review on human cortex development and nomenclature, see [21]. Finally, the laminar structure of six layers of the CP meets the “inside-out” organizational scheme as discussed above. The younger neurons move toward the surface layers of the cortex passing through the deeper layers [11, 13, 17, 22]. The MZ, located under the pial surface, is referenced as layer I. The deepest layer is the layerVI.

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Neuronal Migration

The radial migration of pyramidal neurons between the ventricular zone and the cortical plate of the developing cerebral cortex may occur following two different modes of migration. The first one is named somatic translocation and concerns neurons that display an apical process attached to the basement membrane at the pial surface and that move their nucleus forward, in the direction of the cortical plate. The movement of the nucleus is regular; its speed varies among neurons but is constant (10–50 µm/h in vitro [23]). This mode of migration mainly concerns early postmitotic neurons fated to reach the preplate [24, 25]. The second mode of migration is called locomotion. The vast majority of pyramidal neurons use this mode of migration. They migrate freely within the neuropil, displaying a bipolar morphology with a leading and a trailing process, directed toward the cortical plate and the ventricular zone, respectively. The leading process is short and slightly branched. The migrating neuron is attached to the radial process of a radial glial cell, which exerts a guiding role. Neurons in locomotion migrate radially with a saltatory pattern at a speed that ranges from 10 to 20 µm/h [13, 19, 23, 26]. Once arrived close to its final destination, the migrating neuron is released from the radial glia and may transiently use a somal translocation mode to reach its final position [23]. Nonpolarized cells have been observed for more than 30 years on fixed tissues [18, 27–29]. Further studies investigated this transitory multipolar stage and revealed that the multipolar cells present an unstable morphology, extending and retracting many processes. Their migration speed is relatively slow (2 µm/h) and is not associated with any conspicuous displacement [19, 30]. Lineage analyses provided further evidences that transient multipolar cells can undergo symmetrical divisions late in corticogenesis, giving rise to two neurons. These observations led the authors to conclude that multipolar cell populations partially overlap with the intermediate progenitor cell populations [19, 31, 32]. Following this multipolar migration step, neuroblasts become bipolar.

16.2.3

Polarization Precedes Migration

The polarization step corresponds to the symmetry break undergone by young postmitotic neuroblasts, which occurs right after the differentiative mitosis. This process is regulated as follows: both intrinsic and extrinsic factors activate extracellular receptors (1), which transduce the signal through a variety of signaling pathways (2) that target both microtubules and actin microfilaments (3). To our knowledge, four signaling pathways have been described that are involved in the initiation and maintenance of neuronal polarity (for review, see [33]). The first one requires the activity of PI3K/PIP3/AKT/GSK3β [34–36]. The second one involves the Rho GTPases RhoA, Cdc42, and Rac1 [37, 38]. The PAR complex constitutes the third signaling complex. The fourth complex comprises the kinases LKB1 and SAD [39]. Rather than being independent, these pathways intermingle at many steps, thus creating a complex

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signaling network, whose targets are the microtubule-associated proteins (MAPs, MAP2, tau, APC, Stathmin) and actin-associated proteins (AAPs, Cofilin, Profilin). These proteins are responsible for the modulation of microtubule and actin cytoskeleton dynamics, that is, the local stabilization and destabilization of the microtubules and actin microfilaments, respectively [40–45]. This transition from multipolar to bipolar stage is a prerequisite for the initiation of radial migration and relies on the proper function of the signaling pathways mentioned above as well as the proper downstream modulation of cytoskeleton dynamics. Indeed, it has been shown by different groups that the experimental disruption of various genes related with these processes often leads to the disruption of the transition from a multipolar to a bipolar morphology [46–48]. The modulation of the cytoskeleton stability is not only involved in the early process of polarization, but also in the plasticity and sustainability of the migrating neuron’s morphology. For instance, the growth of the leading process requires a highly dynamic actin cytoskeleton and an elongating stable microtubule network, which are both under the control of molecular complexes involved in the polarization homeostasis [33, 49].

16.3

Human Disorders Associated with Mutated Tubulin Genes

Genetic studies in both humans and mice have identified a spectrum of mutations in genes involved in a large array of critical processes (cell proliferation, cell adhesion, cell migration, chemoattraction and repulsion, posttranslational modifications, and dynamics of the cytoskeleton) that often disrupts the development of the cerebral cortex and further leads to severe cortical malformations. Malformations of cortical development have been traditionally classified based on which biological process is likely to be affected and grouped under disorders of proliferation, migration, and cortical organization [50, 51]. In the disorders of proliferation (or of the balance between proliferation and apoptosis), the number of cells is significantly reduced, resulting in an abnormally small head (microcephaly). In the disorders of migration, neurons do not reach their correct destination in the cortical plate, either by remaining at the ventricular surface (periventricular nodular heterotopia, PNH), arresting in the white matter (subcortical band heterotopia, SBH) or forming a disordered, often thickened, cortical plate. This thicker cortex affects the formation of normal gyration, leading to a reduced gyral pattern or a smooth appearance of the cortical surface (lissencephaly or classic lissencephaly or type I lissencephaly). Other lissencephalies (cobblestone lissencephaly or type II lissencephaly) are associated with overmigration of neurons to the pial surface [52, 53]. Disorders of cortical organization or late migration, comprises mostly the polymicrogyrias, a heterogeneous group of malformations with multiple small gyri and an abnormally thin or thick cortex, sometimes so severely affecting brain structure as to cause clefting between the ventricular and meningeal surface (schizencephaly) [54].

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Clinical and Imaging Features of Tubulin-Related Malformation of Cortical Development

Lissencephaly (LIS, smooth brain) comprises a group of severe brain malformations associated with deficient neuronal migration that results in mental retardation, epilepsy and when severe, a shortened lifespan [50, 55–57]. In the most severe form, the surface of the cerebral hemispheres is completely smooth (agyria), whereas less severe forms of LIS are characterized by reduced folding patterns with abnormally broad gyri (pachygyria). Classic or type 1 LIS (agyria–pachygyria spectrum) is typically characterized by a loosely organized and markedly thickened four-layer cortex (compared with the normal six-layered cortical architecture) and apparently normal cerebellum, although some patients have mild vermis hypoplasia. However, some rare forms of LIS are associated with a disproportionately small cerebellum, referred to as LIS with cerebellar hypoplasia (LCH) [58]. Polymicrogyria (PMG) is a malformation of the cerebral cortex in which the usual gyral pattern is replaced by numerous small foldings, separated by sulci, which fuse inferiorly, and the normally six-layered cortex is replaced by a four-layered or unlayered cortex [50]. PMG can be a focal lesion or a more generalized cortical abnormality and may be accompanied by other malformations, including agenesis of the corpus callosum, microcephaly, or cerebellar vermis hypoplasia. Localized PMG is most commonly perisylvian, but has been described in all areas of the brain, with its extent and location determining the neurological manifestation [50, 59]. Tubulin-related cortical dysgeneses encompass a wide range of complex cerebral malformations from microlissencephaly to focal pachygyria, combined with alterations of interhemispheric commissure (i.e., corpus callosum) and cerebellum and brainstem. This heterogeneity of developmental disorders recapitulates multiple impairments in the different steps of brain development related to mutations in four genes encoding specific isotypes of α-tubulin (TUBA1A and TUBA8) and β-tubulin (TUBB2B and TUBB3) [1–5, 60–63]. Alterations of any of these tubulins are suspected to disrupt several steps of brain development including proliferation, neuronal early differentiation (that might lead to microcephaly), and migration of neuronal cells and pial basement membrane maintenance and integrity (leading to overmigration of neuronal cells into the leptomeningeal space) [5, 64, 65]. Mutations in α-tubulin 1A (TUBA1A) result in a new subset of agyria–pachygyria spectrum Twenty five de novo heterozygous missense mutations in α-tubulin 1A (TUBA1A) gene have been reported [2, 60–62, 66–68]. TUBA1A mutations are responsible for a wider spectrum of phenotypes than any other gene of lissencephaly (LIS1 and DCX). Most children harboring these mutations have microcephaly, severe motor and intellectual disabilities, and epilepsy. A subset has strabismus, pseudobulbar diplegia, and cerebellar symptoms according to the maximal location of brain malformations. On the basis of the maximum location and the severity of gyral abnormalities, the patients can be subdivided into three different groups with homogeneous clinical presentation and emerging common molecular findings [61, 62, 66]. In the

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Fig. 16.2 a T2 weighted sagittal. b T2 weighted axial. c T1 weighted axial. Section of MRI scan of the brain of a 10-year-old girl with R264C mutation in TUBA1A showing bilateral perisylvian, insular and opercular pachgyria (b, c), dysmorphic aspect of the basal ganglia suggesting a fusion between the caudate nucleus and putamen (b, c), and hypoplastic corpus callosum and brainstem (a)

Fig. 16.3 a T1 weighted sagittal. b T2 weighted axial. c T1 weighted axial. Section of MRI scan of the brain of a 6-month-old patient with R402H mutation in TUBA1A showing posteriorly predominant agyria-pachygyria, 12–20-mm-thick cortex (b, c), and hypolastic corpus callosum and mild cerebellar and brainstem hypoplasia (a)

first group, the most striking radiological hallmarks are a perisylvian-predominant pachygyria with thick cortex combined with a hooked aspect of the frontal horn of the lateral ventricles. The dysgenesis of the anterior limb of the internal capsule results in a more spherical shape of the head of the caudate nucleus, which is impinging in the inferolateral wall of the lateral ventricles. Because of this impingement, the shape of the frontal horns becomes more angular or hooked (Fig. 16.2). This combination constitutes a unique pattern that seems to be specifically associated with TUBA1A mutations. Most of the mutations detected in this group of patients correspond to the recurrent R264C mutation. The second group consists of classic LIS defined by extensive posteriorly predominant agyria–pachygyria, 12–20-mm-thick cortex and mild cerebellar hypoplasia (Fig. 16.3). In this group, mutations of the residue R402C/H are more prevalent. The third group consists of mild or moderate lissencephaly with moderate-to-severe cerebellar hypoplasia (LCH) and more consistent hypoplasia

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of the corpus callosum. In this group, distinct mutations have been reported giving more variable phenotypes with more variable severity. Very recently, TUBA1A mutations were also reported in bilateral perisylvian polymicrogyria suggesting a phenotypic overlap between the TUBA1A-, TUBB2B-, and TUBA8-associated spectrum [68]. Thin corpus callosum and hypoplasia of pons and cerebellum are almost common hallmarks of TUBA1A-related cortical dysgenesis. These features can be present in patients with lissencephaly due to mutations in other genes including LIS1, DCX, RELN, or ARX. However, it is the combination of all of the above mentioned features which is highly characteristic of mutations in TUBA1A, much more than the cortical malformation itself, which can range from agyria, pachygyria to polymicrogyria, or almost normal brain with mild brainstem and vermis hypoplasia. Mutations were also identified in five fetuses in which prenatal ultrasound, magnetic resonance imaging (MRI), and neuropathological examination revealed a wide lesional spectrum which consistently involved five brain structures including the neocortex, the hippocampus, the corpus callosum, the brainstem, and the cerebellum [60, 67, 69]. Compared with the phenotype of children mutated for TUBA1A, these prenatally diagnosed fetal cases display a more severe phenotype strongly suggestive of pathological mechanisms including impairment of neuronal migration and differentiation as well as axonal guidance disturbances. Fetal brain and infratentorial weights are below the 5th percentile. The brain surface is either agyric or pachygyric, with a postero-anterior gradient of severity. Cortical lamination is significantly disturbed and the six-layer cortex is reduced to four thicker layers and in some individuals only to two thin layers with an only discernible molecular layer and numerous heterotopic neurons arrested in the white matter of the cerebral hemispheres indicated a failure of the majority of neurons to migrate properly to their final destination. Basal ganglia are hypoplastic and dysplastic, sometimes asymmetric and thalamic nuclei appear prominent and displaced. Caudate and hypothalamic nuclei are dramatically reduced in size and shape. The internal capsule is hypoplastic and fragmented with aberrantly directed fascicles. The corpus callosum is either completely or partially absent. Cerebellar hypoplasia and brainstem hypoplasia (i.e., mesencephalon, pons, and medulla) are also prominent. Corticospinal tracts are hypoplastic and aberrantly placed. Mutation of the variant alpha-tubulin TUBA8 results in polymicrogyria with optic nerve hypoplasia TUBA8 is the second alpha-tubulin to be implicated in a specific aspect of cerebral cortical development [3]. Two consanguineous pedigrees segregating severe developmental delay, seizures, and optic nerve hypoplasia as a recessive trait have been reported to harbor the same homozygous 14-base pair intronic deletion upstream of exon 2 of the TUBA8 gene, altering transcript splicing and resulting in greatly reduced TUBA8 protein levels in patient-derived lymphoblastoid cells. Brain imaging of affected family members revealed extensive bilateral polymicrogyria, dysplastic or absent corpus callosum. In addition, there is a distinctive abnormality of the brainstem, in which the normal sharp demarcation between the pons and medulla was absent, with the pontine bulge extending too far caudally. Although these phenotypes converge with those described above, additional mutations are needed to definitively support TUBA8 as a polymicrogyria-associated gene.

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Fig. 16.4 T1 weighted axial (a, c) and sagittal (b, d) section of MRI scan of the brain of two different patients with TUBB2 mutation. The first patient (a, b) presented with diffuse polymicrogyria, dysmorphic basal ganglia, corpus callosum agenesis, or dysgenesis and brainstem and cerebellar atrophy. The second patient (c, d) presented with bilateral, perisylvian polymicrogyria, mild hypoplasia of the corpus callosum, and brainstem atrophy

Mutations in β-tubulin 2B (TUBB2B) result in a new subset of polymicrogyria TUBB2B is the first gene encoding a β-tubulin to be involved in malformations of cortical development [1]. Five heterozygous missense mutations in TUBB2B that are all sporadic and de novo have been reported. Similar to the TUBA1A phenotype, children harboring TUBB2B mutations are typically microcephalic and have severe motor and intellectual disabilities often accompanied by seizures. Patients with TUBB2B mutations share a complex brain disorder with bilateral polymicrogyria (Fig. 16.4). The polymicrogyria in affected individuals is bilateral,

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Fig. 16.5 T1 weighted axial (a, b) and sagittal (b) section of MRI scan of the brain of a 6-yearold boy with TUBB3 mutation showing global microgyria and dysmorphic and hypertrophic basal ganglia (a), vermian dysplasia (b), and corpus callosum hypogenesis combined with hypoplastic brainstem (c)

asymmetric, and typically more predominant in the frontal and temporal lobes. Similar to TUBA1A mutations, partial or complete agenesis of the corpus callosum, dysmorphisms of the basal ganglia, and cerebellum are common radiological findings. In addition, they all share a distinctive abnormality of the brainstem, in which the normal sharp demarcation between the pons and medulla was absent with the pontine bulge extending too far caudally that was also observed in patients with TUBA1A and TUBA8 mutations. Only one fetus with TUBB2B mutation (S172P mutation) is described in the literature [1]. Similar to living patients, the macroscopic examination demonstrates a complex brain disorder with bilateral, asymmetrical, and anteriorly predominant polymicrogyria, dysmorphic basal ganglia, corpus callosum agenesis or dysgenesis and, in most cases, cerebellar and pons atrophy. Neuropathological examination reveals perturbed cortical cell migration, disorganization of cortical layering, and overmigrating neurons in the leptomeningeal space through breaches of a defective pial basement membrane. This observation recalls the “cobblestone-like” phenotype of mice inactivated for Gpr56 [70], a gene associated with bilateral frontoparietal polymicrogyria in humans. Fetal observations, together with imaging features on living patients, suggest that TUBB2B plays a critical role in the development and/or maintenance of the pial basement membrane and suggest that disruption of the pial membrane contributes to the formation of polymicrogyria. Mutations in the neuronal β-tubulin subunit TUBB3 result in malformation of cortical development and ocular motility disorder: one gene, several phenotypes Missense mutations in the TUBB3 gene, encoding β-tubulin isotype III, were shown to be causative of a spectrum of brain malformations and neurological disabilities. Interestingly, certain phenotypes frequently segregate with particular amino acid substitutions [4, 63] (Fig. 16.5). Fourteen heterozygous missense mutations have been reported, which are a mixture of familial and de novo mutations, and six have been identified in more than one unrelated individual.

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The first eight TUBB3 missense mutations to be reported cause the paralytic eye movement disorder CFEOM3, which results from hypoplasia of the oculomotor nerves and secondary atrophy of extraocular muscles. Depending upon the specific amino acid substitution, the patients can develop sensorimotor polyneuropathy due to the progressive degeneration of motor and sensory axons in the limbs, and are sometimes born with wrist and finger contractures, facial paralysis, and mildto-moderate intellectual and behavioral disabilities. Brain malformations include agenesis or hypoplasia of commissural axon tracts, hypoplasia of the corticospinal tract, and dysmorphic basal ganglia with fusion of the caudate and putamen. Overall, this initial combination of clinical and radiological findings points to a generalized defect in axon guidance and maintenance [63]. The TUBB3 phenotypic spectrum was further expanded to complex cortical dysgenesis. Six additional TUBB3 missense mutations have been identified in individuals with malformations of cortical development [4]. These are characterized by cortical disorganization ranging from gyral simplification to polymicrogyria, cerebellar dysplasia, and hypoplastic brainstem. Basal ganglia dysmorphisms are also present, and thus appear to be a defining phenotype of the dominant tubulin-related disorders. These cortical malformations were not associated with ocular motility defects. Interestingly, the severity of TUBB3-related phenotype is related with mutation type, with a specific mutation associated with a particular set of symptoms. Of these, patients with R262C recurrent mutation, the most commonly observed in Tischfield’s series clearly demonstrate isolated CFEOM3, while those with R262H, E410K, or D417H also showed varying degrees of facial paralysis and sensorimotor polyneuropathy. More interestingly, the patients with cortical dysgenesis related to missense TUBB3 mutations also harbor different mutations with distinct consequences on microtubule dynamics and function as well as their interacting proteins [4, 63]. Similar to TUBB2B, only one fetus with TUBB3 mutation (M388V) was reported [4]. Fetopathological analysis reveals microlissencephaly with a severely disorganized cortical plate associated with a disturbed cortical cytoarchitecture and neuronal differentiation. Caudate nucleus, putamen, and pallidum are absent, and the thalami are hypoplastic and crudely shaped. Brainstem and cerebellar hypoplasia is present. Basal ganglia dysmorphism, corpus callosum agenesis, and corticospinal tract hypoplasia also support a generalized defect in axonal guidance.

16.3.2

Other Microtubule (MT) Cytoskeleton-Related Molecules Associated with Human Diseases

a. Loss of function of Filamin A causes periventricular nodular heterotopia in females The importance of the cytoskeleton in neuronal migration is also demonstrated by the contribution of actin and FLNA and its associated proteins, such as Filamin A, because mutations in the FLNA (Filamin A) gene impairs the migration of neurons out of the germinal zone and leads to periventricular nodular heterotopia

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Fig. 16.6 T1 weighted axial (a) and coronal (b) section of MRI scan of the brain of a 26-year-old female with FILA mutation showing isointense tissue with gray matter lining bilaterally the ventricular walls (white arrows) compatible with nodular heterotopia. Note the preserved architecture of the rest of the cerebral structures, including the cerebral cortex and the subcortical white matter

(PNH). PNH occurs most frequently in women as an X-linked trait [71] and constitutes the “classical bilateral PNH.” This form is associated with high rates of prenatal lethality in male fetuses and 50% recurrence risk in the female offspring [72]. Almost 100% of affected families and 25% of sporadic patients, harbor mutations of the filamin A gene (FLNA) [73], which also causes coagulopathy, cardiovascular abnormalities, and Ehlers-Danlos syndrome (joint hypermobility and aortic dilatation in early adulthood) in some patients. Only a few living male patients with PNH owing to FLNA mutations have been reported [74]. Mild missense mutations or mosaic mutations, probably causing limited functional defects of the FLNA protein, account for survival of affected males, who can, in turn, pass their genetic defect to their daughters. Many patients have epileptic seizures, with normal or borderline cognitive level. However, a wide spectrum of clinical presentations and associated features is on record, with a loose correlation between sizes of PNH, structural abnormality of the cortex, and clinical severity. On magnetic resonance imaging (MRI), the PNH is characterized by bilateral confluent nodules of gray matter abnormally located, most frequently in the periventricular region (Fig. 16.6). This form may be associated with hypoplasia of corpus callosum and cerebellum [10, 71, 73]. b. LIS1 and DCX mutations cause most classical lissencephaly, but different patterns of malformation In addition, two major genes have been associated with classical LIS and SBH, DCX (Doublecortin) and LIS1 (or PAFAH1B1, platelet-activating factor acetylhydrolase, isoform Ib) genes both encoding MAPs. Functional studies revealed that the LIS1 protein (also known as PAFAH1B1) is involved in the regulation of the molecular motor complex formed by Dynein and Nudel [22] and that

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Fig. 16.7 Axial T1 weighted brain images obtained in two patients aged 6 months with LIS1-related LIS (a) and DCX-related LIS (b). LIS1-related LIS is more severe in the posterior brain regions (p > a gradient or LIS1-associated pattern, white arrows) (a), whereas DCX-related LIS is more severe in the anterior brain (a > p gradient or DCX-associated pattern, white arrows) (b)

Doublecortin is an MAP that both induces nucleation and stabilizes MTs by linking adjacent tubulin protofilaments [8, 75–78]. The LIS1 gene is responsible for the autosomal form of LIS [9], while the double cortin gene (DCX or XLIS) is X-linked [8, 79]. Although either gene can result in either LIS or SBH, most cases of classical LIS are due to deletions or mutations of LIS1 [80, 81], whereas most cases of subcortical band heterotopia (SBH) are due to mutations of DCX [82]. Clinically, patients with LIS often suffer from moderate-to-severe mental retardation and refractory epilepsy [51, 81, 83]. LIS1-related LIS is more severe in the posterior brain regions (p > a gradient or LIS1-associated pattern, Fig. 16.7a), whereas DCX-related LIS is more severe in the anterior brain (a > p gradient or DCX-associated pattern, (Fig. 16.7b) [9, 80, 81, 84, 85]. LIS can be associated with other abnormalities: enlarged lateral ventricles, absence of claustra and external capsules, malformations of corpus callosum, hypoplasia of pyramidal tracts, and cerebellar malformations. Codeletion of the nearby LIS1 and CRK and YWHAE genes, telomeric to LIS1, underlies Miller–Dieker syndrome, which consists of severe classic LIS and characteristic facial dysmorphism [86, 87]. While hemizygous DCX mutations lead to classic LIS in males (with anteroposterior gradient), the heterozygous DCX mutation in females are associated with

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Fig. 16.8 Axial section in T1 weighted (a) and IR sequence (b) MRI scan of the brain of a 6-year-old girl with DCX mutation. Beneath the cortex, and separated from it by a thin layer of white matter, is present a subcortical laminar (band) heterotopia (white arrows) showing the same signal intensity as the cortex

subcortical band heterotopia (SBH), also known as subcortical laminar heterotopia or double cortex syndrome [8, 79, 88]. SBH is a related disorder characterized by the presence of symmetrical and bilateral bands of heterotopic gray matter located between the ventricular wall and the cortical mantle, and clearly separated from both [56, 89] (Fig. 16.8). DCX mutations are seen in 100% of familial cases and in more than 50% [82, 90] of sporadic, diffuse, or anteriorly predominant band heterotopia cases. Patients with SBH may have mild-to-moderate cognitive abnormalities and frequently have seizures. The brain malformation is often revealed by onset of seizures. In the remaining patients, the investigations that led to the diagnosis are prompted by the presence of developmental delay, sleep disorders, behavioral or learning problems alone or in association [91].

16.3.3

Neurological Diseases Associated with Microtubule Homeostasis-Related Proteins

Interestingly, another protein indirectly related to microtubules homeostasis has been involved in neurological disorders. The HRD syndrome (hypoparathyroidism/mental retardation/facial dysmorphism) or Sanjad-Sakati syndrome, has been associated

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Fig. 16.9 Tubulin-folding and heterodimerization pathway. The tubulin-folding pathway [94, 95] involves a series of molecular chaperones whose function is to facilitate the assembly of the α-/β-tubulin heterodimer. Newly translated α-tubulin (α) and β-tubulin (β) polypeptides are first captured and stabilized by prefoldin (PFD) that acts as a shuttling protein to deliver its bound target protein to the cytosolic chaperonin (CCT) with which it interacts. CCT generates folding intermediates via one or more cycles of ATP binding and hydrolysis. These intermediates then interact with a set of downstream tubulin-specific chaperones (TBCs). Two TBCs (TBCB, TBCE) capture CCT-generated α-tubulin intermediates in which the encapsulating GTP-binding pocket (the Nsite) is already formed producing TBCB/α-tubulin (B/α) and TBCE/α-tubulin (E/α) cocomplexes. Two others (TBCA, TBCD) capture and stabilize CCT-generated β-tubulin intermediates forming TBCA/β-tubulin (A/β) and TBCD/β-tubulin (D/β) cocomplexes. The TBCA/β-tubulin cocomplex can act as a donor of its target protein to TBCD. TBCD/β-tubulin (D/β) and TBCE/α-tubulin (E/α) converge to form a supercomplex with TBCC (C-D/β − E/α). Interaction with TBCC (C) results in the triggering of GTP hydrolysis by β-tubulin. This reaction acts as a switch to signal the release of newly formed GDP-bound α/β heterodimers (light red β-tubulin), which are then competent (following spontaneous exchange with GTP at the E-site) for incorporation into microtubules (MTs). Several cortical dysgeneses-associated mutations in α- or β-tubulin were shown to disrupt this pathway [2, 5, 64]

with a 12-bp deletion in the TBCE gene, encoding the ubiquitous tubulin-specific chaperone (TBC) E [92, 93]. This chaperone belongs to the folding and heterodimerization pathway of α- and β-tubulin. This pathway involves a series of molecular chaperones whose function is to facilitate the assembly of the α-/βtubulin heterodimer [94, 95] (Fig. 16.9). The tubulin heterodimerization can be disrupted by the presence of mutations in the tubulin α1A, β2B or β3 (encoded by TUBA1A, TUBB2B, and TUBB3, respectively), which affects the interaction of tubulin-folding intermediates with different molecular chaperones of the pathway [2, 4, 5]. A recent report states that the TBCE mutation might be causative of a hypoplastic corpus callosum, as observed in six HRD patients [96]. Although this condition is poorly understood and does not seem to present evidence of neuronal migration defects, it supports the idea that tubulins and MT homeostasis are crucial not only for neuronal migration but also for axonal growth. Moreover, despite a rather limited knowledge of the genetic causes of forms of autosomic recessive microcephaly, four genes have been shown to be associated with a significant reduction of head circumference. Mutations in the ASPM gene (Abnormal Spindle-like Microcephaly-associated) are the most common cause [97]

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of microcephaly. This gene is specifically expressed in ventricular zone neuronal progenitors and encodes a protein that interacts with microtubules. The drosophila ASPM mutant is characterized by an impairment of progenitor proliferation, which stops during the metaphase. It has been suggested that the absence of the protein would affect microtubules anchoring to the poles and mitotic spindle assembly. Mutations in CDK5RAP2 (Cyclin-dependent kinase 5 regulatory-associated protein 2) and CENPJ (Centromere-associated protein J) are also responsible for microcephaly cases [97]. They encode two proteins that are involved in the regulation of microtubule assembly from the microtubule-organizing center (MTOC). The last gene, microcephaline (MCPHN1), is unrelated with microtubule-based functions and is rather thought to be part of the DNA reparation machinery [98]. Taken together, these investigations offered compelling evidence that proper microtubule functions are necessary at every step of cortical development: during proliferation and neurogenesis or during postmitotic neuronal migration and differentiation.

16.4 16.4.1

Pathophysiological Mechanisms Molecular Defects Associated with the Mutations

The biochemistry investigations carried out by independent groups revealed that interactions of α- and β-tubulin monomers along with the different molecular chaperones of the folding and heterodimerization pathway are highly sensitive to the presence of mutations, either in TUBA1A, TUBB2B, or TUBB3 [1, 2, 4, 63–65]. A wide array of molecular disruptions was associated with different mutations; the interactions with TBCA, TBCB, and TBCD being the most often affected interactions. These impairments result either in a reduction or in an absence of formed α-/β-tubulin heterodimers (alpha-tubulin 1A, mutants L397P, R264C or beta-tubulin 2B, mutants F265L, S172P) [1, 64, 65]. It is worth mentioning that other mechanisms must be affected by the mutations as some mutants are properly incorporated into microtubules in vitro and upon transfection in cells. These observations strongly suggest that a complete or partial loss of function might be associated with these mutations and that a haploinsufficiency mechanism could explain the neuronal migration disorders. Interestingly, the overexpression of a β-tubulin 2B mutant (T312M), which has not been associated with any molecular abnormality in vitro, could not rescue the radial migration arrest induced by an shRNA in vivo. This in vivo investigation shows that despite the proper incorporation of the mutant into microtubules, the T312M mutation could cause a loss of function of the protein. For those mutations that did not reveal any defects in vitro or in vivo, the consequences might be on the interactions of tubulins with MAPs, motor proteins (kinesins and dyneins), or with some critical effectors of microtubule functions. On the one hand, the S419 α-tubulin 1A residue is indeed located in the H12 helix of the protein, which has been recently shown to be crucial for the control of kinesin processivity [99]. On the other hand, the R402 residue locates between helix 11 and helix 12 that correspond to an interface with

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the MAP Doublecortin (Dcx) and the kinesin KIF1A [76, 100]. Further investigations are needed to explore the alterations in tubulin interactome associated with the different mutations that are silent to date. Microtubule growth and stability alterations associated with different mutations have been recently revealed using in vitro explorations. One alpha-tubulin mutant (P263T) revealed a delayed incorporation into microtubules following a coldinduced, or a nocodazole-induced depolymerization [65]. These observations have been further confirmed using a live assay based on the assessment of velocity of the end-binding protein EB3-GFP. This parameter is indicative of microtubule growth speed, which is reduced upon overexpression of the P263T mutant. These analyses carried out in primary cultured cortical neurons revealed an interesting compartmentalization of the defect for the P263T mutant; the growth rate of microtubules being affected in neurites rather than in the neuronal cell body. These results might reflect a differential composition of microtubules in different cellular compartments (cell body vs. neurites), or the loss of an interaction with a microtubule growth-regulating MAP, which would be specifically expressed in neurites versus soma. The MAP CLIP-170 or the Dis1/XMAP215 family are preferentially located in neurites and stimulate microtubule growth [101–105]. An independent study revealed a similar defect in a yeast system. The R262C mutant of the β3-tubulin induces a stabilization of the microtubule cytoskeleton and a reduced rate of polymerization [63]. These observations underline the possible occurrence of a dominant negative effect for some mutants in addition to the haploinsufficiency observed for many others. Finally, at the extremity of the severity spectrum, several mutations have been shown to be completely reluctant to incorporate into microtubules [1]. To conclude, the different mutations in TUBA1A, TUBB2B, and TUBB3 are translated into a variety of defects: (1) folding and heterodimerization impairments, (2) defective incorporation into microtubules, and (3) alterations of microtubule dynamics and stability.

16.4.2

Cellular Defects Associated with the Mutations

Different investigations carried out on the various forms of agyria, pachygyria, and polymicrogyria associated with mutations in tubulin genes (TUBA1A, TUBB2B, and TUBB3) confirmed the existence of defective neuronal migration mechanisms. Neuropathological studies of TUBA1A- and TUBB2B-mutated fetus revealed neuronal migration defects through the observation of a striking cortical layering disorganization and heterotopic neurons [1, 2, 60, 67, 69]. This was confirmed by investigations of the Tuba1aS140G/+ mouse model and through an in utero RNAi approach. First, Tuba1aS140G/+ -mutant mice displayed hippocampus lamination defects and a discrete lamination defect in layers II/III and IV in the visual, somatosensory, and auditory cortex [2]. Second, the reduction of Tubb2b expression in vivo using shRNA blocked radially migrating neurons in the intermediate zone [1]. Moreover, as described in a previous section, axonal tract organization abnormalities have been reported in patients carrying mutations in the TUBA1A, TUBB2B, and TUBB3 genes

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as well as in the TUBB3R262C/R262C mouse model [1, 2, 4, 60, 61, 66, 69]. Corpus callosum, anterior commissure, subcortical tracts, and internal capsule abnormalities were indeed reported by our group and others, in addition to optic nerve defects reported by Tischfield et al. [6]. More insights have been provided using diffusion tensor imagery on patients with TUBB3 mutations. These MRI investigations permitted to underline the misorientation and mistargeting of internal capsule axonal fibers rather than their absence [4]. On the basis of existing observations and results of animal model investigations, diverse hypotheses can be drawn to explain the occurrence of the abnormal axonal targeting and the abnormal neuronal migration.

16.5

Perspectives and Concluding Remarks

Two nonmutually exclusive phenomenons could explain the abnormal development of subcortical and callosal tracts. The first one is based on the existence of a cell population named corridor cells, which transiently participate in the guidance of corticothalamic and thalamocortical axonal fibers [106]. These cells while migrating from the lateral ganglionic eminence toward the ventromedial area of the telencephalon form a permissive corridor to the growing axons. One could hypothesize that mutations in the tubulin genes of interest in human could affect the migration of a similar population of cells and thus perturb the guidance of axons dependent on this population. Interestingly, a recent study reported the existence of another transient cell population playing an analogous role in the corpus callosum [107]. Thus, we cannot exclude that neuronal migration defects observed in tubulin-related cortical malformations of cortical development could extrinsically contribute to the abnormal guidance of axonal tracts. The second hypothesis involves potential intrinsic mechanisms. Axonal guidance indeed requires properly regulated microtubules, whose polymerization, stabilization, and fasciculation must adequately respond to guidance cues and signals in order to maintain neuronal polarity as well as growth cone displacements and directionality [108]. Interestingly, conditional removal of the serine/threonine kinase 11 (LKB1) in the mouse cortex (Emx1Cre/+ ; Lkb1c/c ) severely disrupts the formation of pyramidal neuron axonal tracts directed toward the internal capsule and corpus callosum [39]. It is worth mentioning that LKB1 underlies microtubule stability through the regulation of MAPs [39, 109, 110]. Further investigations are required to bridge the gap between the observation of neuronal migration defects and the molecular abnormalities affecting the tubulin-folding and heterodimerization pathway and also to explain the phenotypic discrepancies observed between the different forms of tubulin-related malformation of cortical development associated with mutations in TUBA1A, TUBA8, TUBB2B, and TUBB3. In addition to potential subtleties in the spatiotemporal expression of these genes that may partially explain the different phenotypes, one could strongly suspect that the different tubulin isotypes might carry specific intrinsic properties and interaction abilities with given MAPs. Indeed, since the discovery in the 1980s that α- and β-tubulin genes belong to a multigene family, it has been speculated that

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the different isotypes might bear specific intrinsic properties. This concept has been named “multi-tubulin hypothesis” [111–113] and has been highly debated since then. Several studies provided evidences that a differential microtubule composition in tubulin isotypes might partially account for given functional specificities. Drosophila has three β-tubulins (β1 to β3). The β2-tubulin is expressed in male germline where it participates in the formation of mitotic spindle during meiosis of cytosolic microtubules, axonema, and flagellum. The overexpression of the β3-tubulin encoding gene in a β2-tubulin null drosophila does not rescue the defective spindle organization caused by the loss of the β2-tubulin [114, 115]. Only β2-tubulin reexpression can complement the defect, suggesting that certain isotypes carry particular nonredundant functional properties. It is also worth mentioning that the isotypic composition of microtubules has been shown to influence their organization through differential interaction affinities with MAPs [116, 117]. Moreover, it has also been shown that the dynamic properties of the polymer assembly and disassembly can also be altered by its isotypical composition [118–122]. Thus, the modulation of genetic expression of given tubulin isotypes at a given time could contribute to the regulation of microtubule dynamics, which is also operated by MAPs. Genetic expression of tubulin isotypes is indeed submitted to a spatiotemporal regulation during development [1, 60, 123, 124]. For instance, in the absence of MAPs, microtubule in vitro assembly carried out in presence of αβIII tubulin heterodimers is faster than in the presence of αβII or αβIV heterodimers [119]. The presence of mutations in four different tubulin isotypes (as far as we know) is causative of neuronal migration disorders with axonal growth and guidance defects. These mutations have been associated with an array of molecular impairments, from a total absence of incorporation into microtubules to altered dynamic properties. It is possible that the inhibition of the complete heterodimerization and proper incorporation of mutant tubulin into microtubules might perturb the equilibrium of the microtubules isotypic composition and affect their dynamics. This in turn may result in polarization and polarity defects and could explain the occurrence of neuronal migration and axon defects. To conclude, one interesting feature of TUBB2B-related forms of polymicrogyria is the presence of radial glial cell disorganization, which has been observed in several fetal cases [1]; (J. Chelly, unpublished observations). The radial processes are not correctly anchored to the pial basement membrane sporadically throughout the cerebral cortex. The local disruption of radial glial cell architecture is associated with the presence of breaches in the basement membrane that allow migrating neurons to overpass the pia and settle aberrantly within the meninges [1]. It is worth mentioning that these features have been observed in the cortex and cerebellum of the mouse model of GPR56-related bifronto-parietal polymicrogyria (BFPP) and more recently in human fetal cases of BFPP associated with mutations in the G protein-coupled receptor 56, GPR56 [70, 125, 126]. This transmembrane protein is expressed by radial glial cells where it is enriched in the glial-end feet lining the pial basement membrane [70]. All these studies suggest that GPR56 might be involved in the development and the maintenance of the pial basement membrane. In vitro investigations have shown that several mutations identified in humans disrupt the intracellular trafficking of the receptor, which is no longer located in glial-end

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feet and does not participate in the molecular scaffolding sustaining the basement membrane [127]. The radial glia disruption observed in TUBB2- and TUBB3-related malformations of cortical development might result from an impaired intracellular trafficking of transmembrane receptors and adhesion molecule normally present in glia-end feet caused by the alteration of the microtubule cytoskeleton. This feature is specific of TUBB2B and TUBB3 mutations and could be one of the underlying defects leading to polymicrogyria, although it is highly likely that other pathophysiological mechanisms could contribute to the occurrence of neuroanatomical abnormalities in addition to the migration defects that we discussed above.

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Part IV

Intermediate Filaments and Disease

Chapter 17

Spectrins in Human Diseases Marie-Christine Lecomte

Abstract Spectrin is one of the major components of a plasma membrane-associated network, known as the spectrin-based skeleton. The spectrin-dependent skeleton was first identified at the membrane of human erythrocytes, the best known membranes in terms of structure, function, and genetic disorders. Mutations in genes encoding erythroid spectrins cause hemolytic anemia such as hereditary elliptocytosis and hereditary spherocytosis. Present in all mammalian cells, spectrins, and the spectrin-based skeleton are required for organization of specialized membrane domains. Recent advances reveal that these proteins actively participate in cellular localization of a surprisingly diverse set of proteins in many cell types. Recent studies, such as the description of nonerythroid spectrin defects in cerebellar ataxia, lead to an emerging concept for pathogenesis of human disease where failure in cellular localization of membrane proteins results in loss of physiological function.

17.1

Introduction

Plasma membranes of multicellular animals are organized into numerous highly specialized domains which confer to the cell the capacity to participate in diverse and appropriate physiological functions. Their plasma membranes have to be able to: (1) establish precise connections to other cells or to the extra cellular matrix, (2) withstand the forces generated by movement and local environment, (3) send, receive and transmit to the cells interior multiple signals of different nature (electrical signaling, hormones or other molecules), and (4) assume vectorial transport. Therefore, the cells must segregate functionally related membrane proteins within the same domains but also maintain the topographical organization of such domains. These issues require mechanisms to prevent diffusion of membrane-spanning proteins in the fluid phospholipid bilayer. During evolution, a set of proteins emerged to address these functions: they are organized in a scaffold known as the spectrin-based M.-C. Lecomte () INSERM, U665, 75015 Paris, France e-mail: [email protected] Institut National de la Transfusion Sanguine, 75015 Paris, France Université Paris 7-Denis Diderot, 75005 Paris, France

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Fig. 17.1 Visualization of membrane skeleton derived from triton-treated red cell ghosts, a negativestaining electron microscopy reveals the hexagonal lattice of junctional complexes, containing short F-actin and band 4.1, cross-linked by spectrin tetramers skeletons, b assignment of these structural elements is given in the schematic diagram [100]

membrane skeleton (SBMS) which is located at the inner surface of the plasma membranes via links to a number of integral membrane protein (for review see [16]). Such a structure has been first identified in mammalian red blood cells [196]; components of this membrane skeleton have been further characterized in all animal tissues that have been examined so far, indicative of their presence in likely all metazoan cells. Recent progress demonstrates that all the components of this membrane skeleton collaborate in both the establishment and the maintenance of a diverse specialized plasma membrane domain. Spectrins are the major structural components of this membrane-associated scaffold as they constitute the filaments of this network. This review summarizes the new insights into the human diseases related to spectrins.

17.1.1

The Spectrin-Based Skeleton as Defined in Red Blood Cells

The human red blood cell (RBC) is characterized by its discoid shape, its ability to undergo extensive reversible deformations, and its resistance to shear stress during repeated passages though small vessels during its 120-day life span. Such particular shape and mechanistic properties are supported by the spectrin-based membrane skeleton as it has been clearly demonstrated in hereditary hemolytic anemia such as hereditary elliptocytosis (HE) and spherocytosis (HS), in which the underlying molecular defects reside within one of its components. The spectrin-based skeleton was first visualized in electron micrographs of detergent-extracted erythrocytes ([100, 196], Fig. 17.1). It is organized as a polygonal network formed by five to six flexible filaments made of spectrin as tetramers, cross-linked by short actin filaments (40 nm in length made of 14–16 nanomers). Spectrin tetramers, which are the functional units, are made up of two α- and two β-subunits which are encoded by separate genes [48]. The α- and β-chains are associated side to side in an antiparallel manner and intertwine to form rod-like dimers

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Fig. 17.2 Schematic representation of the red cell membrane spectrin-based skeleton [102]

which self-associate head to head to form tetramers: the N-terminal end of each α-subunit associates with the C-terminus of each β-subunit. When stretched, spectrin tetramer filaments can reach approximately 200 nm length. Each extremity of tetramers via β-spectrin binds actin microfilaments, allowing spectrin to form cross-links between actin filaments, thus generating an extended network. The spectrin-actin interaction is modulated by accessory proteins such as protein 4.1, together with dematin (protein 4.9), adducin, tropomyosins, and tropomodulin (Fig. 17.2, [102]). Their functions are to stabilize the actin-spectrin complex, to maintain actin filament length (adducin acts as a capping protein), and to bind the spectrin-based network to the transmembrane proteins (the glycophorin C, the anionic exchanger, AE1/band 3) via adapter proteins (protein p55 and protein 4.2, [86, 102]). Another major binding site to the membrane is mediated via ankyrin, which binds to β-spectrin and the anionic exchanger, AE1/band 3 [75]. The Rh/RhAG-ankyrin complex can be also a link between the red cell membrane and the spectrin-based skeleton, the disruption of which might result in the stomato-spherocytosis typical of Rh null red cells [127]. All these interactions are highly regulated by protein posttranslational modifications, mainly phosphorylation. Spectrins interact also directly with phospholipids such as phosphatidylserine, a component actively confined to the inner leaflet of the lipid bilayer [164, 36, 173].

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The phosphatidylserine-binding sites in β-spectrin are grouped in close proximity to the attachment sites for both ankyrin and 4.1, the proteins engaged in spectrin links to the membrane [4].

17.1.2

The Diversity of the Spectrins and Spectrin-Based Skeletons in Nucleated Cells

The components of the erythrocyte spectrin-based skeleton are, in fact, products of closely related gene families which have widespread distribution in all mammal cells. Isoforms of the proteins that form the erythrocyte spectrin-based skeleton are detected in all metazoan cells that have been examined so far. These observations led to the assumption of the existence of a spectrin-based skeleton as a ubiquitous protein scaffold under all metazoan plasma membranes. Although the principles of the basic organization established in erythrocytes could be likely applied in nonerythroid tissues, the regulation of gene expression, the protein-protein interactions and functions are considerably more diverse. In contrast to the red blood cell, several kinds of spectrin-based skeletons exist in nonerythroid cells, but their composition, organization, and function are not so well characterized owing to the diversity of the genes (for example, two genes coding for α-spectrin [αI and αII], five for β-spectrin [βI–βV], three for ankyrin [Ankyrin-R, -G and -B], four for protein 4.1) and the complexity of the gene products (at least 200 proteins, [16]). The diversity of the gene products can be correlated to the structural complexity of the nucleated cell. These multiple protein isoforms distribute to various intracellular locations besides the plasma membrane, such as the Golgi apparatus, vesicles, the endoplasmic reticulum, and the nucleus. These diverse cell locations indicate that the spectrin-based skeleton fulfils a variety of functions at multiple sites in nucleated cells. Therefore, in mammals, the different spectrin isoforms originate by extensive mRNA splicing from seven genes (Table 17.1). Two genes, SPTA1 and SPTAN1, encode αI- and αII-spectrin subunits, respectively. The SPTA gene seems to have a restricted expression pattern, encoding for the well-known αI1-spectrin isoform in RBC and also for a not well-defined isoform (αI ∗ ) in brain. The αII isoform expressed in human RBC contains 2,428 residues given a calculated MW of 280 kDa [156]. In contrast, the SPTNA gene codes for several αII-spectrin isoforms present in all nonerythroid cells, resulting from three alternative splice variants (corresponding to 60, 15, and 18 nt insert in spectrin repeats 9, 14, and 20, respectively, [26, 121, 200]). Five genes code for β-spectrins: four “conventional” β genes, SPTB, SPTBN1, SPTBN2, SPTBN4, encoding the βI–βIV spectrins, respectively, and one gene, SPTBN5 encoding one large βV-spectrin (β-Heavy, [191, 192]). The SPTB gene is predominantly expressed in erythrocytes, coding for the βI1 isoform (2,137 residues with a calculated MW of 246 kDa), but also for isoforms in brain and muscle such as the βI2 isoform presenting a large C-terminal end including a PH domain. βI-spectrin was also detected in lymphocytes [148]. The βII-spectrins are widely expressed in all nucleated cells. βIII-spectrin likely constitutes a major component

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Table 17.1 Spectrin genes and subunits Cenorhabditis elegans

Drosophila melanogaster

Homosapiens

Subunit

Gene

Subunit

Gene

Subunit

Gene

Chromosome

α

Spc-1

α

I(3)dre3

β-G

Unc-70/bgs-1

β-G

b-Spec

β-H

Sma-1

β-H

karst

αI αII βI βII βIII βIV βV

SPTA1 SPTAN1 SPTB SPTBN1 SPTBN2 SPTBN4 SPTBN5

1q21-q23 9q33-q34 14q22-q23.2 2q21 11q13 19q13.13 15q21

of the Golgi and vesicular membrane skeletons (actively transported vesicles in the secretory and endocytic pathway), but βIII-spectrin was also found to be associated with the plasma membrane in neurons and epithelial cells [72, 170]). βIV-spectrins are most abundantly expressed in neurons (axon, initial segment, nodes of Ranvier) and pancreatic islets [18]. Five isoforms have been detected with isoform 5 located in the nucleus [178]. βV-spectrin expressed at low levels in many tissues is detected prominently in outer segments of photoreceptor rods and cones, at the basolateral membrane of gastric epithelial cells and in the outer hair cell (OHC, [96, 169]). The expression of the diverse isoforms is regulated in a complex tissue- and time-specific manner. Invertebrates have a smaller repertoire. The Cenorhabditis elegans and Drosophila melanogaster genomes include a single gene coding for an α-spectrin close to the mammalian αII-spectrin (spc-1 and I(3) dre3, respectively, [23, 38]) and two genes coding for β-spectrin, one codes for a protein βG resembling the mammalian βIIspectrin referred to as “conventional β spectrin” (Unc-70/bgs-1 and β-Spc, [24]) and the other one gene (sma1 in C. elegans and karst in D. melanogaster) encodes βH-spectrin (βHeavy similar to mammal βV 430 kDa, [37, 112]). There is greater sequence conservation between spectrins from Drosophila and nonerythroid spectrin than between the erythroid and nonerythroid forms within mammals. For example, the α-spectrin from fly has 64% identity to the human αII-spectrin and the βG− subunit of 265 kDa corresponds with 55.8% identity to the product of the human βII gene. Sequence analyses suggest that the erythroid spectrin genes arose during vertebrate evolution and some of the sequence changes might correspond to neofunctionalization of the erythroid spectrin genes [5, 6, 158].

17.1.3

The Protein Architecture of Spectrins Has Been Conserved From Invertebrates to Humans

Despite the multigenic origin, each spectrin subunit has a common basic architecture: each chain consists of an elongated backbone made up of a succession of repetitive units (roughly 106 amino acids long) defined as spectrin repeats, flanked by nonhomologous N- and C-terminal sequences [156, 166, 192].

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Both α-subunits can be divided into 22 segments, among them 20 are spectrin repeats (designed as α1–α20 repeat), the tenth segment is an SH3 domain and is included in the α9-repeat, the last COOH segment (the 22nd segment) consists of two EF-hands motifs (40 residues motifs structurally related to calmodulin and involved in calcium binding, [103, 174]). The β-spectrins have a central domain made of 17 spectrin repeats (β1–β17), except the βV/βH isoform which is made of 30 spectrin repeats. The N-terminal domain of β-spectrins contains an actin-binding site (defined between residues A47 and K186 within βI-spectrin), which comprises two calponin homology (CH) domains in a tandem arrangement [8]. This region presents the greatest homology among spectrins.

17.1.3.1 The Spectrin Repeat The 3-D structure of the spectrin repeat established by X-ray crystallography [194] and NMR [104, 137, 138] confirm the predicted model based on both protein sequence and circular dichroism studies indicating a high α-helicity content [193]: the 106 amino-acid spectrin repeat folds in a triple α-helical coiled-coiled structure made of three helices (A, B, and C) interconnected by two nonhelical loops (AB and BC). The two A and B helices form up in an antiparallel fashion and build a groove in which the C helix resides. These coiled-coil triple helical bundles are linked through interconnection between C helix of one repeat and A helix of the following one to form a continuous CA helix. Sequence homology between the repeats is only about 20%, but highly conserved residues like leucine at position 26 and tryptophan at positions 22 and 45 contribute through hydrophobic and electrostatic interactions to the particular triple helical structure of each repeat (about 5-nm long and 2-nm across). The connecting BC and AB loops are sites of major differences between the repeats, differing in length and conformation. In situ analysis of the spectrin dimers has shown that its length can vary between 30 and 100 nm [162]. Such flexibility in length might occur through conformational rearrangements of the BC-connecting loop [63, 118]. Many spectrin repeats contain additional sequences (between 8 and 60 residues) which are integrated into the repeat units, respecting likely the triple helical structure. These sequences, but also the repeats themselves, are sites for numerous protein interactions.

17.1.3.2 Spectrins Present Numerous Binding Sites for a Broad Diversity of Partners Spectrins appear as large platforms of interactions, exhibiting numerous binding sites given by the nonhomologous sequences specific to the different isoform, but also by the spectrin repeats themselves.

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The first set of interactions which are conserved in all spectrins, mediates the formation of the spectrin filament, that is, the formation of the dimer and the tetramer, involving the site of “nucleation” and the site of “tetramerization,” respectively. The lateral dimer assembly between α- and β-subunits begins at a specific nucleation site, near the actin-binding domain, involving the four last spectrin repeats of the αchain (α17–α20) and the first spectrin repeats of the β-chain (β1–β4, [98, 167, 180]). The α19 and α20 repeats as well as the β1 and β2 repeats are somewhat atypical, having an eight-residue sequence insertion and exhibiting strong homology with αactinin. These repeats zip together to the head of the molecule. Then, heterodimers self-associate head-to-head to form tetramers by two α–β interactions. These α–β interactions lead to the reconstitution of a complete triple helical repeat as present along the spectrin (Sp) molecule [66, 75, 87, 175]. Indeed, the N-end of α-chain begins by an isolated C helix (followed by 20 complete repeat units, α1–α20), when the last repeat unit, β17 at the C-end of β-chain, is incomplete consisting only in A and B helices, followed by a nonconsensus structure, which can include a PH domain according to the isoforms [106, 183]. The two A and B helices of the last repeat β17 interact with the first helix C of the α-chain N-terminal domain. The first full triple helix adjacent to the partial repeats modulates the affinity of interaction. While the spectrin dimer-dimer interaction was long though to be static, recent studies have revealed that dissociation of spectrin tetramers can be induced by membrane deformation [3]. A second group of interaction sites is involved in the formation of the skeleton such as the actin-binding site, the 4.1-binding site, and sites which participate in the attachment of the spectrin tetramers to the membrane such as the ankyrin-binding site located on β-spectrin (within β15 repeat, [75, 76, 82]).These sites are thought to be present in all spectrin tetramers. A third set comprises sites, which mediate interactions with specific partners, contributing to definite functions of the particular spectrin isoform, such as the SH3 domain present in α9 repeat [19, 22, 153, 202].

17.1.4

Spectrins for Which Functions?

The functions clearly determined up to date for the spectrin network emerge from human mutations associated with diseases as well as from animal models. Numerous studies on red cells, particularly those in hereditary hemolytic anemia, have clearly established its importance for supporting cell shape and for maintaining cell membrane integrity. In nucleated cells, the spectrin-based skeleton has been shown to participate in the stabilization or activation of several specialized membrane proteins. One consistent feature observed when spectrin or its binding partner ankyrin are lost or defective, is a failure of interacting membrane proteins to accumulate at the appropriate site. The diverse cellular environments and the multiple protein interactions put the spectrin-based skeleton in the context with numerous different physiological pathways which include cell proliferation and differentiation.

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Spectrins in Hemolytic Anemia Spectrin-Based Skeleton Confers Membrane Stability in RBC

The best understood function of the spectrin-based skeleton (SBMS) is to confer integrity and flexibility to red cell membranes [184]. To assume their function, erythrocytes need a highly flexible sheathing in order to edge through small blood vessels, a characteristic which it owes to the firmly attached SBMS with its extremely flexible spectrin molecules at the inner surface of the membrane. An intact SBMS is critical for the structural integrity of the plasma membrane in erythrocytes. Defects in its components are associated with red cell fragility, fragmentation and premature destruction, and are clinically expressed as hemolytic anemia, such as hereditary elliptocytosis (HE), pyropoikylocytosis (HPP), and spherocytosis (HS). In HE and HPP diseases, most of the molecular defects described so far occur essentially within components concerned in horizontal interactions, namely Sp dimers and protein 4.1. Sp defects have been of interest in regard to the analysis of the structure-function relationship and have provided information on the definition of the structure involved in some binding sites, in particular, the self-association site. Conversely, molecular defects described in HS involve proteins concerned in vertical interactions, such as deficiencies in Sp or ankyrin or protein band 3 or protein 4.2.

17.2.2

Hereditary Elliptocytosis is Often Related to Defects in Spectrin Which Alter the Formation of Tetramer

Hereditary elliptocytosis (HE) is a relatively common autosomal dominant disorder characterized by the presence of elliptocytic-shaped erythrocytes on peripheral blood smear. HE has a worldwide distribution, with an incidence ranging from one in 5,000 to 10,000 among Caucasians, but HE appears more common in malaria endemic regions (prevalence approaching 2% in West Africa, [35, 61, 91]). However, HE differs from the Southeast Asian Ovalocytosis (SAO) which is very common in malaria areas (and related to AE1/band 3 defect, [43, 83, 115]). HE is heterogeneous in terms of clinical severity, red cell morphology abnormalities, and underlying molecular defects. Its clinical presentation ranges from an asymptomatic condition to severe hemolytic anemia characterized by the occurrence of red cell fragmentation and poikylocytosis (also defined as HPP); a few cases of fatal hydrops fetalis were also reported [54]. Most of the patients present mild HE, showing compensated hemolysis. Mild-to-severe anemia is observed in patients either homozygous for HE variants or compound heterozygous HE variants [46]. HPP which exhibits great membrane fragility (numerous poikylocytes on blood smear) and thermal sensitivity of red cells [197] was considered as a distinct clinical entity, but it corresponds in fact to a severe picture of HE [90]. The increase in severity is usually accompanied by the replacement of elliptocytes by poikylocytes (cells of all sizes and shapes, reflecting a great fragility of red cells and their fragmentation).

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A common feature of all forms of HE is a mechanically unstable membrane: in most HE, the molecular defect described so far occurs essentially within components involved in horizontal linkages, namely with the Sp dimer-dimer interaction or spectrin-actin-protein 4.1R junction complex. HE results from mutations altering the SPTA1, the SPTB, and the EL1 genes that encode the Sp αI-chain, the Sp βI chain, and the protein 4.1, respectively. Mutations in spectrins are the most common cause of HE, accounting for approximately 90% of cases. Almost all HE cases related to a spectrin molecular defect present a more or less pronounced deficiency of the ability of the Sp spectrin dimer to self-associate into tetramers, indicating that Sp spectrin dimer self-association is crucial for the mechanical properties of the membrane. This defect can be evaluated in vitro through the percentage of Sp dimers present in the 4◦ C crude spectrin extracts. The increased percentage of Sp dimer in the 4◦ C extracts ranged from 8 to 62% in heterozygous HE and from 29 to 82% in homozygous HE compared to 3–5% in controls [93]. This functional defect has been related to numerous mutations of either αI- or βI-spectrins, the SPTA gene being more affected than SPB: more than 25 α-spectrin mutations have been reported (Table 17.2). Most of these HE mutations occur at or near the spectrin repeats involved in the formation of the tetramer, either in the first C helix and α1 repeat of α-spectrin or in the incomplete β17 repeat, they are also distributed along the first six repeat units of αI-spectrin. Most of them are missense mutations, but some mutations lead to truncations of various sizes within the C-end of the β-chain. Importantly, four mutations have been observed in codon 28 within α-spectrin suggesting that the CpG dinucleotide is a hotspot for mutation. Attempts were made to establish the possible relationship between the different mutations and the various clinical expressions of the disease. However, it appeared that the same mutation could result in different clinical features, even in HE subjects from the same family and each clinical phenotype does not necessarily correspond to a specific mutation, but each defect occurs in more than one clinical phenotype [28, 93]. The main correlation was found between the intensity of the Sp self-association defect and the severity of clinical phenotype [93]. In fact, the severity of clinical expression, reflected by the intensity of hemolysis as well as the decrease in mechanical stability of the membrane, is proportional to the extent of the self-association defect. Asymptomatic HE showed the lowest Sp self-association defect as indicated by percentages of Sp dimer below 35%. Severe hemolytic anemia, requiring blood transfusions and most often splenectomy, was always observed in HE patients with a proportion of Sp dimer greater than 43–45%. Between these extreme clinical phenotypes (asymptomatic and highly hemolytic diseases), HE patients with more or less compensated chronic hemolysis showed intermediate percentages of Sp dimer in membranes (from 30 to 40%). The severity of the self-association defect appears to depend on the location of the molecular defect in relation to the self-association site as it was first defined by Tse et al. [175]. It was generally more pronounced when the mutation is located within the helices directly implicated in the formation of the tetramer, (the first C helix of α-spectrin or the last two A and B helices of the β-chain, [126] and when the mutation

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Table 17.2 Spectrin mutations in hereditary elliptocytosis Designation

Nucleotide change

Amino-acid change

Repeat

References

CAT-CGT ATC-AGC CGT-CAT CGT-TGT CGT-AGT CGT-CTT GTG-GCG CGG-TGG CGG-TGG AGG-ACG AGT-AGG GGT-GTT CTT-TTT AAG-AGG GGT-GAT CTG-CCG CTG-CCG TCC-CCC CAG-CCG GAC-GAA

I24T I24S R28H R28C R28S R28L V31A R34W R41W R45T R45S G46V L49F K48R G151D L207P L260P S261P Q471P D791E

First C helix First C helix First C helix First C helix First C helix First C helix First C helix First C helix First C helix First C helix First C helix First C helix First C helix First C helix Repeat 1 Repeat 2 Repeat 3 Repeat 3 Repeat 4 Repeat 8

[55] [136] [56] [29] [29, 42] [29, 42] [107] [143] [123] [144] [92] [124] [124] [42] [21] [52] [155] [155] [155] [1]

L154 dup 49 del H471del

Repeat 1 Repeat 2 Repeat 5

[154] [67] [53]

363–371 del 822–862 del 926–958 del

Repeat 4 Repeat 8 Repeat 8

[7] [2] [44]

S2019P A2023V W2024R L2025R A2023P W2061R R2064P E2069stop GLN1946stop

Repeat 17 Repeat 17 Repeat 17 Repeat 17 Repeat 17 Repeat 17 Repeat 17 Repeat 17 Repeat 16

[54] [136] [136] [107] [175] [49] [150] [108] [108]

2041 2053 2058 2044

Repeat 17 Repeat 17 Repeat 17 Repeat 17

[58] [188] [81] [176]

Repeat 17 Repeat 17 Repeat 17 Repeat 17 Repeat 17

[78] [9] [195] [51] [57]

SPTA1 mutations Missense mutation

Sp Genova Sp Tunis Sp Anastasia Sp Culoz Sp Lyon Sp Ponte deSor

Sp Jendouba Insertion/deletion

TTCins Sp Dayton INS ME Sp Alexandria CGTcatCG Splicing site mutation Sp Sfax IVS8 A-G Sp Oran IVS17, G-A Sp St Claude IVS19, T-G-13 SPTB mutations Missense and nonsense mutations Sp Providence TCT-CCT GCG-GTG TGG-AGG CTG-CGG GCT-CCT TGG-AGG Sp Cosenza CGC-CCC Sp Nagoya GAG-TAG CAG-TAG Insertion/deletion Sp Tandil Del7nt Sp Napoli Del8nt 6255–6262 Sp Tokyo Del1nt Sp Nice Ins2nt Splicing site mutation Sp Prague IVS29 G-C Sp Campinas IVS30 G-A nt + 1 Sp Gottingen IVS30 T-A Sp Le Puy IVS30 A-G Sp Rouen IVS31 G-T

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affects residues involved in the stability of the spectrin repeat [45, 80, 88, 134, 198]. A second parameter appears to affect the severity of the functional defect: the percentage of mutated α-spectrin present in the membrane, the expression of the αHE allele being modulated by the presence in trans of a low expression allele (such as the αLELY allele, for Low Expression Lyon). The αLELY allele contains a C-T mutation at nucleotide-12 of intron 45 associated with partial inframe skipping of exon 46 and leading to deletion of residues 2177–2182 within α20 spectrin repeat, which participates in α–β spectrin nucleation. This deletion alters the dimer assembly [11, 187, 189, 190]. The allele αLELY occurs in various ethnic groups investigated so far (Caucasians, African Blacks, Japanese, and Chinese) with a fairly uniform incidence between 20 and 30% [109]. To conclude, the severity of both the self-association defect and clinical expression in HE related to β-spectrin mutations mainly depends on the location and the kind of the mutation. Conversely, the severity of the disease in HE related to α-spectrin mutations depends on the presence of low expression α-allele in trans of the αHE allele. However, a set of HE mutations within α-spectrin did not produce any deleterious effect on the tetramer formation; the damaging mechanism related to these mutations is not yet elucidated. These mutations are located at the center of αI-spectrin around the SH3 domain: Sp Jendouba, Sp Oran, Sp St Claude. Sp St Claude allele is characterized by a splicing mutation that leads in part to a defective α-spectrin chain that cannot be assembled in the membrane. The patient homozygous for the Sp St Claude allele experienced a severe poikilocytic hemolytic anemia. Molecular epidemiologic studies indicated that HE related to α-spectrin defects have a higher incidence in black ethnic populations than in Caucasian populations. Indeed, it has been observed that some mutations are largely distributed in Africa in contrast to mutations characterized in Caucasians HE which appear more or less silent. Therefore, the Leu154 dup has a broad diffusion in WestAfrica, in NorthAfrica as well as in southern Italy, the West Indies, and among the African-American population. The Leu260Pro mutation often associated in cis with particular polymorphisms, defining the Sp αNigerian allele, is frequent in southern Benin and Togo among two ethnic groups (Fon and Yoruba). The Leu207Pro mutation was observed in other ethnic groups from West Africa in the Dendi people. This mutation is present in the low-expressed αI-spectrin allele (αLely allele), defining the Sp αSt Louis allele. Sp St Claude allele was identified in 3% of asymptomatic individuals from Benin [44, 61]. The broad diffusion of these αHE alleles raises the question of whether these spectrin mutations provide some protection against malaria [35]. None of the above spectrin mutations, however, reaches such a high frequency as, for instance, the protein band 3 mutation accounting for ovalocytosis in Southeast Asia (up to 35% in particular areas). Certain elliptocytogenic alleles of spectrin could be supplementary genetic factors of malaria resistance in vitro [35]. It is not known, however, whether this potential resistance is expressed in vivo. Most of the parasite proteins exported to the RBC membrane appear to be distributed around the spectrin-based skeleton and direct molecular interactions between host and parasite proteins begin to be characterized [71, 130, 139–141]. All these data suggest that the relationships between the parasite and the spectrin-based skeleton should be explored more closely.

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Spectrin Deficiencies in Hereditary Spherocytosis

Hereditary spherocytosis (HS) is a common inherited hemolytic anemia with a worldwide distribution but it appears more common in individuals of Northern European ancestry affecting 1–5 persons in 10,000 individuals in Europe [40, 47]. HS has also been frequently described in Japan [125]. Hereditary spherocytosis has mainly an autosomal dominant inheritance (roughly 75% of cases); the remaining cases are likely recessive cases or result from de novo mutations. The clinical manifestations of HS vary widely: HS is characterized by a chronic hemolysis with a broad spectrum of clinical severity ranging from asymptomatic to severe anemia requiring splenectomy, and rare cases of hydrops fetalis were observed. In the common forms, the symptoms appear 1 or 2 weeks following birth. Typical HS consists of evidence of hemolysis with anemia, jaundice, reticulocytosis, gallstones, splenomegaly, as well as spherocytes on peripheral blood smear. Splenectomy significantly reduced the severity of anemia by increasing the circulatory lifespan of spherocytes [145]. HS is also characterized by the heterogeneity of the causal molecular defect. The analysis of the RBC membrane proteins reveal several subsets of HS characterized by isolated or combined protein deficiencies. These protein deficiencies result from mutations affecting either ankyrin (the most common cause of HS in Northern European populations accounting for 50–60% of cases) or AE1/band 3 (15–20%) or spectrin (15–30%) or protein 4.2 (common in Japan and Korea) and Rh complex [94, 110, 152]. The underlying mutations are very heterogeneous, they are not largely distributed, and nearly every family has a unique mutation. They are widely distributed along the affected genes and include frame-shift, nonsense or missense mutations, consensus splice site mutations, and large genomic deletions. Such molecular defects alter the stability of the transcriptional message, or the stability of the protein, or lead to the disruption of specific functions of the protein. The result may be a total silencing of one allele (null allele), a partial disabling of one gene with reduced expression of a normal protein, or production of a dysfunctional peptide (low express allele). These adverse effects ultimately result in the deficiency of the affected protein and their associated proteins. Despite the heterogeneity of the underlying molecular defects, a common feature of HS is a loss of membrane surface leading to a spherocytic phenotype and increased osmotic fragility. The severity of the disease seems directly related to the extent of membrane surface loss [25]. The common mechanistic basis for membrane loss in HS results from defective anchoring of the spectrin-based skeleton to the membrane and involves the disruption of vertical protein interactions. The disruption of vertical links such as spectrin-ankyrin, ankyrin-band 3, band 3-protein 4.2 interactions, which occur via the failing of one of its components, is proposed as the basis for the uncoupling of the membrane from its supportive skeleton, resulting in the shedding of membrane vesicles and surface area deficiency in HS. Hereditary spherocytosis related to SPTA1 gene defects (αI-spectrin) shows a recessive inheritance pattern and appeared to be rare. They are often associated with

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a severe phenotype; the spectrin deficiency could reach up to 40%. Recessive HS is most often due to compound heterozygosity of defects in α-spectrin, but also paired with ankyrin and protein 4.2 [117]. Common combinations include a low expression allele, facing in trans a null allele or a missense mutation. Only a few productiondefective alleles of the αI-spectrin gene SPTNA1 have been characterized: (1) The αLEPRA allele (Low Expressed Prague) presents a C→T transition at position −90 of intron 30, IVS30, causing an alternative aberrant splicing, frame-shift, and premature termination of translation. The incidence of αLEPRA allele was estimated around 5% among Caucasian populations [32, 34, 177, 186]; (2) The αLELY allele which is also observed in severe HE (HPP); (3) the St Louis allele contains a G to A substitution at position + 5 of the donor consensus splice site of intron 22 resulting in insertion of intronic fragments with an in-frame premature termination codon, St Louis allele is also observed in severe HE [30]; (4) The null αLELY-Bicetre allele contains an additional G-to-A mutation in the last position of exon 51; and (5) The αPRAGUE allele shows an A to G in the penultimate of intron 36, IVS36, leading to exon 37 skipping and frame shift. The particular feature of recessive HS results from a combination of null alleles, facing in trans a low expressed allele could be related to the excess of α-spectrin synthesis during erythropoiesis. Indeed, pulse-labeling studies in cultured erythroid cell lines have established that α-spectrin is synthesized by threefold excess compared to β-spectrin synthesis. Therefore, a weak allele or null allele of the SPTA1 gene in the heterozygous state will have no consequence on the spectrin amount at the membrane. Inversely, homozygosity for null SPTA1 alleles is likely to be lethal. In contrast, β-spectrin is predicted to play a regulatory role in the heterodimer assembly onto the membrane, and therefore β-spectrin defects support an autosomal dominant transmission hypothesis. Animal models give further insight into the pathogenetic consequences of membrane protein defects as well as the causes of the variability of disease severity. Several spontaneous allelic mutations in the αI-spectrin gene, Spna1, have been identified in mice causing severe hemolytic anemia with spectrin deficiency [151, 182]. In (sph), mice spherocytosis is due to a single base deletion in repeat 5 causing a frame shift and premature termination. No α-spectrin could be detected in the membrane whereas some β-spectrin is still present. In sph(2BC), a splicing mutation initiates the skipping of exon 41 and premature protein termination before the site required for dimerization of α-spectrin with β-spectrin. In sph(Ihj), a premature stop codon occurs (Q1853Stop) in α18 repeat. In sph(3J), a missense mutation (H2012Y) in α19 repeat introduces a cryptic splice site resulting in premature termination of translation. Both mutations result in markedly reduced membrane spectrin content, decreased band 3, and absence of β-adducin. In sph(1J), a nonsense mutation in exon 52 eliminates the last COOH-terminal 13 residues, facing the 4.1/actin-binding region at the junctional complex providing new evidence that this 13 amino-acid segment at the COOH-terminus of α-spectrin is crucial to the stability of the junctional complex. In sph(4J), a missense mutation occurs in the C-terminal EF hand domain (C2384Y). Notably, a severe hemolytic anemia occurs despite a weak decreased membrane spectrin with normal band 3 content and reduced β-adducin. The

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Table 17.3 Beta-spectrin mutations in hereditary spherocytosis Designation

Nucleotide change

Amino acid change

Reference

Nonsense mutation Sp Baltimore Sp Tabor Sp Alger Anonymous

CAG-TAG CAG-TAG CAG-TAG AGA-TAG

Q845stop Q1946stop Q514stop R1756stop

[107] [107] [33] [105]

Deletion/insertion Sp Ostrava Sp Bicetre Sp Sta Barbara Sp Houston Sp Sao PauloII Sp Philadelphia Sp Bergen Sp Dhuram Anonymous Anonymous

699delT 1328–1335del 1912delC 2872delA 4176delC 1862insA 2351insA 4473-4842del 2344delT 4508insA

201-frameshift 443-frameshift 638-frameshift frameshift 1391-frameshift 589-frameshift 783-frameshift L1492-K1614 deletion 781-frameshift 1502-frameshif

[107] [33] [50] [70] [13] [70] [70] [68] [105] [105]

Splicing site mutation Sp Guemene-Penfao Sp Bari Sp Winston-Salem Anonymous Anonymous

IVS 3-1C IVS 16 IVS 17 + 1A Sp IVS22nt-4G > A Sp IVS20nt-2A > G

100-frameshift frameshift 935-frameshift frameshift frameshift

[59] [146] [69] [105] [105]

Missense mutation Sp Promissao Sp Atlanta Anonymous Sp Kissimmee Sp Oakland Sp Columbus Sp Birmingham Sp Sao Paulo

ATG-GTG TGG-GGG c567C TGG-CGG ATC-GTC CCT-TCT CGC-TGC GCG-GTG

M1V W182G G189A W202R I220V P1227S R1684C A1884V

[10] [70] [14] [70] [70] [70] [50]

severe phenotype in sph(4J) indicates that the highly conserved cysteine residue at the C-terminus of α-spectrin participates in critical interactions for membrane stability. The data reinforce the notion that a membrane bridge in addition to the classic protein 4.1-p55-glycophorin C linkage exists at the RBC junctional complex that involves interactions between spectrin, adducin, and band 3. In contrast to HS-related SPTA1 gene, many more mutations (more than 25 different mutations) have been elucidated in the SPTB gene. They include initiation codon disruption, frame shift, nonsense and missense mutations, gene deletions, and splicing defects. Most of these mutations are associated with a phenotype of mildto-serious autosomal dominant form [10, 12, 14, 70, 105]. Not infrequently, SPTB mutations appear de novo [116]. Most of the mutations within the SPTB gene result in the silencing of the mutant allele and are distributed along the gene (Table 17.3). Some HS present missense mutations—such as Sp Atlanta (W182G) and Sp Kissimmee (W202R)—located in the highly conserved region of β-spectrin involved in

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the interaction with protein 4.1 and actin. These mutations alter likely the binding of spectrin to actin as showed by in vitro studies. Spectrin Dhuram has a deletion within the ankyrin-binding site altering spectrin attachment to the membrane. Any of these mutations have been observed so far in humans with a homozygous or composite heterozygous status. It is possible that other β-spectrin mutations leading to the reduced expression of a normal protein, or leading to the production of a dysfunctional protein, may have less dramatic consequences and may exhibit a recessive pattern of inheritance.

17.3

Spectrin Mutations in Brain Disease

Spectrin is particularly expressed in brain, comprising between 2 and 3% of total protein [31]. In neurons, spectrin is, among other sites, localized to synapses. Spectrin was shown to be involved in the connection of synaptic vesicles with the presynaptic neuronal membrane [62], probably through interaction with two synaptic vesiclespecific proteins, synapsin [163] and MUNC13–1 [157]. Isoform βII1 spectrin was demonstrated to be an essential component of the synaptic transmission, participating in the adhering of synaptic vesicles within nerve terminals [165]. Drosophila null mutants lacking α- or β-spectrin showed severely disrupted neurotransmission and wrong-located classes of synaptic proteins [41]. The spectrin-based skeleton was hence suggested to determine the localization of the synaptic proteins CSP (Cystein string protein) and DLG (Disc-large) in neurons. Spectrins interact with “Shank” proteins [20] in postsynaptic densities, the site of neurotransmitter reception in excitatory synapses in the brain. The ankyrin-repeat containing Shank plays a central role in organizing the subsynaptic scaffold through interaction with several synaptic proteins. However, although C. elegans with disrupted βG -spectrin (unc-70) showed a defective pattern of axon outgrowth, the animals had normally clustered synaptic vesicles at nerve terminals and a functioning secretory pathway [64, 65, 122].

17.3.1

Spectrin Defects in Spinocerebellar Ataxias

Autosomal dominant spinocerebellar ataxias (SCAs) belong to a heterogeneous group of neurodegenerative disorders usually of adult age. SCAs display classic cerebellar signs and most commonly brainstem dysfunction, resulting from degeneration of the cerebellum, spinal cord, and brainstem. They are characterized by uncoordinated gait, limb and eye movement abnormalities, slurred speech, and swallowing difficulties. Slowly progressive ataxia accompanied by cerebellar degeneration is often of genetic origin. Nearly 30 distinct genetic causes of SCA are known, numbered chronologically in order of discovery (SCA1–30). The most common form of mutation that causes SCA is the expansion of trinucleotide (CAG) repeat encoding polyglutamine as found in SCA1, SCA2, Machado-Joseph disease, SCA6, SCA7, SCA17, and DRPLA. Other dynamic mutations, such as a noncoding one

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(CAG/CTG) expansion has been identified as the cause of SCA8 and SCA12, and a pentanuclotide (ATTCT) expansion in SCA10. In addition to these dynamic mutations, static mutations, such as missense mutations and deletions, have been identified to cause SCA5, SCA11, SCA13, SCA14, SCA15, and SCA27. Among them, mutations in SPTBN2 gene, coding for βIII-spectrin (2,390 residues) have been recently recognized as the cause of SCA5 in three large families of American and European origin [74]. A 39-bp deletion of βIII-spectrin gene (exon 12) that causes an in-frame 13-residue deletion (E532-M544del) was found in the American family. This American family descends from the paternal grandparents of President Lincoln. Similar to the American family, a 15-bp deletion in exon 14 leading to an in-frame deletion (1886–1900del; L629-R634delinsW) was found in a French family. This deletion does not disrupt the open reading frame but introduces a tryptophan. Both deletions are located within the β3 spectrin repeat, which is included in the initial interaction between α- and β-spectrin to constitute the dimer (in the nucleation site). In a German family, a T to C transition in exon 7 causes a leucine to proline change (L253P) in the calponin homology domain containing the actin-/Arp1-binding site. This region is highly conserved, with the L253 residue found in all five human β-spectrins as well as in mammals and flies. βIII-spectrin is highly expressed in Purkinje cells [131, 170]. βIII-spectrin participates in the membrane stabilization of membrane proteins such as the Purkinje cell-specific glutamate transporter EAAT4 [77]. Cell fractionation studies suggest that the mutant βIII-spectrin (with the 39-bp deletion) affects the plasma membrane localization of the synaptosomal proteins EAAT4 and GluRg2. Cell culture experiments (coexpression of GFP-EAAT 4 with either wt or mutant 39-bp deletion βIII-spectrin) showed that wt spectrin stabilized EAAT4 at the membrane. Several studies suggested the possible role of EAAT4 in ataxia [99, 161, 203]: loss of EAAT4 and GlurRg2 at the plasma membrane could lead to glutamate signaling abnormalities that over time could cause Purkinje cell death in SCA5. A homozygous mouse lacking full-length βIII-spectrin has a phenotype closely mirroring symptoms of SCA5 patients. However, heterozygous animals show no signs of ataxia or cerebellar degeneration (even up to 2 years of age) arguing against haploinsufficiency as a disease mechanism, but suggesting a dominant-negative of human mutations on wild-type βIII-spectrin function [142]. βIII-spectrin has been also reported to be associated with Golgi and vesicle membranes [170] and to bind to the dynactin subunit ARP1 suggesting a possible role in transport [72]. The mutation found in the German family in the calponin homology (CH) domain may disrupt the ability of spectrin to bind to the actin cytoskeleton and consequently may affect the stabilization of membrane protein or this mutation may cause alterations in transport by disrupting binding to ARP1 and the dynein motor complex. Interaction of βIII-spectrin with Arp1 is lost with the L253P substitution. Cell culture studies reveal that the L253P mutant βIII-spectrin, instead of being found at the cell membrane, appears trapped in the cytoplasm associated with the Golgi apparatus. Moreover, the L253P βIII-spectrin prevents correct localization of wt βIII-spectrin and prevents EAAT4 from reaching the plasma membrane. These data provide evidence for a dominant-negative effect of a SCA5 mutation and show

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that trafficking of both βIII-spectrin and EAAT4 from the Golgi is disrupted through failure of the L253P mutation to interact with Arp1 [27]. Loss of βG -spectrin in Drosophila melanogaster and Cenorhabditis elegans, which have one form of β-spectrin closed to mammal βII-spectrin, results in destabilization of the neuromuscular junction through loss of synaptic cell-adhesion molecules [147] and axonal breakage [65], respectively. Expression of either human βIII-spectrins or fly βG -spectrins containing SCA5 mutations in the eye of Drosophila causes a progressive neurodegenerative phenotype, and their expression in larval neurons results in posterior paralysis, reduced synaptic terminal growth, and axonal transport deficits. These phenotypes are genetically enhanced by both dynein and dynactin loss-of-function mutations. So, SCA5 mutant spectrin causes adult-onset neurodegeneration in the fly eye and disrupts fundamental intracellular transport processes that are likely to contribute to this progressive neurodegenerative disease. Defects in protein trafficking have been observed in other neurodegenerative diseases, such as a dominantly inherited motor neuron disease caused by mutations in p150Glued, a subunit of dynactin [149], in Huntington disease [111, 179] and in Alzheimer disease [132]. Identifying additional mutations in the SPTBN2 gene that cause ataxia in families having unknown mutations will provide further insight into the functions of βIIIspectrin and the molecular mechanism of neurodegenerative diseases. Moreover, it will be of interest to look for mutations in other β-spectin genes such as SPTBN1 and SPTBN5 which map to the SCA11 and SCA25 critical regions in ataxia. Consistent with the possibility that β-spectrin may have additional roles in disease, dominantly inherited mutations in the βII-spectrin homologue (unc70/bsg1) in C. elegans cause an uncoordinated phenotype [133].

17.3.2

Spectrin Defects in the West Syndrome

West syndrome is an uncommon to rare epileptic disease that affects babies under the age of 1 year. Also known as infantile spasms, it affects about 1 in 5,000 babies and accounts for about 1 in 20 cases of childhood epilepsy. West syndrome is characterized by spasms, a specific electroencephalogram pattern called hypsarrhythmia, developmental regression and often learning disability which occur as a result of one of a variety of possible causes. In about 70–80% of cases the cause can be identified, such as birth trauma and starvation of oxygen from the baby’s brain, or a condition known as tuberose sclerosis. Unfortunately, the long-term prognosis is very poor, although this depends to some extent on the underlying cause and whether this is a progressive disease. It is conjectured that it is a defective neurotransmitter function, or more precisely, a defect in the regulation of the GABA transmission process. Besides the only two contributing genes, ARX (Aristaless-related homeobox gene) and CDKL5 (Cyclin-dependent kinase-like 5 gene) described in a subset of familial and sporadic X-linked WS cases, three cases of early-onset WS with cerebral hypomyelination harboring SPTAN1 defects were recently reported. In two subjects, it was found de novo heterozygous mutations in βII-spectrin: an in-frame 3-bp

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deletion (c. 6619–6621 del) leading to E2207 del in the continuous helix region between the last two α19 and α20 spectrin repeats, and in-frame 6-bp duplication (c.6923–6928dup, R2308-M2309 dup) within the α20 spectrin repeat. These mutations are located within the spectrin repeats involved at the initial nucleation site between α-and β-spectrins and they were predicted to affect formation of αβspectrin heterodimers. They cause early-onset WS with spastic quadriplegia, poor visual attention, and severe developmental delay. Both subjects showed severe cerebral hypomyelination, decreased white matter, widespread brain atrophy including brainstem, hypoplasia and/or atrophy of the cerebellum, and a thinned and shortened corpus callosum. These in-frame mutations could cause aggregation of αII (mut)/βII and αII (mut)/βIII spectrin heterodimers, suggesting dominant negative effects of the mutations. βII- and βIII-spectrins have been shown to participate in stabilization of membrane proteins and axonal transport [72, 101]: they could disturb clustering of ankyrinG and voltage-gated sodium channels (VGSC) at axon initial segments, together with an elevated action potential threshold. In metazoans, they coordinate activities of VGSC, underlie cellular excitability and control neuronal communication, cardiac excitation-contraction coupling, and skeletal muscle function. Sodium channel dysfunction is associated with arrhythmia, epilepsy, and myotonia. In myelinated nerve fibers, action potential initiation and propagation requires that voltage-gated ion channels be clustered at high density in the axon initial segments and nodes of Ranvier. Previous results indicate that αII-spectrin is enriched at the paranodes which flank the node of Ranvier, notably in nodes and paranodes at early stages of development as observed in zebrafish [129, 181]. The nodal expression diminishes as nodes mature in zebrafish, αII-spectrin mutants harboring a nonsense mutation destabilize nascent clusters of sodium-channel clusters (VGSC) and affect normal development of mature nodes of Ranvier. The mutants also showed impaired myelination in motor nerves and in the dorsal spinal cord, suggesting that αII-spectrin plays important roles in the maintenance of the integrity of myelinated axons. These findings revealed essential roles of a αII-spectrin in human brain development and suggest that abnormal axon initial segment (AIS) is possibly involved in pathogenesis of infantile epilepsy.

17.3.3

BIV-Spectrin Defects in Mice

The involvement of spectrin in human disease outside the context of red blood cells has recently been revealed. The contributing effect of αII- and βIII-spectrin in degenerative disease has been clearly demonstrated. However, the other spectrin genes such as the βIV-spectrin might also be involved in the pathological process as suggested by different mouse models. Indeed, spontaneous autosomal recessive mutations in the mouse spectrin βIV gene (spnb4 coding for a protein of 2,554 residues/2,559 in human sharing 95% identity), an orthologue of human βIV spectrin (SPTBN4) affect the nervous system

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and cause a progressive ataxia with hind limb paralysis, deafness, and tremor in quivering mice (also known as qv, [135]. This phenotype was associated with six spontaneous alleles: qv1j , qv2j , qv3j, qv4j, qvInd, and qvInd2j. Each of the five quivering mutations described results in truncation of the βIV-spectrin: 81, 65, 86, 53, and 19% of the wild-type length, respectively. The two-point mutations introducing a stop codon, C6397T (R2078-stop, repeat 17) in qv allele and the C4234T (Q1358stop) in qv4J (β10 repeat) leads to the loss of the COOH end domain involved in the tetramer formation and ankyrin binding. These two point mutations are predicted to result in peptide truncation and loss of functional domains. The qv2J allele carries a 7-base deletion (5185–5191) that causes a frame shift at amino-acid R1675 and extension of an aberrant 35 amino-acid peptide in β13 repeat, the qv3J allele contains a single-base insertion (InsT6786) that produces a frame shift at amino acid G2209 and a new 49 amino-acid extension (PH domain). The qvInd allele has a single-base pair deletion delC1601, causing a frame shift and an extension of a 12 amino-acid-paptide downstream of residue A480 (α2 repeat). Loss of peptide length or reduced expression correlated with the severity of the mutant phenotype. Progressive deletion of β-spectrin repeats may compromise the formation of the tetramer (as observed in hereditary elliptocytosis), reducing its ability to anchor peptides including ion channels at specialized subcellular domains. Characterization of βIV-spectrin knockout mice [85] demonstrated that βIV-spectrin acts as a multifunctional regulatory platform for sodium channels and has important roles in the structure and stability of excitable membranes in the heart and brain, targeting critical structural and regulatory proteins. Ankyrin-G and voltage-gated sodium channels, which normally cocluster with βIV-spectrin at nodes of Ranvier and axon initial segments [18] were mislocalized in the mutants. Inversely, βIV-spectrin was mislocated in ankyrinG knockout mice, suggesting mutually dependent targeting of these two proteins. Voltage gate-dependent sodium channels, neurofascin (the L1 family cell-adhesion molecule), and βIV-spectrin (all of which interact with ankyrin in vitro) were mislocated in ankyrin-G knockout mice [79, 201]. In a similar way, the ankyrinB (-/-) mice exhibit a severe phenotype that partially overlapped the phenotype of human patients with L1 mutations [159]. Besides its role in the formation of the functional domain (targeting and stabilization of proteins), βIV-spectrin might be involved in a regulatory mechanism for Na+ channels (Nav1.5), via direct phosphorylation by βIV-spectrin-targeted calcium-/calmodulin-dependent kinase II [73]. These findings provide evidence for an unexpected yet commanding molecular platform involving spectrin that determines vertebrate membrane excitability. Thus, knockouts of spectrin and ankyrin have direct consequences for the development of the mammalian nervous system. They affect neuronal physiology by interfering with the normal targeting and stabilization of interacting sodium channels, adhesion molecules, and perhaps other physiologically important membrane activities. Based on observations in mice, humans with mutations in βIV-spectrin might show symptoms ranging from nonsyndromic auditory neuropathy to syndromic peripheral motor neuropathies such as Charcot-Marie-Tooth syndrome, one form of which maps to human chromosome 19q13.3 in the region homologous to quivering [89].

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Conclusions and Perspectives

Characterization of the underlying defects in human hemolytic anemia was fruitful in the understanding of not only the role of the spectrin and the spectrin-based skeleton in red blood cells, but also of their structure and their organization. Recent genetic studies in human disease are now helping to reveal the functions of spectrin outside the erythrocyte. Spectrins and the spectrin-based skeleton system are starting to reveal its secrets in nonerythroid cells and the ensuing clinical implications are just beginning to be appreciated. Spectrin defects manifest in neuropathies, and other components of the spectrin-based skeleton and associated proteins are also involved in human disease such as ankyrin mutations which affect the heart (cardiac arrhythmias linked to sudden death, [17, 119, 120, 160]). In a similar way, the data obtained from several animal models (nematode, fruit fly, zebrafish, and mice) have firmly established that one important role of spectrins consists in organizing specialized functional domains of the plasma membrane, and in some cases spectrin mutations have lethal phenotypes with drastic effects on cell morphology and tissue organization [39, 171]. Moreover, spectrins are involved in interactions with a broad panel of proteins such as channels (TRCP channels, [128], receptors (NMDA receptor, [185], and adhesion molecules (Lu/BCAM, N-CAM, [60, 97]). All these interactions suggest an involvement of spectrins in numerous cell processes: proliferation [114, 172], cell-cell contact establishment [15, 84], cell adhesion and migration [19, 114], signaling [202], actin dynamics [153], and trafficking, endoexocytosis and DNA repair [95, 113, 168, 199].

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M.-C. Lecomte beta-III spectrin leads to Purkinje cell dysfunction recapitulating the behavior and neuropathology of spinocerebellar ataxia type 5 in humans. J Neurosci 30:4857–4867 Perrotta S, Miraglia del Giudice E, Alloisio N, Sciarratta G, Pinto L, Delaunay J, Cutillo S, Iolascon A (1994) Mild elliptocytosis associated with the alpha 34 Arg → Trp mutation in spectrin Genova (alpha I/74). Blood 83:3346–3349 Perrotta S, Iolascon A, De Angelis F, Pagano L, Colonna G, Cutillo S, Miraglia del Giudice E (1995) Spectrin Anastasia (alpha I/78): a new spectrin variant (alpha 45 Arg → Thr) with moderate elliptocytogenic potential. Br J Haematol 89:933–936 Perrotta S, Gallagher PG, Mohandas N (2008) Hereditary spherocytosis. Lancet 372:1411– 1426 Perrotta S, Della Ragione F, Rossi F, Avvisati RA, Di Pinto D, De Mieri G, Scianguetta S, Mancusi S, De Falco L, Marano V, Iolascon A (2009) Beta-spectrinBari: a truncated beta-chain responsible for dominant hereditary spherocytosis. Haematologica 94:1753–1757 Pielage J, Fetter RD, Davis GW (2005) Presynaptic spectrin is essential for synapse stabilization. Curr Biol 15:918–928 Pradhan D, Morrow J (2002) The spectrin-ankyrin skeleton controls CD45 surface display and interleukin-2 production. Immunity 17:303–315 Puls I, Jonnakuty C, LaMonte BH, Holzbaur EL, Tokito M, Mann E, Floeter MK, Bidus K, Drayna D, Oh SJ, Brown RH Jr, Ludlow CL, Fischbeck KH (2003) Mutant dynactin in motor neuron disease. Nat Genet 33:455–456 Qualtieri A, Pasqua A, Bisconte MG, Le Pera M, Brancati C (1997) Spectrin Cosenza: a novel beta chain variant associated with Sp alphaI/74 hereditary elliptocytosis. Br J Haematol 97:273–278 Robledo RF, Lambert AJ, Birkenmeier CS, Cirlan MV, Cirlan AF, Campagna DR, Lux SE, Peters LL (2010) Analysis of novel sph (spherocytosis) alleles in mice reveals allele-specific loss of band 3 and adducin in alpha-spectrin-deficient red cells. Blood 115:1804–1814 Rocha S, Costa E, Rocha-Pereira P, Ferreira F, Cleto E, Barbot J, Quintanilha A, Belo L, Santos-Silva A (2010) Erythrocyte membrane protein destabilization versus clinical outcome in 160 Portuguese Hereditary Spherocytosis patients. Br J Haematol 149:785–794 Rotter B, Bournier O, Nicolas G, Dhermy D, Lecomte MC (2005) AlphaII-spectrin interacts with Tes and EVL, two actin-binding proteins located at cell contacts. Biochem J 388:631–638 Roux AF, Morle F, Guetarni D, Colonna P, Sahr K, Forget BG, Delaunay J, Godet J (1989) Molecular basis of Sp alpha I/65 hereditary elliptocytosis in North Africa: insertion of a TTG triplet between codons 147 and 149 in the alpha-spectrin gene from five unrelated families. Blood 73:2196–2201 Sahr KE, Tobe T, Scarpa A, Laughinghouse K, Marchesi SL, Agre P, Linnenbach AJ, Marchesi VT, Forget BG (1989) Sequence and exon-intron organization of the DNA encoding the alpha I domain of human spectrin. Application to the study of mutations causing hereditary elliptocytosis. J Clin Invest 84:1243–1252 Sahr KE, Laurila P, Kotula L, Scarpa AL, Coupal E, Leto TL, Linnenbach AJ, Winkelmann JC, Speicher DW, Marchesi VT et al (1990) The complete cDNA and polypeptide sequences of human erythroid alpha-spectrin. J Biol Chem 265:4434–4443 Sakaguchi G, Orita S, Naito A, Maeda M, Igarashi H, Sasaki T, Takai Y (1998) A novel brain-specific isoform of beta spectrin: isolation and its interaction with Munc13. Biochem Biophys Res Commun 248:846–851 Salomao M, An X, Guo X, Gratzer WB, Mohandas N, Baines AJ (2006) Mammalian alpha Ispectrin is a neofunctionalized polypeptide adapted to small highly deformable erythrocytes. Proc Natl Acad Sci U S A 103:643–648 Scotland P, Zhou D, Benveniste H, Bennett V (1998) Nervous system defects of AnkyrinB (-/-) mice suggest functional overlap between the cell adhesion molecule L1 and 440-kD AnkyrinB in premyelinated axons. J Cell Biol 143:1305–1315 Sedlacek K, Stark K, Cunha SR, Pfeufer A, Weber S, Berger I, Perz S, Kaab S, Wichmann HE, Mohler PJ, Hengstenberg C, Jeron A (2008) Common genetic variants in ANK2 modulate QT interval: results from the KORA study. Circ Cardiovasc Genet 1:93–99

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161. Serra HG, Byam CE, Lande JD, Tousey SK, Zoghbi HY, Orr HT (2004) Gene profiling links SCA1 pathophysiology to glutamate signaling in Purkinje cells of transgenic mice. Hum Mol Genet 13:2535–2543 162. Shotton DM, Burke BE, Branton D (1979) The molecular structure of human erythrocyte spectrin. Biophysical and electron microscopic studies. J Mol Biol 131:303–329 163. Sikorski AF, Terlecki G, Zagon IS, Goodman SR (1991) Synapsin I-mediated interaction of brain spectrin with synaptic vesicles. J Cell Biol 114:313–318 164. Sikorski AF, Hanus-Lorenz B, Jezierski A, Dluzewski AR (2000a) Interaction of membrane skeletal proteins with membrane lipid domain. Acta Biochim Pol 47:565–578 165. Sikorski AF, Sangerman J, Goodman SR, Critz SD (2000b) Spectrin (betaSpIIsigma1) is an essential component of synaptic transmission. Brain Res 852:161–166 166. Speicher DW, Ursitti JA (1994) Spectrin motif. Conformation of a mammoth protein. Curr Biol 4:154–157 167. Speicher DW, Weglarz L, DeSilva TM (1992) Properties of human red cell spectrin heterodimer (side-to-side) assembly and identification of an essential nucleation site. J Biol Chem 267:14775–14782 168. Sridharan DM, McMahon LW, Lambert MW (2006) alphaII-Spectrin interacts with five groups of functionally important proteins in the nucleus. Cell Biol Int 30:866–878 169. Stabach PR, Morrow JS (2000) Identification and characterization of beta V spectrin, a mammalian ortholog of Drosophila beta H spectrin. J Biol Chem 275:21385–21395 170. Stankewich MC, Tse WT, Peters LL, Ch’ng Y, John KM, Stabach PR, Devarajan P, Morrow JS, Lux SE (1998) A widely expressed betaIII spectrin associated with Golgi and cytoplasmic vesicles. Proc Natl Acad Sci U S A 95:14158–14163 171. Tang Y, Katuri V, Dillner A, Mishra B, Deng CX, Mishra L (2003) Disruption of transforming growth factor-beta signaling in ELF beta-spectrin-deficient mice. Science 299:574– 577 172. Tang Y, Katuri V, Srinivasan R, Fogt F, Redman R, Anand G, Said A, Fishbein T, Zasloff M, Reddy EP, Mishra B, Mishra L (2005) Transforming growth factor-beta suppresses nonmetastatic colon cancer through Smad4 and adaptor protein ELF at an early stage of tumorigenesis. Cancer Res 65:4228–4237 173. Thompson JM, Ellis RE, Green EM, Winlove CP, Petrov PG (2008) Spectrin maintains the lateral order in phosphatidylserine monolayers. Chem Phys Lipids 151:66–68 174. Trave G, Pastore A, Hyvonen M, Saraste M (1995) The C-terminal domain of alpha-spectrin is structurally related to calmodulin. Eur J Biochem 227:35–42 175. Tse WT, Lecomte MC, Costa FF, Garbarz M, Feo C, Boivin P, Dhermy D, Forget B G (1990) Point mutation in the beta-spectrin gene associated with alpha I/74 hereditary elliptocytosis. Implications for the mechanism of spectrin dimer self-association. J Clin Invest 86:909–916 176. Tse WT, Gallagher PG, Pothier B, Costa FF, Scarpa A, Delaunay J, Forget BG (1991) An insertional frameshift mutation of the beta-spectrin gene associated with elliptocytosis in spectrin nice (beta 220/216). Blood 78:517–523 177. Tse WT, Gallagher PG, Jenkins PB, Wang Y, Benoit L, Speicher D, Winkelmann JC, Agre P, Forget BG, Marchesi SL (1997) Amino-acid substitution in alpha-spectrin commonly coinherited with nondominant hereditary spherocytosis. Am J Hematol 54:233–241 178. Tse WT, Tang J, Jin O, Korsgren C, John KM, Kung AL, Gwynn B, Peters LL, Lux SE (2001) A new spectrin, beta IV, has a major truncated isoform that associates with promyelocytic leukemia protein nuclear bodies and the nuclear matrix. J Biol Chem 276:23974–23985 179. Twelvetrees AE, Yuen EY, Arancibia-Carcamo IL, MacAskill AF, Rostaing P, Lumb MJ, Humbert S, Triller A, Saudou F, Yan Z, Kittler JT (2010) Delivery of GABAARs to synapses is mediated by HAP1-KIF5 and disrupted by mutant huntingtin. Neuron 65:53–65 180. Ursitti JA, Kotula L, DeSilva TM, Curtis PJ, Speicher DW (1996) Mapping the human erythrocyte beta-spectrin dimer initiation site using recombinant peptides and correlation of its phasing with the alpha-actinin dimer site. J Biol Chem 271:6636–6644 181. Voas MG, Lyons DA, Naylor SG, Arana N, Rasband MN, Talbot WS (2007) alphaII-spectrin is essential for assembly of the nodes of Ranvier in myelinated axons. Curr Biol 17:562–568

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182. Wandersee NJ, Birkenmeier CS, Gifford EJ, Mohandas N, Barker JE (2000) Murine recessive hereditary spherocytosis, sph/sph, is caused by a mutation in the erythroid alpha-spectrin gene. Hematol J 1:235–242 183. Wang DS, Miller R, Shaw R, Shaw G (1996) The pleckstrin homology domain of human beta I sigma II spectrin is targeted to the plasma membrane in vivo. Biochem Biophys Res Commun 225:420–426 184. Waugh RE, Agre P (1988) Reductions of erythrocyte membrane viscoelastic coefficients reflect spectrin deficiencies in hereditary spherocytosis. J Clin Invest 81:133–141 185. Wechsler A, Teichberg VI (1998) Brain spectrin binding to the NMDA receptor is regulated by phosphorylation, calcium and calmodulin. Embo J 17:3931–3939 186. Wichterle H, Hanspal M, Palek J, Jarolim P (1996) Combination of two mutant alpha spectrin alleles underlies a severe spherocytic hemolytic anemia. J Clin Invest 98:2300–2307 187. Wilmotte R, Marechal J, Morle L, Baklouti F, Philippe N, Kastally R, Kotula L, Delaunay J, Alloisio N (1993) Low expression allele alpha LELY of red cell spectrin is associated with mutations in exon 40 (alpha V/41 polymorphism) and intron 45 and with partial skipping of exon 46. J Clin Invest 91:2091–2096 188. Wilmotte R, Miraglia del Giudice E, Marechal J, Perrotta S, de Mattia D, Delaunay J, Iolascon A (1994) A deletional frameshift mutation in spectrin beta-gene associated with hereditary elliptocytosis in spectrin Napoli. Br J Haematol 88:437–439 189. Wilmotte R, Harper SL, Ursitti JA, Marechal J, Delaunay J, Speicher DW (1997) The exon 46encoded sequence is essential for stability of human erythroid alpha-spectrin and heterodimer formation. Blood 90:4188–4196 190. Wilmotte R, Marechal J, Delaunay J (1999) Mutation at position −12 of intron 45 (c→t) plays a prevalent role in the partial skipping of exon 46 from the transcript of allele alphaLELY in erythroid cells. Br J Haematol 104:855–859 191. Winkelmann JC, Forget BG (1993) Erythroid and nonerythroid spectrins. Blood 81:3173– 3185 192. Winkelmann JC, Chang JG, Tse WT, Scarpa AL, Marchesi VT, Forget BG (1990) Full-length sequence of the cDNA for human erythroid beta-spectrin. J Biol Chem 265:11827–11832 193. Winograd E, Hume D, Branton D (1991) Phasing the conformational unit of spectrin. Proc Natl Acad Sci U S A 88:10788–10791 194. Yan Y, Winograd E, Viel A, Cronin T, Harrison SC, Branton D (1993) Crystal structure of the repetitive segments of spectrin. Science 262:2027–2030 195. Yoon SH,Yu H, Eber S, Prchal JT (1991) Molecular defect of truncated beta-spectrin associated with hereditary elliptocytosis. Beta-spectrin Gottingen. J Biol Chem 266:8490–8494 196. Yu J, Fischman DA, Steck TL (1973) Selective solubilization of proteins and phospholipids from red blood cell membranes by nonionic detergents. J Supramol Struct 1:233–248 197. Zarkowsky HS, Mohandas N, Speaker CB, Shohet SB (1975) A congenital haemolytic anaemia with thermal sensitivity of the erythrocyte membrane. Br J Haematol 29:537–543 198. Zhang Z, Weed SA, Gallagher PG, Morrow JS (2001) Dynamic molecular modeling of pathogenic mutations in the spectrin self-association domain. Blood 98:1645–1653 199. Zhang P, Sridharan D, Lambert MW (2010a) Knockdown of mu-calpain in Fanconi anemia, FA-A, cells by siRNA restores alphaII spectrin levels and corrects chromosomal instability and defective DNA interstrand cross-link repair. Biochemistry 49:5570–5581 200. Zhang Y, Resneck WG, Lee PC, Randall WR, Bloch RJ, Ursitti JA (2010b) Characterization and expression of a heart-selective alternatively spliced variant of alpha II-spectrin, cardi+, during development in the rat. J Mol Cell Cardiol 48:1050–1059 201. Zhou D, Ursitti JA, Bloch RJ (1998) Developmental expression of spectrins in rat skeletal muscle. Mol Biol Cell 9:47–61 202. Ziemnicka-Kotula D, Xu J, Gu H, Potempska A, Kim KS, Jenkins EC, Trenkner E, Kotula L (1998) Identification of a candidate human spectrin Src homology 3 domain-binding protein suggests a general mechanism of association of tyrosine kinases with the spectrin-based membrane skeleton. J Biol Chem 273:13681–13692 203. Zuo J, De Jager PL, Takahashi KA, Jiang W, Linden DJ, Heintz N (1997) Neurodegeneration in Lurcher mice caused by mutation in delta2 glutamate receptor gene. Nature 388:769–773

Chapter 18

Laminopathies Nadir M. Maraldi and Giovanna Lattanzi

Abstract The laminopathies are a group of rare diseases characterized by a vast range of phenotypic alterations, due to mutations in lamin A and C or other nuclear envelope proteins. A-type lamins, as well as B-type lamins, belong to the type V intermediate filaments and, by polymerization, form the nuclear lamina, a component of the nuclear envelope. Following a brief description of the complex interactions between lamins and proteins of the nuclear membrane, this Chapter describes disease phenotypes that characterize each laminopathy, the possible mechanisms involved into the pathogenesis, as well as potential therapies based on the use of existing drugs.

18.1

Introduction

For many years, the nuclear lamina has been considered a structural component of the nuclear envelope (NE, [1]) and has been studied by cell biologists working on the description of the different phases of mitosis [2]. After the discovery that mutations in the genes encoding A-type lamins and associated NE proteins cause an impressive range of human diseases [3], new and unexpected functions have been attributed to lamins and the interest in the nuclear lamina has been greatly enhanced. Here, a description of the nuclear lamina intermediate filament network and of laminlinked diseases known as “laminopathies” will be provided. Although a detailed description of the NE is outside the scope of this Chapter, a brief overview of the general organization of this nuclear domain, with references to recent reviews on the subject will be presented in order to help the comprehension of disease mechanisms. As schematically indicated in Fig. 18.1, in the NE the outer and inner nuclear membranes ONM and INM) are not equivalent; in fact, the ONM is continuous N. M. Maraldi () Laboratory of Musculoskeletal Cell Biology, Rizzoli Orthopedic Institute, Via di Barbiano 1/10, 40136 Bologna, Italy e-mail: [email protected] Department of Anatomical Sciences, University of Bologna, Bologna, Italy G. Lattanzi National Research Council of Italy CNR-IOR, Institute of Molecular Genetics, Unit of Bologna, Bologna, Italy e-mail: [email protected]

M. Kavallaris (ed.), Cytoskeleton and Human Disease, DOI 10.1007/978-1-61779-788-0_18, © Springer Science+Business Media, LLC 2012

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Fig. 18.1 The nuclear envelope and the nuclear lamina. The nuclear envelope is composed by the inner and outer nuclear membranes, which delimit the perinuclear space (pns). Inner nuclear membrane proteins are emerin, MAN1, LBR, SUN1, and nesprin 1. Nesprin 2 is an outer nuclear membrane protein. TorsinA is located in the pns. The nuclear lamina is a meshwork of intermediate filaments located in the nucleoplasmic face of the inner nuclear membrane. Lamin B1, Lamin B2, Lamin A and C, and prelamin A are nuclear lamina proteins. LAP2α is associated with the nuclear lamina. BAF (barrier to autointegration factor) links nuclear envelope/lamina proteins to chromatin. Transcription factors interacting with nuclear envelope/lamina proteins are represented as black labels

with the endoplasmic reticulum, to which it is functionally related, while the INM is characterized by a set of proteins that mediate the attachment of the cytoskeleton to the nuclear lamina and to the underlying chromatin [4]. Integral proteins of the INM include the lamin B receptor (LBR), MAN1, emerin, lamina-associated polypeptide 1 (LAP1), LAP2β, small nesprin 1 isoforms, SUN1, and SUN2. SUN proteins bind nesprin 1 in the INM and the large nesprin 2 isoform in the ONM, which in turn interact with cytoskeletal actin filaments ([5], Fig. 18.1). The nuclear lamina (Fig. 18.1) is a protein three-dimensional meshwork that lines the nucleoplasmic face of the INM [4]. The nuclear lamina is essential in maintaining the structural integrity of the interphase nucleus, and allows the disassembly and reassembly of the nucleus during the different steps of mitosis. A- and B-type lamins are type V intermediate filament (IF) proteins, able to assemble into higher-order filamentous structures [4]. In humans, the A-type lamins, with lamin A and C being the product of alternative splicing, are encoded by the LMNA gene on chromosome 1. The B-type lamins, lamin B1 and lamin B2, are encoded by the LMNB1 gene on chromosome 5 and the LMNB2 on chromosome 19, respectively. The LMNA gene consists of 12 exons (Fig. 18.2); lamin C is encoded by exons 1–9 and a portion of exon 10, while lamin A results from alternative splicing, which adds exon 11 and 12 and removes the lamin C-specific portion of exon 10. Like other IF proteins, lamin A and C present conserved α-helical central domains, while head and tail domains are variable globular domains (Fig. 18.2). Lamin polymerization requires the association of monomers to form parallel coiled-coil homodimers that interact in a head-to-tail fashion and then laterally to form antiparallel protofilaments; eight of these protofilaments combine to form the characteristic 10-nm IF structure [6]. Higher-order polymer formation might involve a separate polymerization of A- and B-type lamins as distinct homopolymers [6].

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Fig. 18.2 The LMNA gene and prelamin A maturation. a The LMNA gene is composed by 12 exons. Representative mutations causing laminopathies are shown in the rectangles. Lamin A and lamin C protein structure are reported. b Prelamin A processing. Lamin A is translated as a 664-kDa precursor (prelamin A), which undergoes farnesylation, cleavage, methylation in its C-terminal CaaX box, and it is finally endoproteolysed to yield mature lamin A. The four prelamin A forms produced during processing are represented

B-type lamins are expressed in all cell types, while A-type lamins are expressed in a tissue-specific manner during differentiation [7]. Lamins are also specifically modified posttranslationally; B-type lamins are isoprenylated in order to be able to attach to INM proteins [8]. While lamin C does not undergo posttranslational modification, lamin A is isoprenylated, methylated, and subsequently cleaved by a specific metalloproteinase (Fig. 18.2, [9, 10]).

18.2

Short Story of Diseases Related to the Nuclear Lamina

The relevance of the study of the so-called laminopathies apparently exceeds the clinical prevalence of these disorders. Laminopathies present a broad spectrum of disease phenotypes which include striated muscle disorders, lipodystrophies, metabolic disorders, and premature aging syndromes. Among the inherited disorders caused by mutant nuclear envelope proteins, Emery-Dreifuss muscular dystrophy (EDMD) has been regarded with a particular interest, since its genetic characterization [11]. This disease is caused by mutations

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affecting the EMD gene on chromosome X (X-linked form of EDMD, EDMD1) that encodes the inner nuclear membrane-associated protein emerin. An indistinguishable disease phenotype (EDMD2) is caused by mutations in the LMNA gene on chromosome 1 [12, 13], which encodes lamin A and C. EDMD1- and EDMD2-affected patients present identical phenotypic features, consisting of slowly progressive wasting of specific muscles, early contractures of the ankles, elbows, and neck, and dilated cardiomyopathy with severe cardiac conduction defects [13]. This represents the first paradox of laminopathies; in fact, two genes, when mutated, cause the very same morphological and functional alterations which characterize a single pathological entity, phenotypically recognized as EDMD. The second paradox of laminopathies depends on the fact that several disease phenotypes, corresponding to distinct pathological entities, have been then demonstrated to depend on mutations of a single gene [14]. In fact, mutation in LMNA have been demonstrated to cause several tissue-specific diseases: the lamin-linked autosomal dominant form of EDMD (EDMD2, [12]) and its recessive form, known as EDMD3 [15], the limb-girdle muscular dystrophy type 1B (LGMD 1B, [16]), the dilated cardiomyopathy and conduction-system disease (CMD1A, [17]), the Dunningan-type familial partial lipodystrophy (FPLD2, [18]), and the Charcot-Marie-Tooth disorder type 2 (CMT 2B1, [19]). Each disease selectively strikes one or more specific tissues, including skeletal and cardiac muscle, tendons, adipose tissue, and peripheral neurons. A further group of laminopathies has been then identified, characterized by a systemic involvement of almost all the tissues, which may undergo premature senescence. The progeric laminopathies include the Hutchinson-Gilford progeria syndrome (HGPS, [20, 21]), Atypical Werner syndrome (AWS, [22]) and atypicalprogeroid syndrome (APS, [23]), and Mandibuloacral dysplasia (MADA, [24]). Restrictive dermopathy (RD, [25]) and lethal fetal akysinesia [26] are severe developmental syndromes with features resembling other laminopathic syndromes [14]. While the reported laminopathy phenotypes can be considered as separate entities, some overlapping phenotypes have been also reported. For example, the coexistence of EDMD2, CMD1A, and LGMD1B has been recognized within a family [27]. Moreover, some patients have been reported to carry features of more than one laminopathy phenotype, such as FPLD2 with cardiac involvement [28] or mandibuloacral dysplasia with myopathy [29]. These findings suggest that laminopathies are a complex group of multisystem disorders, whose phenotypes cannot be always considered as discrete entities, but rather show wide overlap, some individuals and families having features of several different laminopathies and some mutations causing diverse manifestations in different individuals [30]. In very recent years, new diseases associated with A-type lamins or NE proteins have been identified [14]. It is conceivable that new pathological entities will increase the growing group of these inherited diseases in the near future. Laminopathies, including pathologies linked to LMNA mutations or prelamin A processing defects, will be described in Sects. 2.1–2.7. Diseases associated with nuclear membrane protein mutations and their phenotypes will be described in Sect. 2.8. The latter diseases are referred to, in this Chapter, as “nuclear envelopathies.”

18 Laminopathies

18.3

379

Laminopathies

The inherited diseases associated with LMNA mutations that result into alterations of a restricted type of tissues are referred to as tissue-specific laminopathies. Laminopathies in which several tissues are involved have been named systemic laminopathies and include premature aging syndromes and developmental syndromes. In Table 18.1, besides the main disease features, the characteristic alterations at both tissue and cell level, when available, are reported for each clinical entity.

18.3.1

Laminopathies With Skeletal Muscle Involvement

18.3.1.1 EDMD2 and EDMD3 About 60–70% of cases due to LMNA mutations show involvement of striated muscles. The autosomal dominant form of Emery-Dreifuss muscular dystrophy (EDMD2) is characterized by slowly progressive weakness and wasting of the humeroperoneal and limb girdle muscles, associated with contractures of the Achilles tendons, elbow, and postcervical muscles [12]. Cardiac involvement with conduction system defects, arrhythmia and, later, dilated cardiomyopathy are very common [13, 31]. The onset of the pathology is in the first or second decade, but early onset has been also reported, while cardiomyopathy may occur later [13]. Muscle biopsy findings include fiber-type disproportion, variation in fiber size, enlarged nuclei and occasional degeneration or necrosis. Lamin A and C and emerin immunostaining has been reported to be quite normal in muscle biopsies [32, 33]; on the other hand, characteristic nonuniform distribution of both Lamin A and C and emerin have been reported in skin fibroblasts from EDMD2 patients [34]. Ultrastructural alterations have been reported in muscle biopsies, including nuclear lamina thickening, nuclear pore clustering, as well as focal absence of heterochromatin and loss of contact between heterochromatin and the nuclear lamina [32]. Prelamin A is not accumulated neither in biopsies nor in cultured cells from EDMD2 patients [34, 35]. No significant differences in the clinical phenotype have been reported in the few cases of autosomal recessive EDMD (EDMD3, [15]). Clinical and genetic data on EDMD are continuously updated by Bonne and colleagues at http://www.ncbi. nlm.nih.gov/books/NBK1436/. 18.3.1.2 LGMD 1B Limb girdle muscular dystrophy type 1B (LGMD1B) due to missense mutations or splicing defects in LMNA is a dominantly inherited disorder characterized by a slowly progressive weakness and wasting of shoulder and pelvic muscles, without

TMPO

LMNA

CMD1A

CMD1T

HHS

Dilated cardiomyopathy with conduction defect

Dilated cardiomyopathy Heart hand syndrome

LMNA

LMNA

MDCL

Muscular dystrophy congenital, LMNA-related

LMNA LMNA

LGMD1B LMNA

EDMD2 EDMD3

Acronym Mutated gene

Limb girdle muscular dystrophy, type 1B

Laminopathies Emery-Dreifuss muscular dystrophy, type 2/3

Disease

Lamin A and C

LAP2alpha

Lamin A and C

Lamin A and C

Lamin A and C

Lamin A and C Lamin A and C

Mutated NE protein

AD

AD

AD

AD

AD

AD AR

Common mutations

Table 18.1 Diseases of the nuclear lamina and of the nuclear envelope

Early contractures (stiff/fixed joints) of elbows, Achilles tendon, neck, and spine. Progressive muscle weakness in upper arms and lower limbs. Dilated cardiomyopathy with conduction abnormalities Progressive weakness of shoulder, upper arm, hip and leg muscles with later development of dilated cardiomyopathy with conduction abnormalities Neonatal onset. Selective axial weakness and wasting of the cervicoaxial muscles, upper limbs with proximal wasting and lower limbs with distal wasting. Contractures Ventricular dilation and impaired systolic function. Sudden death due to cardiac pump failure may occur after conduction abnormalities. No skeletal muscles affected Left ventricular severe dilated cardiomyopathy Conduction system disease, atrial and ventricular tachyarrhythmias, sudden death, dilated cardiomyopathy, brachydactyly

Phenotype

Nuclear lamina disorganization, lamin A aggregates

NA

Nuclear lamina disorganization (honeycomb appearance)

Nuclear lamina disorganization (honeycomb appearance)

Cellular phenotype

#610140

A 68-kDa Lamin A and C isoform is detected in cellular lysates

* 188380 NA

#115200

#613205

#159001

#181350

OMIM

380 N. M. Maraldi and G. Lattanzi

APS

Atypical progeria syndrome

HGPS

Adult-onset autosomal dominant leukodystrophy Hutchinson–Gilford progeria syndrome

AWS

LMNA

ADLD

Charcot-MarieTooth disorder, type 2B1

Atypical Werner syndrome

CMT2B1 LMNA

LMNB1

FPLD2

Dunningan-type familial partial lipodystophy

LMNA

LMNA

LMNA

Acronym Mutated gene

Disease

Table 18.1 (continued)

Lamin A and C

Lamin A and C

Lamin A and C

Lamin B1

Lamin A and C

Lamin A and C

Mutated NE protein

AD

AD

AD

AD

AR

AD

Common mutations Loss of subcutaneous fat from limbs and trunk with simultaneous accumulation in face and neck. Insulin resistance and diabetes mellitus Progressive deterioration of motor and sensory nerves leading to atrophy of limb muscles and numbness/sensory problems. Nerve conduction velocities not affected Slowly progressive neurological disorder, symmetrical widespread myelin loss in the central nervous system Childhood onset of premature aging including growth retardation, baldness, facial hypoplasia, delayed tooth formation, aged skin, osteoporosis, atherosclerosis, arthritis. Teenage mortality due to cardiovascular disease Adult onset of premature aging. Hard, tight skin, cataracts, subcutaneous calcification, premature atherosclerosis, diabetes mellitus, premature aging of face Early onset of disease. Hard, tight skin, cataracts, subcutaneous calcification, premature atherosclerosis, diabetes mellitus, premature aging of face

Phenotype

*150330

*150330

#176670

#169500

#605588

#151660

OMIM

Nuclear dysmorphism

Prelamin A accumulation, dysmorphic nuclei

Progerin accumulation, nuclear dysmorphism, heterochromatin loss

Prelamin A accumulation, chromatin disorganization, lamin A aggregates NA

Cellular phenotype

18 Laminopathies 381

MADB

RD

Mandibuloacral dysplasia type B

Restrictive dermopathy

Nuclear envelopathies Emery-Dreifuss EDMD1 muscular dystrophy, type 1

MADA

Mandibuloacral dysplasia, type A

ZMPSTE24

Lamin A and C

Mutated NE protein

EMD

Emerin

FACE1 ZMPSTE24 LMNA LaminA and C

FACE1

LMNA

Acronym Mutated gene

Disease

Table 18.1 (continued)

X-linked

AR AD

AD

AR AD

Common mutations

Early contractures (stiff/fixed joints) of elbows, Achilles tendon, neck, and spine. Progressive muscle weakness in upper arms and lower limbs. Dilated cardiomyopathy with conduction abnormalities

Postnatal growth retardation, craniofacial abnormalities (especially crowding or loss of teeth), skeletal malformations, mottled skin pigmentation, stiff joints and autoimmune hair loss. Partial lipodystrophy, insulin resistance, and diabetes Postnatal growth retardation, craniofacial abnormalities (especially crowding or loss of teeth), skeletal malformations, mottled skin pigmentation, stiff joints, and autoimmune hair loss. Generalized lipodystrophy, insulin resistance, and diabetes Abnormally rigid and translucent skin, joint contractures, and pulmonary hypoplasia. Impaired fetal body movements lead to deformity. Early neonatal death due to respiratory insufficiency.

Phenotype

Prelamin A accumulation, nuclear lamina defects

Prelamin A accumulation, chromatin and nuclear lamina defects

Cellular phenotype

#310300

Nuclear lamina disorganization (honeycomb appearance)

#275210 Prelamin A *606480 accumulation, absence *150330 of mature lamin A

#608612

#248370

OMIM

382 N. M. Maraldi and G. Lattanzi

EDMD4 EDMD5

PHA

GSD/ HEM

BOS

DYT1

Emery-Dreifuss muscular dystrophy, type 4/5

Pelger–Huet anomaly

Greenberg/HEM skeletal dysplasia

Buschke–Ollendorff syndrome

Torsion dystonia

DYT1

LEMD3

LBR

LBR

SYNE1 SYNE2

Acronym Mutated gene

Disease

Table 18.1 (continued)

TorsinA

MAN1/LEMD3

Lamin B receptor

Lamin B receptor

Nesprin 1 Nesprin 2

Mutated NE protein

AD

AD

AR

AD

AD

Common mutations Early contractures (stiff/fixed joints) of elbows, Achilles tendon, neck, and spine. Progressive muscle weakness in upper arms and lower limbs. Dilated cardiomyopathy with conduction abnormalities Neutrophil nuclei in heterozygotes have fewer segments and course chromatin, with no effect on normal death. Homozygotes are also prone to epilepsy and skeletal abnormalities, e.g., polydactyly and metacarpal shortening Widespread tissue edema in fetus. Disorganized bone structure, short limbs, and conversion of cartilage to bone. Early in utero lethality Skeletal defects include multiple spots of increased bone density (osteopoikilosis) and band of sclerosis in a flowing pattern (melorheostosis). Sometimes accompanied by joint contractures, skin lesions, muscle atrophy, hemangiomas, and lymphedema Prolonged, involuntary muscle contractions induce abnormal posture and twisting or repetitive movements in arms and legs. Caused by CNS dysfunction rather than neurodegeneration

Phenotype

#128100

#166700

#215140

#169400

#612998 #612999

OMIM

Altered endoplasmic reticulum under stress

NA

Chromatin condensation, heterochromatin clumps

Neutrophil nuclei in heterozygotes have fewer segments and course chromatin

Nuclear lamina disorganization

Cellular phenotype

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contractures. Cardiac involvement, with conduction system disease and dilated cardiomyopathy, is common [16]. The onset of the pathology is in the first decade, but may occasionally occur later.

18.3.1.3 MDCL LMNA mutations have been identified in muscular dystrophies with early onset and severe phenotype. The disease entity is best classified as a congenital muscular dystrophy (LMNA-related congenital muscular dystrophy, or MDCL, [36–38]). Dropped head is typical of patients affected by MDCL [38], while cardiac arrythmias are mostly asymptomatic, but may cause sudden death [37]. Contractures and spine rigidity are always reported [36].

18.3.2

Laminopathies With Heart Involvement

18.3.2.1 CMD1A Autosomal dominant mutations in LMNA are frequently associated with dilated cardiomyopathy with conduction system disease (CMD1A). The disease is characterized by a progressive ventricular dilation and systolic dysfunction, accompanied by conduction defects and may include skeletal muscle involvement [17]. Because of the risk of sudden death, the use of an implantable cardioverter-defibrillator can prevent possible lethal ventricular arrhythmias [39, 40]. The prevalence of LMNA mutations in unselected, nonfamilial dilated cardiomyopathy cases is low, but the yield increases by including familial cases with evidence of skeletal muscle weakness [40]. A comprehensive description of clinical and genetic data on CMD1A can be found at the following Web site: http://www.ncbi.nlm.nih.gov/books/NBK1674/#dcmlmna.REF.hershberger.2008.1

18.3.2.2 HHS Heart-hand syndrome (HHS) is due to an LMNA mutation that creates a new cryptic splicing site with the retention of 11 intronic nucleotides in the mRNA. From the clinical point of view, HHS has similarities with CMD1A; however, progressive cardiac system disease, tachyarrhythmias, and dilated cardiomyopathy have an adult onset and are associated with brachydactily [41]. In HHS fibroblasts, a 550 amino acid truncated Lamin A and C is produced due to the abnormal splicing [41]. By immunofluorescence, nuclei appear dysmorphic and show nuclear envelope herniations; emerin is normally localized at the nuclear rim, while aberrant foci of accumulated Lamin A and C are localized within the nucleoplasm [41].

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385

Laminopathies With Adipose Tissue Involvement

18.3.3.1 FPLD2 Dunnigan-type familial partial lipodystrophy (FPLD2) is an autosomal dominant disorder due to LMNA mutations often located in exon 8, with substitutions at Arginine-482 in 75% of cases [18, 42]. The disease phenotype is characterized by loss of fat from extremities and excess fat accumulation on the face and neck, beginning at puberty. The phenotype is less pronounced in males. Patients are affected by insulin-resistant type 2 diabetes, hyperinsulinemia, hyperlipidemia, and atherosclerosis [18, 43]. Other metabolic disorders including polycystic ovary are frequently associated with the disease. Cardiomyopathy and muscle weakness have been reported in some families. A mutation at the intron 8 consensus splice donor site, leading to a prematurely terminated lamin A isoform has been reported in a severe FPLD2 case [44]. Partial lipodystrophy is also present in MADA [24], while generalized lipodystrophy characterizes HGPS [45] and atypical Werner Syndrome [22]. Fibroblasts from FPLD2 patients present characteristic nuclear alterations [46] due to the accumulation of abnormal amounts of prelamin A [35]. The dysmorphic FPLD2 nuclei present intranuclear prelamin A aggregates [35], an enlarged and irregular nuclear profile as well as a reduced amount of peripheral heterochromatin [46].

18.3.3.2 Metabolic Syndromes A peculiar form of generalized lipoatrophy, insulin-resistant diabetes, leukomelanodermic papules, liver steatosis, and hypertrophic cardiomyopathy has been reported to be associated with a heterozygous mutation in the rod domain of Lamin A and C; nuclear abnormalities were also found in primary fibroblast cultures [47]. Another form of generalized lipodystrophy with features of progeroid syndromes has been reported in a patient with a T10I LMNA mutation [48]. Polycystic ovary syndrome and insulin-resistant diabetes without lipodystrophy has been reported in a case showing a unique LMNA mutation (G602S). Also, in this case, the skin fibroblasts exhibited nuclear alterations similar to those described in FPLD2 and other laminopathies [49].

18.3.4

Laminopathies With Nerve Involvement

18.3.4.1 CMT2B1 Autosomal recessive Charcot-Marie-Tooth type 2 disease (CMT2B1) due to homozygous mutation in the LMNA gene is characterized by an onset of the clinical features in the second decade of life. These features include weakness, wasting, and areflexia

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of distal lower limb muscles and pes cavus. CMT disease includes both the type 1 demyelinating forms of hereditary motor and sensor neuropathies with highly reduced nerve conduction velocity, and the type 2 axonal forms with slightly reduced nerve conduction velocity. In CMT2B1, the motor nerve conduction velocities are quite normal or slightly reduced [19]. 18.3.4.2 AD-CMT2 An autosomal dominant form of CMT2 (AD-CMT2) has been reported, in one family, as associated with muscular dystrophy, cardiac disease, and leukonychia, or associated with cardiac disease and partial lipodystrophy, in other two families [50, 51]. Molecular and cell biology studies concerning the CMT2B1 pathogenesis are not yet available. 18.3.4.3 ADLD Adult-onset autosomal dominant leukodystrophy (ADLD) is a slowly progressive neurological disorder characterized by symmetrical widespread myelin loss in the central nervous system [52]. The disease is linked to mutations in LMNB1 encoding the type V intermediate filament lamin B1 and it is caused by a duplication in the gene [52].

18.3.5

Premature Aging Syndromes

18.3.5.1 HGPS Hutchinson–Gilford progeria syndrome (HGPS) is an autosomal dominant condition which develops in the first or second year of life, followed by severe and rapid features of premature senescence which involve almost all tissues [45, 53]. The patients present delayed growth, short stature, alopecia, thinning of skin, loss of subcutaneous fat, midface hypoplasia, skeletal involvement with osteolysis and fractures, premature atherosclerosis, cardiac failure leading to death or stroke at about 13.5 years [53]. A de novo missense mutation in exon 11 leads to creation of an abnormal splice donor site that results in expression of a truncated, permanently farnesylated prelamin A, termed progerin [20, 21, 54]. This is the most common mutation in HGPS. Mutations in other exons of the LMNA gene give rise to less severe forms of the disease. 18.3.5.2 AWS and APS Atypical Werner syndrome (AWS) is a condition due to missense LMNA mutation characterized by premature aging with a later onset and milder course with respect

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to HGPS [22]. Because some typical features of WS, such as cataracts, are absent in these patients, while they present dilated cardiomyopathy and phalangeal osteosclerosis, it has been argued that these cases should be considered as atypical progeria syndrome (APS, [23]). 18.3.5.3 MADA Mandibuloacral dysplasia type A (MADA) is an autosomal recessive disorder prevalently associated with the homozygote R527H LMNA mutation [24]. MADA has been also linked to heterozygote compound mutations in the Lamin A and C rod domain [29] or other homozygote mutations. The disease is characterized by postnatal growth retardation, hypoplasia of the mandible and clavicles, acro-osteolysis, delayed closure of the cranial sutures, joint contractures, partial or generalized lipodystrophy, mottled skin pigmentation, premature loss of teeth, acanthosis nigricans, and moderate signs of premature aging [24]. Patients also present insulin resistance with diabetes and hypertriglyceridemia [24]. The missense homozygote R527H LMNA mutation has been mainly identified in Italian and Mexican families [24, 55, 56], suggesting a founder effect. 18.3.5.4 MADB Mandibuloacral dysplasia type B (MADB) is caused by heterozygote mutations in the FACE1 gene yielding low enzyme levels and accumulation of prelaminA in cells [57]. MADB differs from MADA in the lipodystrophy phenotype, which appears more severe, and in the more accelerated aging process [55, 57]. The typical bone defects of MADA consisting in acro-osteolysis of phalanges and clavicles are also observed in MADB, associated with more severe disorders including multiple fractures and neonatal tooth eruption [57]. The increased severity of the disease with respect to MADA appears to be associated with the greater retention of farnesylated prelamin A in MADB cells [57]. In dermal fibroblasts, obtained from HGPS, AWS, APS, and MADA patients, typical nuclear alterations have been observed, mainly consisting in local or total loss of peripheral heterochromatin, associated with blebs and invaginations of the nuclear lamina [35, 54, 58]. In HGPS and MADA cells, the worsening of chromatin alterations and altered methylation of the heterochromatin marker HeK9me3 have been reported to increase with the age of the patient, as well as with the increasing amount of prelamin A [54, 58–60].

18.3.6

Developmental Syndromes

Mutations in the LMNA gene have been associated with developmental syndromes resulting in lethal phenotypes within the first hours or day after birth.

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18.3.6.1 RD Restrictive dermopathy (RD) is characterized by intrauterine growth retardation, thinning of the dermis, reduction of elastic fibers and skin erosions, prominent superficial vasculature, micrognathia, pulmonary hypoplasia, mineralization defects of the skull, clavicle osteolysis, and multiple joint contractures [25]. Neonatal lethal course generally occurs within the first week of life. Heterozygous splicing LMNA mutations resulting in truncated prelamin A were found in the reported cases [61]. Most RD cases are associated with homozygote mutations in the prelamin A endoprotease ZMPSTE24, a membrane protein of the endoplasmic reticulum and the nuclear envelope, which catalyzes cleavage and maturation of the lamin A precursor [25]. The genetic defect, typically the 1085_1086 insT mutation in the FACE1 gene, causes a null phenotype, with undetectable levels of enzyme activity [62]. Consistently, the cellular phenotype of RD is determined by accumulation of farnesylated prelamin A, which is toxic to cells and causes severe nuclear envelope and chromatin abnormalities [63]. Diagnosis of RD can be performed by genetic testing, as well as by immunocytochemical detection of prelamin A, a protein which is almost undetectable in normal cells [25, 63].

18.3.6.2 Lethal Fetal Akynesia Lethal fetal akynesia due to LMNA homozygous nonsense mutation has been reported in a newborn deceased child presenting dysmaturity, facial dysmorphism with retrognathia, severe contractures of the fingers and the toes, long-bone fractures, and severe generalized muscular dystrophy [26]. Lamin A and C-null skin fibroblasts showed severe nuclear abnormalities with blebbing and protrusions of chromatin in the cytoplasm, and reduced levels of emerin staining and local loss of lamin B and other nuclear envelope proteins [26, 53].

18.3.7

Overlapping Phenotypes

Isolated or familial cases of laminopathy have been reported, showing involvement of several tissues and suggesting overlapping phenotypes. These cases include: phenotypes combining clinical features of lipodystrophy presenting also variable combinations of skeletal and/or cardiac muscle alterations [28, 64, 65]; CMT2 axonal neuropathy phenotypes associated with myopathic features and cardiac disease [50]; phenotypes combining axonal neuropathy, partial lipodystrophy, and scapuloperoneal myopathy [51]; phenotypes combining early-onset myopathy and progeria [66]; phenotypes combining MAD and myopathy [29]. These findings confirm that laminopathies are a complex group of multisystem disorders and suggest that they could be regarded as a continuum of clinical phenotypes.

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18.3.7.1 Nuclear Envelopathies As indicated in Fig. 18.1, A-type lamins interact directly or indirectly with integral proteins of the inner nuclear membrane (INM proteins). Mutations affecting the expression of some INM proteins have been associated with diseases here referred to as “nuclear envelopathies” and share some aspects of laminopathies or completely overlap with the laminopathy phenotypes (Table 18.1).

18.3.7.2 EDMD1 The first nuclear-envelope-linked disease was discovered by the Daniela Toniolo group in 1994 [11]. It is the X-linked form of Emery-Dreifuss muscular dystrophy (EDMD1), which has been associated with mutations in the EMD gene encoding the inner nuclear membrane protein, emerin. The clinical phenotype of the disease overlaps with the autosomal forms (EDMD2, EDMD3, EDMD4, and EDMD5, described in other paragraphs), consisting of contractures, muscular weakness, and wasting and cardiomyopathy [11]. The genetic defect usually yields a null cellular phenotype (complete absence of emerin from the nuclear envelope), which allows rapid diagnosis of the pathology by immunofluorescence or Western blot analysis [67]. Missense mutations in EMD have been also reported [68]. Typical nuclear abnormalities consisting of altered nuclear envelope/lamina structure and focal loss of heterochromatin are observed in fibroblasts, myoblasts, and muscle tissue [69, 70].

18.3.7.3 EDMD4/EDMD5 EDMD4 and EDMD5 are caused by mutations in SYNE1 or SYNE2 genes encoding nesprin 1 and nesprin 2, respectively [71]. These proteins are located in the nuclear membrane and connect the nuclear envelope to the actin or plectin cytoskeleton [72]. The clinical phenotype of EDMD4 and EDMD5 [71] completely overlaps with EDMD1 and EDMD2, suggesting that emerin, Lamin A and C, and nesprins are major players in a muscle-specific regulatory pathway. They appear to play a major role in nuclear migration and anchorage [73], a function recently linked to the Lamin A precursor.

18.3.7.4 Pelger-Huet Anomaly Two diseases have been reported to be due to mutation of the gene encoding the Lamin B receptor (LBR). Autosomal dominant mutations in the LBR gene cause the PelgerHuet anomaly (PHA, [74]). PHA is a benign condition characterized by alterations in the morphology of neutrophil nuclei in heterozygotes, while homozygotes present polydactily, metacarpal shortening, and are prone to epilepsy [75].

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18.3.7.5 Greenberg/HEM Skeletal Dysplasia The Greenberg/HEM skeletal dysplasia is an autosomal recessive rare disease due to null mutations in the LBR gene. Greenberg dysplasia causes early in utero lethality due to severe and diffuse edema; also the skeletal organization is severely affected [76]. Nuclear alterations in cells consist of altered membrane profile and heterochromatin compaction with large heterochromatin clumps [76]. This heterochromatin alteration is different from all the other laminopathies and nuclear envelopathies, which feature heterochromatin loss.

18.3.7.6 Buschke–Ollendorff Syndrome (BOS) Mutations in the LEMD3 gene encoding MAN1, an inner nuclear membrane protein that interacts with lamins and emerin, cause the Buschke–Ollendorff syndrome (BOS). Multiple tissues—including bone, muscle, and joints—are affected in BOS [77], the prominent phenotype being associated with increased bone density and osteosclerosis. Other diseases associated with LEMD3 mutations are osteopoikilosis and melorheostosis all characterized by increased bone density [77].

18.3.7.7 Néstor-Guillermo Progeria Syndrome (NPGS) A homozygous mutation in BANF1 (c.34G > A (p.Ala12Thr), encoding barrier-toautointegration factor 1 (BAF), a nuclear envelope protein which interacts with emerin and lamins, has been associated with a new laminopathy called NéstorGuillermo progeria syndrome (NGPS, [78]). NGPS is characterized by mild progeroid tracts and severe bone abnormalities. The disease features bone resorption of phalanges, clavicles and mandible, resembling the MADA phenotype, but patients suffer more severe skeletal abnormalities, including complete absence of clavicles and scoliosis, that affect their quality of life. Since BAF is a key binding partner of prelamin A involved in chromatin dynamics [79], the pathogenetic pathway of NGPS likely involves altered interplay of BAF and prelamin A leading to chromatin misfunctioning.

18.3.7.8 Torsion Dystonia Torsion dystonia (DYT1) is a rare disease characterized by involuntary muscle contractions that induce twisting in arms and legs, due to mutation in the gene encoding torsinA, a nuclear membrane ATPase that mediates nuclear movements and maintains nuclear envelope integrity [80, 81]. Importantly, torsinA could be implicated in nucleo-cytoplasmic interplay, a function involving other nuclear envelope proteins such as nesprins, SUN1, and SUN2 [82, 83].

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18.3.7.9 Genotype-Phenotype Correlation To date, about 300 mutations have been reported to be distributed throughout the LMNA gene in more than 1,000 patients [84]. The majority of mutations are missense or frameshift mutations ([84], Table 18.1). Laminopathies with striated muscle involvement, account for about 60% of all laminopathies. EDMD2 and CMD1A mutations are distributed throughout the gene, while LGMD1B mutations are clustered in both the Ig-like fold and coil 2. In any case, these laminopathies present an inconsistent genotype-phenotype link. CMT2B1 appears to arise from a prevalent mutation (R298C). About 75% of patients with FPLD2, harbor mutations of codon 482, located in the Ig-like fold, while the remaining 25% mutations are distributed along the protein [14]. A different approach to the genotype-phenotype correlation is based on the analysis of mutation type with respect to particular Lamin A domains. All the phenotypic clustering studies indicate that the underlying disease mechanism may be associated with functional properties of the protein domains rather than with the type of mutation [84, 85]. For example, both the sterol-regulatory element-binding protein 1 (SREBP1) and barrier to autointegration factor (BAF) have binding sites in the Cterminal globular tail of Lamin A, where the most common FPLD2 mutations lie. So, disturbed interaction with the chromatin-binding protein BAF could lead to global higher-order chromatin alterations, while the aberrant binding with SREBP1 might interfere with adipocyte-specific differentiation processes [35]. By contrast, most mutations causing laminopathies with neuromuscular phenotypes are prevailingly located within the Lamin A hydrophobic core, potentially destabilizing the structural interactions with type-B lamins. Furthermore, it must be considered that disease phenotypes almost undistinguishable with respect to those caused by LMNA mutations have been reported to be due to mutations of other genes. EDMD1, due to mutation in the EMD gene encoding emerin [11], localized at the inner nuclear membrane, may be phenotypically indistinguishable from that of the autosomal dominant form [12]. The molecular interaction between A-type lamins and emerin [86] may account for a common pathogenic mechanism in these distinct diseases. EDMD has been also reported to be due to mutation of SYNE1, SYNE2 [71], and FHL1 genes [87]. The latter genes encode for nesprin 1, nesprin 2, and the LIM protein FHL1, all involved in nuclear anchorage and/or positioning. This argues in favor of a major pathogenetic role of altered nuclear positioning in EDMD. On the other hand, CMD1A, which presents cardiac conduction defects similar to those in EDMD patients, besides to be caused by mutations of the LMNA gene [17], has been also reported to be caused by mutations in the gene coding LAP2α [88], a nuclear lamina-associated protein which regulates cell cycle progression and stem cell activity [89]. Finally, some systemic laminopathies—including HGPS, MAD, and RD—are caused by mutations in either LMNA or FACE1 [57, 61, 90], strongly supporting the major role of altered prelamin A cleavage by the endoprotease ZMPSTE24 in the pathogenesis of those diseases.

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The emerging evidence shows that the laminopathies are a complex group of multisystem disorders, whose molecular disease mechanisms may be caused by a combination of effects of disease-causing mutations on structural or regulatory functions of Lamin A.

18.4

Pathogenesis of Laminopathies

Besides their role in maintaining, in association with B-type lamins, the mechanical stability of the NE throughout the phases of the cell cycle, A-type lamins and associated NE proteins represent scaffolds for molecular interaction with elements that regulate DNA synthesis and repair, higher-order chromatin organization, nuclear positioning, gene transcription, and cell differentiation [4]. Many of these functions involve Lamin A interplay with signal transduction pathways, transcription factors, and chromatin-associated proteins [91, 92].

18.4.1

Nuclear Envelope Defects as Determinants of Chromatin Misfunctioning

The expression of A-type lamins is modulated in a tissue-specific manner during the intermediate stages of embryonic development and during postnatal development [93] and can be considered as a marker of embryonic differentiation [94]. A-type lamins are required for postnatal growth and the maintenance of quiescence and differentiation. In fact, while Lamins A and C are expressed in the primary neuronal progenitor stem cells as well as in mature neurons, they are underexpressed in early and late progenitor cells [95]. Because hallmarks of laminopathy cells include characteristic deformation of the nuclear profile, nuclear envelope blebbing or thickening of the nuclear lamina [46, 54, 63, 69, 96], and nuclear fragility upon stress [97], it has been suggested that mutations in A-type lamins can affect the structural organization of lamin dimmers and filaments, leading to nuclear lamina disorganization and nuclear envelope deformation under stress [98, 99]. The mechanical stress pathogenic mechanism has been thus longer considered to be effective, particularly in tissues which undergo mechanically induced strain such as skeletal and cardiac muscles, vascular smooth muscles, or fibroblasts of some connective tissues. Many of the mutations causing the muscular laminopathies alter the lamin incorporation in the nuclear lamina [100], as well as the localization of emerin and nesprins [100], which could result in increased levels of apoptosis following mechanically induced strain [101]. However, the degree of nuclear blebbing does not correlate with the severity of the phenotype, and emerin deficiency in mice does not result into skeletal or cardiac abnormalities, while satellite-cell-mediated muscle regeneration appears to be affected [102]. These findings suggest that it may be too simplistic to consider physical weakening of the nuclei as the main cause of muscle defects in the laminopathies.

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Alterations of nuclear morphology are present in many types of laminopathies, frequently associated with alterations of the chromatin arrangement; these alterations occur also in cells not subjected to mechanical strain. Severe nuclear abnormalities have been reported in HGPS, AWS, RD, and MADA cells, including lobulation, blebbing, and loss of heterochromatin [46, 54, 58, 60, 63]. Aberrant nuclear morphology results in cellular senescence, downregulation of transcription, and apoptosis [54, 58, 103, 104]. Furthermore, there is increasing evidence of a role for lamins in the regulation of epigenetic marks in chromatin. Loss of heterochromatin in HGPS and RD cells is associated with downregulation of trimethylation at Lys-9 of histone H3 (H3K9), and reduced association with heterochromatin protein 1α (HP1α), [54, 60, 63]. Cells from older MADA patients also exhibit loss of peripheral heterochromatin, together with mislocalization of HP1β, trimethylated H3K9, and LBR [58]. Experimental evidence exists that association of certain gene loci with the nuclear periphery has a repressive effect on the transcription of these genes [105]. It is thus conceivable that alterations in the heterochromatin arrangement could influence gene transcription of specific gene loci or chromosome domains [105–107]. Alterations in the normal peripheral localization of chromosome 13 and 18 to a more interior position have been reported in fibroblasts from patients with Lamin A mutations [104]. Changes in histone methylation associated with the expression of mutant lamins could also account for the telomere shortening reported in cells from HGPS and AWS patients, or in cells overexpressing HGPS Lamin A mutants [106, 108]. Interestingly, a potential mediator of histone methylation is the retinoblastoma gene product (pRb, [109]). The pRb binds A-type lamins [110] and regulates the expression of EZH2, which is required for trimethylation of H3K27 [111]. Moreover, the pRb binds the histone methyl transferase SUV39H1, which is responsible for H3K9 methylation. These interactions may be impaired due to the expression of mutant lamins, leading to alterations in histone methylation patterns.

18.4.2

Nuclear Envelope Defects as Determinants of Altered Nucleo-Cytoplasmic Interplay

Positioning of the nucleus is an active mechanism essential to the formation of functionally polarized cells, cell division, cell migration, and formation of multinucleated syncytia. To properly position the nucleus, a class of transmembrane proteins, referred to as KASH (Klarscicht, ANC-1; Syne Homology) proteins, is expressed at the inner and outer nuclear membranes [112]. KASH proteins connect the ONM to the actin and intermediate filament cytoskeleton. In mammals, KASH proteins originally called Syne-1 and Syne-2 are also referred to as Nesprin-1 and Nesprin2 [112]. Nesprins require SUN (Sad1 and UNC-84) proteins for localization to the nuclear envelope [112, 113]. SUN proteins are transmembrane proteins localized at the INM which are expressed in a tissue-specific manner to maintain the even spacing between the INM and ONM [113]. SUN1 and SUN2 interact with Lamin A and

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prelaminA [114] and mediate nuclear positioning in muscle [115, 116]. SUN proteins recruit KASH proteins to the ONM, forming a bridge across the nuclear envelope and connecting the nucleoskeleton to the cytoskeleton. Nesprin 1 and nesprin 2, interacting with the actin network, might transmit mechanical stimuli from the plasma membrane to the chromatin, affecting gene transcription [73]. These mechanisms may be altered in EDMD4 and EDMD5 patients carrying heterozygous mutations in SYNE-1 and SYNE-2 genes. Because nesprins bind emerin, it has been suggested that disrupting the interaction with emerin could impair the coupling of the nucleus and the cytoskeleton, leading to the progression of EDMD [117]. This has been further confirmed by the demonstration that disruption of nesprin-1 causes an EDMD-like phenotype in mice and cardiomyopathy [118]. Importantly, EDMD2 muscle appears to be affected by altered prelamin A-SUN1 interplay leading to mislocalization of nuclei [116, 119]. Furthermore, abnormal nesprin-2 localization has been reported to correlate with the severity of nuclear defects in fibroblasts from a patient affected by AWS [120]. A tissue-specific pathogenetic mechanism involving nucleo-cytoplasmic interplay could be linked to the altered or disrupted functional interaction between emerin and proteins of the intercalated discs, including β-catenin [3, 68]. In fact, emerin has been localized both in the nuclear envelope and in the intercalated discs of myocardium, a localization suggesting its involvement in mechanosignaling processes in the heart. Finally, emerin-actin interplay in the nucleus, within a protein platform also containing Lamin A and C, might be involved in pathogenetic mechanisms, as shown in a mouse model [86].

18.4.3 Altered Cell Cycle Control A direct role for lamins in the organization of DNA replication is supported by the colocalization of lamin A with replication factors at specific stages of S-phase in mammalian cells [121]. When N-terminal deletion mutants of human Lamin A are added to assembled nuclei, the disruption of lamin organization results into DNA synthesis inhibition [122]. Transition from G1 - to S-phase is mainly regulated by the tumor suppressor Rb that in its hypophosphorylated active state binds to the E2F transcription factors and represses E2F target genes involved in the G1 - to S-phase transition, thereby arresting the cell cycle and promoting transition to quiescence, senescence, or differentiation. A-type lamins and LAP2α bind to E2F-dependent promoter sequences and repress E2F-dependent transcription [89, 123, 124]. Cells lacking either Lamin A or LAP2α show abnormal regulation of the cell cycle [89], although differences have been observed between mice and humans and different mechanisms of lamin-directed control of Rb function have been envisaged. In any case, deregulation of Rb activity could contribute to the pathogenesis of lamin-linked diseases; in fact, disruption of Lamin A and C and LAP2α localization may result in destabilization of Rb complexes,

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causing a compensatory upregulation of Rb and MyoD target genes and defects in muscle regeneration observed in EDMD muscle cells [125]. HGPS fibroblasts lose the ability to proliferate. This is associated with altered Lamin A-Rb signaling, as determined in genome-wide expression studies of HGPS cells [126].

18.4.4 Altered DNA Repair ROS accumulate at a higher rate in HGPS fibroblasts, as well as in normally aged fibroblasts [127]. This may contribute to increased levels of DNA damage and may trigger senescence in HGPS cells [105]. Moreover, oxidative stress appears involved in the pathogenesis of FPLD, whose fibroblasts undergo p16INK-dependent senescent arrest [103]. Impaired DNA repair and genomic instability also characterize MEFs of Zmpste24-null mice, as well as bone marrow cells and HGPS or RD fibroblasts undergoing premature senescence [128]. Fibroblasts from HGPS and MADA patients show increased amounts of basal phosphorylated histone variant H2AX (γH2AX), a marker of unprepared DNA damage sites [129]. Gamma-H2AX colocalizes with XPA foci, an essential factor of nucleotide excision repair (NER) and not with double-strand breaks (DBSs), suggesting that the damage in HGPS cells may be different from that accrued by other genotoxic agents [129]. Furthermore, fibroblasts from HGPS patients show a marked delay in the recruitment of p53-binding protein 1 (53BP1) to sites of DNA repair [128]. In HGPS fibroblasts and in ZMPSTE24−/− MEFs the recruitment of the repair factor p53-binding protein (53BP1) to sites of DNA damage is impaired, as well as the presence of fragmented DNA after irradiation [130]. Expression of HGPS mutant Lamin A in HeLa cells increases the levels of phosphorylated histone H2AX (γ-H2AX), a hallmark of double-strand breaks, and this is correlated with defects in DNA repair foci [131]. Other DNA damage signaling pathways, including ATM and ATR kinases as well as Rad50 and Rad51, are affected in HGPS and RD fibroblasts [132]. These findings suggest that defects in the DNA repair machinery are due to expression of abnormal levels of farnesylated prelamin A in progeric laminopathies. DNA damage-initiated genomic instability as well as p53-mediated cell senescence and apoptosis in response to DNA damage may contribute to aging [133].

18.4.5

Telomere Dysfunction

Cell proliferation defects in HGPS cells may also be linked to telomere dysfunction [108, 134]. This is consistent with the observation that telomeres cluster in intranuclear foci containing Lamin A and C during senescence and apoptosis in human mesenchymal stem cells [135]. Consistent with the hypothesis that cell proliferation

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defects in progeric cells may be due to telomere dysfunctions is the evidence that these defects can be rescued either by expression of the catalityc subunit of telomerase, or by the inactivation of p53 [136], although HGPS fibroblasts are refractory to telomerase-mediated immortalization [137]. Moreover, telomere shortening and dysfunction can also trigger DNA damage responses and induce cellular senescence in animal model of progeric syndromes [138]. Increased telomere dysfunction, on the other hand, appears to be linked to genomic instability associated with the aging process [138]. These findings suggest that both telomere shortening and DNA damage, along with DNA repair deficiencies, could result in genomic instability and accelerated cellular senescence; these mechanisms, which are both dependent on p53 status, can act cooperatively to trigger the onset of aging phenotypes.

18.4.6 Altered Cellular Signaling Signaling pathways that modulate transcription factor activity may be regulated by interactions with components of the NE and lamina. Several recent studies indicate that lamins and INM proteins provide a platform for sequestering, through direct chromatin-independent interactions, transcriptional regulators operating in different signal transduction pathways. This association could restrict access to their target genes and limit their transactivation or transrepression abilities [92]. FPLD and MADA present partial lipodystrophy and share the molecular defect, consisting of accumulating prelamin A [35, 58, 59, 103]. In cells accumulating prelamin A, an in vivo binding occurs between prelamin A and the adipocyte transcription factor SREBP1; prelaminA sequesters SREBP1 at the nuclear rim, reducing the pool of DNA-bound active transcription factors; retention of SREBP1 causes downregulation of PPAR-γ expression and reduces the rate of preadipocyte differentiation [35]. Treatment of preadipocytes accumulating prelamin A with the PPAR-γ ligand TDZ elicited rescue of adipogenic program, giving insight into possible therapeutic approaches to lipodystrophy [43, 139, 140]. Evidence has been obtained, in animal models, that A-type lamins, emerin [102], and LAP2-α [141] are involved in the regulation of cell cycle exit of myoblasts through the modulation of the pRb expression, localization, and phosphorylation [141, 142]. It has been thus suggested that altered expression of Lamin A and C and/or lamin-interacting NE partners should result into impaired pRb-mediated muscle cell differentiation [125, 143]. Some evidence suggests that pRb-MyoD signaling cascade could be involved also in the pathogenesis of lamin-linked cardiomyopathy. In fact, mutation in LAP2α that interferes with Lamin A binding has been reported to cause cardiomyopathy in humans [88], and knockout mice for LAP2α are affected by systolic dysfunction and fibrosis [143]. Absence of A-type lamins affects the localization at the inner nuclear membrane of MAN1, a negative regulator of Smad-mediated signal transduction [144]. Defects in regulatory Smads have been reported in laminopathic cells; by using embryonal fibroblasts from LMNA-null mice, it has been shown that phosphorylation kinetics of Smad2 and Smad3 induced by TGF-β are altered [145]. These effects could also

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be due to the reported interaction of A-type lamins with PPA2 phosphatase, which could result into an A-type lamin-mediated dephosphorylation of rSmads [145]. Cardiac and skeletal muscle from homozygous mice carrying the Lamin A and C H222P substitution, which causes EDMD in humans, show an excess accumulation of phosphorylated Smad2 and Smad3 in nuclei [146]. ERK1/2 activation is reduced in C2C12 skeletal myoblasts treated with siRNA to knock down A-type lamins or emerin [147]. Defective differentiation of myoblasts expressing EDMD-causing mutations did not correlate with ERK activation, although myogenesis was enhanced by a treatment with the ERK1/2 inhibitor PD98059 and insulin-like growth factor II, which increased the pool of dephosphorylated pRb [148]. Thus, although ERK1/2 activation may be not affected in EDMD2 skeletal myoblasts, stimulation of pRb dephosphorylation by inhibiting cdk4 might be utilized to rescue myogenic differentiation. In animal models of EDMD2 that feature dilated cardiomyopathy, the abnormal activation of ERK1/2 and JNK activity has been reported [149]. The block of the development of cardiomyopathy—by the use of PD98059, a specific inhibitor of ERK—further sustains the key role of ERK activation in the pathogenesis of dilated cardiomyopathy due to Lamin A and C mutation [147]. Human subjects with MAN1 mutations exhibited increased bone density and overexpression of TGF-β [77]. This resulted in enhanced expression of genes activated by TGF-β and BMPs, conceivably due to failure of mutant MAN1 to bind Smads [150]. In MADA, as well as in HGPS and RD, bone resorption due to osteolysis has been observed [59]. A hint for the involvement of osteoclasts in bone defects associated with A-type lamin mutations comes from the evidence that Lamin A is able to sequester cFos [151], a transcription factor involved in osteoclast differentiation. Hyperactivation of the p53 pathway occurs in Zmpste24-null mice [152]. Some feature of the Zmpste24-null phenotype, including an increase of the onset of cell senescence, are ameliorated in the Zmpste24−/− Tp53−/− double mutant, indicating that the premature aging phenotype is, at least in part, dependent on p53 hyperactivation [152]. Defective Notch signaling has been implicated in HGPS, because stem cells from HGPS patients present an upregulation in the expression of Notch target genes, possibly due to the increased levels of progerin [153]. Moreover, expression of progerin in human MSCs impairs differentiation toward well-defined lineages by interfering with the Notch signaling pathway, which is essential in stem cell regulation [153]. Because Notch3 modulates the response to vascular injury, it has been suggested that progerin-induced defects in Notch signaling contribute to alterations in the large arteries of HGPS patients [154]. Increased Wnt/β-catenin signaling induces the conversion of aged muscle satellite cells into fibroblasts; this results into a poor regenerative response associated with fibrosis in the aged muscle [155]. Hyperactivation of β-catenin has been demonstrated in Emery-Dreifuss cells lacking emerin expression (EDMD1), in association with increased proliferation of fibroblasts [156]. Altered Wnt signaling has been recently reported in a mouse model of HGPS characterized by accumulation of farnesylated

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prelaminA [157]. In this case, the nuclear localization and activity of the transcription factor Lef1 appear to be altered in both the animal model and in human cells, thus affecting the Wnt signaling pathway [157]. Insulin growth factor 1 (IGF1) signaling is considered an important regulator of longevity in many organisms [158]. Progeroid Zmpste24−/− mice present profound transcriptional alterations in genes that regulate the somatotroph axis, as well as high circulating levels of growth hormone and reduction of IGF1; this suggests that an impaired response to growth hormone (GH) might elicit altered expression of IGF1, as confirmed by the observation that administration of IGF1 extends life span in progeroid mice [158]. These data suggest that decreased level of IGF play a key role in the onset of premature aging processes and propose these signaling effectors as promising targets of therapeutic interventions in progeric laminopathies.

18.4.7 Altered Stem Cell Functioning A-type lamins are expressed in a tissue-specific manner during middle stages of embryonic development and during postnatal development [94]; therefore, they can be considered essential for postnatal growth and the maintenance of tissue homeostasis. Besides being involved in tissue-specific differentiation mechanisms, A-type lamins and NE proteins are linked to the maintenance and regeneration of many mesenchymal tissues [123, 159]. Adult stem cell maintenance requires a fine balance between the multiple signaling pathways which modulate cell self-renewal, growth arrest, and lineage specification during tissue regeneration upon damage. A-type lamins have been proposed to be guardians of somatic cells during their lifetime and to be linked to longevity. Aging is associated with degenerative disorders affecting different tissues, including those derived from mesenchymal stem cells. Many tissues affected by mutations in the A-type lamin gene are also involved in degenerative conditions, characteristic of aging, including sarcopenia, osteopenia, cardiomyopathy, lipoatrophy, atherosclerosis, and dermoarthropathy [123]. These disorders might reflect an altered proliferative homeostasis between cell loss and cell replacement through adult stem cells in aged tissues. A loss of appropriate regulation of pathways intrinsic to adult stem cell maintenance may result into the premature demise of tissues with A-type lamin mutations. Along this line, the Lamin A partner LAP2α has been shown to regulate cellular proliferation and cell cycle exit in stem cells from different niches, including epidermal and muscular precursors [123, 143]. Some signaling pathways central to adult stem cell regeneration have been linked to these functions of A-type lamins and their binding partners. A loss of regulation of regeneration pathways linked to adult stem cell maintenance may result into premature demise of cells with Lamin A and C mutations. This, in turn, may lead to defects in adult stem cell growth and differentiation, and to activation of chronic stress signaling pathways that maintain organism survival but promote the aging phenotype [159]. This is consistent with recent data showing that downregulation of genes encoding extracellular matrix proteins, such as collagen I and fibronectin in

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laminopathic mice, associated with deficient Wnt signaling, affects stem cell proliferation [157]. Importantly, it has been demonstrated that Lamin A pathogenetic mutants are not expressed equally in all stem cell types, suggesting a mechanism for the segmental nature of laminopathies [157].

18.5 Therapy of Laminopathies In this section, the experimental therapeutic approaches to laminopathies and efficient pharmacological tools routinely used for treatment of laminopathic patients are reviewed.

18.5.1

Skeletal Muscle Laminopathies

Since contractures are a major impairment in EDMD, surgical extension of Achilles tendons and cervical ligaments is used to improve the quality of life. The combined treatment with PD98059 (an inhibitor of ERK1/2) and IGF-II (an activator of PI3-kinase) contributes to enhance myogenesis in Lamin A-R453W myoblasts [148]. It has been suggested that this combined treatment promotes a switch from cell proliferation to differentiation, causing a downregulation of phosphorylated pRb and cyclin D3, as well as an upregulation of myogenin [148]. The efficacy of these treatments in animal models has not been tested so far.

18.5.2

Cardiac Laminopathies

Like in other forms of cardiomyopathy, the defects of the cardiac conduction system present in lamin-associated cardiomyopathy and arrhythmias can benefit from implantable cardioverter-defibrillators (ICDs) to prevent the incidence of sudden death [40]. A survey of ICD utilization in 19 subjects with LMNA mutations followed over 3 years indicates that about 40% of these patients had an ICD discharge during the period of observation [160]. The results of this study validate the use of ICDs as prevention of sudden death and indicate that cardiac conduction system disease can precede the development of overt cardiac dilation and arrhythmias. In nongenetic forms of cardiomyopathy, ACE inhibitors and β-adrenergic blockers constitute standard therapy; this therapy has been proven to be effective in reducing alterations in left ventricular function and size, also in dystrophinopathies, and it is also effective in lamin-associated cardiomyopathies [39, 160]. Pharmacological inhibitors of ERK1/2, such as PD98059, whose hyperactivation has been related to the pathogenesis of DCM in EDMD [147], have been found to block the development of cardiomyopathy in LMNAH222P/H222P mice before the appearance of clinical symptoms [147].

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Lipodystrophies

Insulin resistance in lipodystrophic syndrome is due to an insufficient capacity of the adipose tissue to buffer dietary fatty acids, with consequent lipotoxicity, due to deposition of triglycerides and acyl-CoA in insulin-sensitive tissues, increased by leptin deficiency. Expression of PPARγ is crucial in the entraining of adipose tissue metabolism to nutritional state, by upregulating genes that mediate fatty acid uptake [161]. Improvement of the metabolic phenotype through PPARγ activation can be achieved by thiazolinediones (TZDs) therapy [43]. Furthermore, TZDs are likely to enhance the ability of adipose tissue to act as a pump for dietary fatty acids, removing them from other insulin-sensitive tissues, including skeletal muscle and liver [162]. PPARγ pathway is, therefore, a potential target of TZDs also in lipodystrophic laminopathies. Importantly, prelamin A accumulation, which occurs in FPLD2, reduces the expression levels of PPARγ and affects adipocyte activity [35, 43, 103]. TZDs are able to rescue adipogenic differentiation in vitro, which had been inhibited by prelamin A accumulation [35]. These drugs act as coactivators of PPARγ which controls the gene expression pathway leading to adipocyte differentiation. TZDs can also act at a systemic level [162]. Insulin sensitization by PPARγ activation upon TZDs administration could elicit depot-selective responses of adipose tissue, with increased accumulation in subcutaneous regions and reduction of visceral depots. Finally, the insulin-sensitizing effects of TZD might be influenced by the release of specific adipocytokines, such as adiponectin and the retinol-binding protein 4, by adipose tissue cells [161]. Several clinical studies have been reported, in which a positive response to glitazone treatment was observed in patients affected by FPLD2 [43, 140, 161]. Unfortunately, fat loss and ectopic accumulation are not actually rescued by TZD treatment of patients. This unexpected outcome is currently under investigation. FPLD2 patients are often treated with metformin, in combination or not with pioglitazone, to improve the insulin-dependent disorders [140]. Leptin treatment of FPLD2 patients has been attempted with promising results. Long-term efficacy in improving glycemic control and normalization of glucose tolerance has been reported in six FPLD2 patients, in which recombinant leptin was given through twice-daily injections over 12 months [161].

18.5.4

Syndromic Laminopathies

Since the discovery in 2003 that HGPS is associated with LMNA mutations, pharmaceutical tools capable of interfering with toxic accumulation of farnesylated prelamin A have been tested in cellular and animal models and have been shown to improve the senescent phenotype in cellular and animal models. Given that farnesylated prelamin A exerts a pathogenetic effect in HGPS [60, 62], RD, MADA, and MADB [57, 58, 63], FTIs, statins, and bisphosphonates have been used to impair prelamin A farnesylation. Treatment with these drugs reduces the percentage of misshapen nuclei in HGPS

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cells and recovers heterochromatin organization and nuclear transcriptional activity [54]. In animal models of progeria, inhibiting prelamin A farnesylation improves the bone phenotype and extends longevity [163]. FTI treatment of HGPS cells did not result in a reduction in DNA double-strand breaks and damage checkpoint signaling, suggesting that DNA damage accumulation and aberrant nuclear morphology are independent phenotypes due to accumulation of progerin [132]. Statins, that are widely used as cholesterol-lowering agents, by inhibiting the HGM-CoA reductase, the rate-limiting enzyme of the mevalonate pathway of cholesterol synthesis, have been proven to reduce prelamin A farnesylation and progeroid phenotype in cells and animal models [54, 163]. Amino-bisphosphonates (N-BPs) that are used to prevent osteoporosis and the risk of pathological fractures in bone malignancies act as inhibitors of farnesyl-phyrophosphate synthase, thus reducing the synthesis of both geranyl-geranyl and farnesyl groups. Therefore, statins and N-BPs have been tested in animal models of progeria and their efficacy has been proven [163]. Although the molecular mechanisms by which FTIs ameliorate disease phenotype in human progeric cells and in mouse model of progeria remain to be further elucidated, some open-label trials are ongoing in children with HGPS and MADA. A phase II clinical trial is ongoing since 2007 at the Children’s Hospital in Boston, which administered the FTI lonafarnib to 26 HGPS children. Another phase II open trial is ongoing since October 2008 at the Hopitaux de Marseille, France, which plans the administration of a statin and an N-BP (pravastatin and zolendronic acid) to 15 HGPS patients. Three MADA patients have been included in a phase II trial at the Policlinico Tor Vergata, Rome, Italy, receiving a treatment with pravastatin and zolendronic acid for 2 years. The results of these clinical trials appear encouraging, because increased growth, reduced levels of markers of cardiovascular risk, and no side effects have been reported in all the patients. Other approaches to progeria therapy have been recently attempted in mouse models with intriguing results. Low IGF-1 levels have been observed in progeric Zmpste24−/− mice, which accumulate farnesylated prelamin A, associated with overexpression of miR1 [138]. Recombinant IGF-1 treatment is able to restore the correct signaling pathway and growth hormone levels in Zmpste24−/− mice, resulting into a delayed onset of many progeroid features and extending the lifespan of the animals [158]. The treatment of cultured fibroblasts from adult progeric mice bearing the homozygous LMNAL530P/L530P mutation, as well as cell lines from HGPS patients, with a Gsk-3β inhibitor, which activates the Wnt effector β-catenin, has been shown to improve survival and proliferation [157]. Gene delivery systems and in vivo gene therapy have been also suggested as possible therapeutical approaches in HGPS. Lentiviral expression of short hairpin RNAi (shRNAi) targeted against prespliced and mutant LMNA mRNA led to a significant decrease of cell senescence, increase of proliferation, and recovery of nuclear morphology in HGPS fibroblasts. Because the major cause of death in HGPS is from coronary artery atherosclerosis, it has been suggested that shRNAi may be used to prevent arterial wall stenosis, once efficient delivery systems targeting the arterial wall have been developed.

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Modified oligonucleotides (morpholinos) targeted to the cryptic splice site of progerin have been demonstrated to reverse the disease phenotype of HGPS cells, suggesting that this approach deserves translation into the clinic, although morpholinos have been reported to cause some “off-target” effects in gene expression knockdown studies [164].

18.6

Conclusions

Research on laminopathies provides a paramount example of intersection between basic cell biology and clinical medicine. Moreover, the studies on disease pathogenesis and treatments of laminopathies greatly overcome the clinical importance of these rare monogenic diseases, because several processes appear to be also involved in common forms of cardiomyopathy, metabolic disorders, as well as in physiological aging. Acknowledgments The authors wish to thank Marta Columbaro and Cristina Capanni for revising the manuscript and for the helpful discussion. This work was supported by: A. I. Pro. Sa. B., Italy; EU-funded FP6 Euro-Laminopathies project; Italian MIUR PRIN 2008 to G. L. and N. M. M.; ISS Rare Diseases Italy–U. S. A. program (grant number 526/D30); Fondazione Carisbo, Italy, FIRB-MIUR Grant 2010 to N. M. M.

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57. Agarwal AK, Fryns JP, Auchus RJ et al (2003) Zinc metalloproteinase, ZMPSTE24, is mutated in mandibuloacral dysplasia. Hum Mol Genet 12:1995–2001 58. Filesi I, Gullotta F, Lattanzi G et al (2005) Alterations of nuclear envelope and chromatin organization in mandibuloacral dysplasia, a rare form of laminopathy. Physiol Genomics 23:150–158 59. Maraldi NM, Lattanzi G (2007) Involvement of prelamin A in laminopathies. Crit Rev Eukaryot Gene Expr 17:317–334 60. Shumaker DK, Dechat T, Kohlmaier A et al (2006) Mutant nuclear lamin A leads to progressive alterations of epigenetic control in premature aging. Proc Natl Acad Sci U S A 103:8703–8708 61. Navarro CL, De Sandre-Giovannoli A, Bernard R et al (2004) Lamin A and ZMPSTE24 (FACE-1) defects cause nuclear disorganization and identify restrictive dermopathy as a lethal neonatal laminopathy. Hum Mol Genet 13:2493–2503 62. Moulson CL, Go G, Gardner JM et al (2005) Homozygous and compound heterozygous mutations in ZMPSTE24 cause the laminopathy restrictive dermopathy. J Invest Dermatol 125:913–919 63. Columbaro M, Mattioli E, Schena E et al (2010) Prelamin A processing and functional effects in restrictive dermopathy. Cell Cycle 9:4766–4768 64. Muchir A, van Engelen BG, Lammens M et al (2003) Nuclear envelope alterations in fibroblasts from LGMD1B patients carrying nonsense Y259X heterozygous or homozygous mutation in lamin A/C gene. Exp Cell Res 291:352–362 65. Van Der Kooi AJ, Bonne G, Eymard B et al (2002) Lamin A/C mutations with lipodystrophy, cardiac abnormalities, and muscular dystrophy. Neurology 59:620–623 66. Kirschner J, Brune T, Wehnert M et al (2005) p.S143F mutation in lamin A/C: a new phenotype combining myopathy and progeria. Ann Neurol 57:148–151 67. Sabatelli P, Squarzoni S, Petrini S et al (1998) Oral exfoliative cytology for the non-invasive diagnosis in X-linked Emery-Dreifuss muscular dystrophy patients and carriers. Neuromuscul Disord 8:67–71 68. Wheeler MA, Warley A, Roberts RG et al (2010) Identification of an emerin-beta-catenin complex in the heart important for intercalated disc architecture and beta-catenin localisation. Cell Mol Life Sci 67:781–796 69. Ognibene A, Sabatelli P, Petrini S et al (1999) Nuclear changes in a case of X-linked EmeryDreifuss muscular dystrophy. Muscle Nerve 22:864–869 70. Maraldi NM, Lattanzi G, Capanni C et al (2006) Laminopathies: a chromatin affair. Adv Enzyme Regul 46:33–49 71. Zhang Q, Bethmann C, Worth NF et al (2007) Nesprin-1 and -2 are involved in the pathogenesis of Emery Dreifuss muscular dystrophy and are critical for nuclear envelope integrity. Hum Mol Genet 16:2816–2833 72. Wilson KL, Berk JM (2010) The nuclear envelope at a glance. J Cell Sci 123:1973–1978 73. Zhang J, FelderA, LiuY et al (2010) Nesprin 1 is critical for nuclear positioning and anchorage. Hum Mol Genet 19:329–341 74. Hoffmann K, Dreger CK, Olins AL et al (2002) Mutations in the gene encoding the lamin B receptor produce an altered nuclear morphology in granulocytes (Pelger-Huet anomaly). Nat Genet 31:410–414 75. Cohen TV, Klarmann KD, Sakchaisri K et al (2008) The lamin B receptor under transcriptional control of C/EBPepsilon is required for morphological but not functional maturation of neutrophils. Hum Mol Genet 17:2921–2933 76. Waterham HR, Koster J, Mooyer P et al (2003) Autosomal recessive HEM/Greenberg skeletal dysplasia is caused by 3 beta-hydroxysterol delta 14-reductase deficiency due to mutations in the lamin B receptor gene. Am J Hum Genet 72:1013–1017 77. Hellemans J, Preobrazhenska O, Willaert A et al (2004) Loss-of-function mutations in LEMD3 result in osteopoikilosis, Buschke-Ollendorff syndrome and melorheostosis. Nat Genet 36:1213–1218 78. Puente XS, Quesada V, Osorio FG et al (2011) Exome sequencing and functional analysis identifies BANF1 mutation as the cause of a hereditary progeroid syndrome. Am J Hum Genet 88:650–656

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124. Naetar N, Hutter S, Dorner D et al (2007) LAP2alpha-binding protein LINT-25 is a novel chromatin-associated protein involved in cell cycle exit. J Cell Sci 120:737–747 125. Bakay M, Wang Z, Melcon G et al (2006) Nuclear envelope dystrophies show a transcriptional fingerprint suggesting disruption of Rb-MyoD pathways in muscle regeneration. Brain 129:996–1013 126. Marji J, O’Donoghue SI, McClintock D et al (2010) Defective lamin A-Rb signaling in Hutchinson-Gilford Progeria Syndrome and reversal by farnesyltransferase inhibition. PLoS One 5:e11132 127. Viteri G, Chung YW, Stadtman ER (2010) Effect of progerin on the accumulation of oxidized proteins in fibroblasts from Hutchinson Gilford progeria patients. Mech Ageing Dev 131:2–8 128. Richards SA, Muter J, Ritchie P et al (2011) The accumulation of unrepairable DNA damage in laminopathy progeria fibroblasts is caused by ROS generation and is prevented by treatment with N-acetyl cysteine. Hum Mol Genet 20:3997–4004 129. Liu Y, Wang Y, Rusinol AE et al (2008) Involvement of xeroderma pigmentosum group A (XPA) in progeria arising from defective maturation of prelamin A. FASEB J 22:603–611 130. Liu B, Wang J, Chan KM et al (2005) Genomic instability in laminopathy-based premature aging. Nat Med 11:780–785 131. Manju K, Muralikrishna B, Parnaik VK (2006) Expression of disease-causing lamin A mutants impairs the formation of DNA repair foci. J Cell Sci 119:2704–2714 132. Liu Y, Rusinol A, Sinensky M et al (2006) DNA damage responses in progeroid syndromes arise from defective maturation of prelamin A. J Cell Sci 119:4644–4649 133. Vousden KH, Lane DP (2009) p53 in health and disease. Nat Rev Mol Cell Biol 8:275–283 134. Decker ML, Chavez E, Vulto I et al (2009) Telomere length in Hutchinson-Gilford progeria syndrome. Mech Ageing Dev 130:377–383 135. Raz V, Vermolen BJ, Garini Y et al (2008) The nuclear lamina promotes telomere aggregation and centromere peripheral localization during senescence of human mesenchymal stem cells. J Cell Sci 121:4018–4028 136. Kudlow BA, Stanfel MN, Burtner CR et al (2008) Suppression of proliferative defects associated with processing-defective lamin A mutants by hTERT or inactivation of p53. Mol Biol Cell 19:5238–5248 137. Wallis CV, Sheerin AN, Green MH et al (2004) Fibroblast clones from patients with Hutchinson-Gilford progeria can senesce despite the presence of telomerase. Exp Gerontol 39:461– 467 138. Du X, Shen J, Kugan N et al (2004) Telomere shortening exposes functions for the mouse Werner and Bloom syndrome genes. Mol Cell Biol 24:8437–8446 139. Moreau F, Boullu-Sanchis S, Vigouroux C et al (2007) Efficacy of pioglitazone in familial partial lipodystrophy of the Dunnigan type: a case report. Diabetes Metab 33:385–389 140. Gambineri A, Semple RK, Forlani G et al (2008) Monogenic polycystic ovary syndrome due to a mutation in the lamin A/C gene is sensitive to thiazolidinediones but not to metformin. Eur J Endocrinol 159:347–353 141. Gotic I, Schmidt WM, Biadasiewicz K et al (2010) Loss of LAP2 alpha delays satellite cell differentiation and affects postnatal fiber-type determination. Stem Cells 28:480–488 142. Parnaik VK, Manju K (2006) Laminopathies: multiple disorders arising from defects in nuclear architecture. J Biosci 31:405–421 143. Gotic I, Leschnik M, Kolm U et al (2010) Lamina-associated polypeptide 2alpha loss impairs heart function and stress response in mice. Circ Res 106:346–353 144. Worman HJ (2006) Inner nuclear membrane and regulation of Smad-mediated signaling. Biochim Biophys Acta 1761:626–631 145. Van Berlo JH, Voncken JW, Kubben N et al (2005) A-type lamins are essential for TGF-beta1 induced PP2A to dephosphorylate transcription factors. Hum Mol Genet 14:2839–2849 146. Arimura T, Helbling-Leclerc A, Massart C et al (2005) Mouse model carrying H222P-Lmna mutation develops muscular dystrophy and dilated cardiomyopathy similar to human striated muscle laminopathies. Hum Mol Genet 14:155–169

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147. Muchir A, Shan J, Bonne G et al (2009) Inhibition of extracellular signal-regulated kinase signaling to prevent cardiomyopathy caused by mutation in the gene encoding A-type lamins. Hum Mol Genet 18:241–247 148. Favreau C, Delbarre E, Courvalin JC et al (2008) Differentiation of C2C12 myoblasts expressing lamin A mutated at a site responsible for Emery-Dreifuss muscular dystrophy is improved by inhibition of the MEK-ERK pathway and stimulation of the PI3-kinase pathway. Exp Cell Res 314:1392–1405 149. Muchir A, Pavlidis P, Decostre V et al (2007) Activation of MAPK pathways links LMNA mutations to cardiomyopathy in Emery-Dreifuss muscular dystrophy. J Clin Invest 117:1282– 1293 150. Lin F, Morrison JM, Wu W, Worman HJ (2005) MAN1, an integral protein of the inner nuclear membrane, binds Smad2 and Smad3 and antagonizes transforming growth factor-beta signaling. Hum Mol Genet 14(3):437–45 (Epub 2004 Dec 15) 151. González JM, Navarro-Puche A, Casar B et al(2008) Fast regulation of AP-1 activity through interaction of lamin A/C, ERK1/2, and c-Fos at the nuclear envelope. J Cell Biol 183:653–666 152. Ugalde AP, Ramsay AJ, de la Rosa J et al (2011) Aging and chronic DNA damage response activate a regulatory pathway involving miR-29 and p53. EMBO J 30:2219–2232 153. Scaffidi P, Misteli T (2008) Lamin A-dependent misregulation of adult stem cells associated with accelerated ageing. Nat Cell Biol 10:452–459 154. Liu GH, Barkho BZ, Ruiz S, Diep D, Qu J, Yang SL, Panopoulos AD, Suzuki K, Kurian L, Walsh C, Thompson J, Boue S, Fung HL, Sancho-Martinez I, Zhang K (2011) Yates J 3rd, Izpisua Belmonte JC. Recapitulation of premature ageing with iPSCs from HutchinsonGilford progeria syndrome. Nature 472:221–225 155. Pekovic V, Hutchison CJ (2008) Adult stem cell maintenance and tissue regeneration in the ageing context: the role for A-type lamins as intrinsic modulators of ageing in adult stem cells and their niches. J Anat 213:5–25 156. Markiewicz E, Tilgner K, Barker N et al (2006) The inner nuclear membrane protein emerin regulates beta-catenin activity by restricting its accumulation in the nucleus. EMBO J 25:3275–3285 157. Hernandez L, Roux KJ, Wong ES et al (2010) Functional coupling between the extracellular matrix and nuclear lamina by Wnt signaling in progeria. Dev Cell 19:413–425 158. Mario G, Ugalde AP, Fernández AF et al (2010) Insulin-like growth factor 1 treatment extends longevity in a mouse model of human premature aging by restoring somatotroph axis function. Proc Natl Acad Sci U S A 107:16268–16273 159. Dechat T, Pfleghaar K, Sengupta K et al (2008) Nuclear lamins: major factors in the structural organization and function of the nucleus and chromatin. Genes Dev 22:832–853 160. McNally EM (2007) New approaches in the therapy of cardiomyopathy in muscular dystrophy. Annu Rev Med 58:75–88 161. Capeau J, Magre J, Caron-Debarle M et al (2010) Human lipodystrophies: genetic and acquired diseases of adipose tissue. Endocr Dev 19:1–20 162. de Pablos-Velasco P (2010) Pioglitazone: beyond glucose control. Expert Rev Cardiovasc Ther 8:1057–1067 163. Varela I, Pereira S, Ugalde AP et al (2008) Combined treatment with statins and aminobisphosphonates extends longevity in a mouse model of human premature aging. Nat Med 14:767–772 164. Osorio FG, Navarro CL, Cadi˜nanos J et al (2011) Splicing-directed therapy in a new mouse model of human accelerated aging. Sci Transl Med 3:106–107

Chapter 19

Desmin and Heart Disease J. Scott Pattison and Jeffrey Robbins

Abstract Desmin is the major intermediate filament protein in muscle cells and is one of the earliest myogenic markers of skeletal and cardiac muscle differentiation. The muscle-specific expression of desmin is regulated by a unique combination of transcriptional and epigenetic controls. Desmin protein functions in the maintenance of myofibril organization and structural and functional integrity of muscle. Loss-of-function studies in mice suggest an additional role for desmin in mitochondrial distribution, morphology, and function. Mutations in desmin or its chaperone protein αB-crystallin are associated with myopathy of skeletal and cardiac muscle. The “desminopathies” are characterized by the accumulation of intracellular desmin-positive protein inclusions and gain of function transgenic models show that mutations in desmin and αB-crystallin are autosomal dominant and sufficient to recapitulate the most important aspects of the human disease. Further study of these models has revealed that a large number of pathological processes are involved in disease development, which may provide starting points for targeting future therapeutic interventions.

19.1

Introduction

Desmin is a muscle-specific cytoskeletal protein found in smooth, skeletal, and cardiac muscles and is a major component of the intermediate filaments found in these muscles. “Intermediate” filaments are defined based on their diameters being approximately 10 nm in diameter, which is intermediate between that of the actin thin filaments and the myosin thick filaments [1]. Desmin forms a threedimensional cytoskeletal network around the myofibrillar Z-discs and longitudinally connects the Z-discs of adjacent sarcomeres, interlinking neighboring myofibrils. Desmin also maintains spatial relationships between the contractile apparatus and the plasma membrane, nucleus, subsarcolemmal cytoskeleton, mitochondria, and other organelles. Thus, the desmin intermediate filament system serves not only J. Robbins () · J. S. Pattison Division of Molecular Cardiovascular Biology, Department of Pediatrics, Cincinnati Children’s Hospital Medical Center, 240 Albert Sabin Way, MLC 7020, Cincinnati, OH 45229-3039, USA e-mail: [email protected]

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to maintain cellular integrity and organization but is also functionally linked to mechanotransduction, mitochondrial respiration, and signal transduction. Desmin was initially discovered in 1976 during the purification of intermediate filament protein from muscle [2, 3]. Upon biochemical purification, antibodies were produced to the purified protein, and the subsequent staining revealed the spatial arrangement and structure of the filament in muscle. Even at this early investigative stage, it was apparent that the filament was intimately associated with the intercalated disc, where the filament interconnected individual myofibers with one another and the plasma membrane. Lazarides and Hubbard named the protein desmin, (delta epsilon sigma mu os (δεσ μós) meaning link or bond [2]).

19.2

Genetics of Desmin

The desmin gene (DES) was originally identified in chicken by Capetanki et al. in 1984 [4], with the human desmin gene isolated and characterized in 1989 [5]. Desmin is encoded by a single copy gene, 8.4 kbp in length, spanning nine exons and eight introns. In situ hybridization localized DES to human chromosome position 2q35 [5, 6]. Northern blot analysis revealed that a single desmin transcript of 2.2 kb is expressed in human striated muscle, which generates a 55 kDa protein (470 amino acids, [2, 3, 5]). The desmin gene is highly conserved from zebrafish to man (Fig. 19.1, [7]). The human desmin gene promoter contains a positive regulatory element located −973 to −693 bp upstream of the transcriptional start site, with a muscle-specific enhancer that is essential for high-level expression [8]. The human gene contains two independent enhancer elements, one active in myotubes and another active in myoblasts. The myotube-specific region contains two E-box elements, corresponding to one MyoD-binding site and one MEF2-binding site, respectively [9]. Downstream of the myotube enhancer region is a myoblast-specific region between −698 and −228, containing a region with homology to the M-CAT motif (at −587) and multiple GC boxes, with consensus binding sites for Krox factors [9]. This unique means of promoter regulation may suggest why desmin is active in myoblasts while other contractile genes are not. Transgenic mice with the promoter containing these regulatory regions, the desmin gene and a LacZ reporter exhibited no transgene expression in cardiac or smooth muscle, suggesting the regulatory sequences are limited to skeletal muscle expression and that other regions are required for other striated muscle expression [10]. A single MEF2C site appears to be necessary for both cardiac and skeletal muscle expression [11], with a distal CarG/octamer element (−4005 to −2495) required for smooth muscle transcriptional activity. Desmin is known as one of the earliest myogenic markers. In 2006, Tam et al. identified a locus control region (LCR) 18-kb upstream of the transcriptional start site of desmin [12]. Locus control regions are transcriptional regulatory elements that impart an open structure to chromatin that permits transcriptional access. Another gene immediately downstream, SPEG (striated muscle preferentially expressed

Fig. 19.1 Multiple sequence alignment with DbClustal software shows the relative conservation of desmin protein sequence from humans to zebrafish. Individual amino acids have colored background (top) and the degree of conservation is indicated by yellow bars (bottom)

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protein kinase), also demonstrates muscle-specific expression and may be subject to high level, coordinate control mediated through the LCR. These initial studies were extended to an epigenetic analysis of a 500-kb region encompassing DES and its LCR [13]. The data showed that the LCR and transcriptional start site were enriched with hyperacetylated domains of histones H3 and H4. In mouse embryonic development, desmin is first detected at 8.25 days postcoitum (p.c.), in the ectoderm and then in the heart rudiment at 8.5 p.c. with increased expression in cardiogenic cells thereafter [14]. Desmin protein begins to accumulate in somites at 9 days p.c. and progresses in a rostro-caudal gradient with somatic maturation [14]. By day 12 p.c., desmin-positive fibers can be observed in limb buds. The levels of desmin expression remain high throughout subsequent embryogenesis and into postnatal life in skeletal, cardiac, and smooth muscles [10]. Desmin is one of the earliest muscle-specific proteins expressed, with substantial accumulations present before titin, skeletal muscle actin, and the myosin heavy chains begin to form the contractile apparatus [15–17].

19.3

Desmin Protein Structure

Characterization of the desmin protein revealed an α-helical head domain, a series of seven peptide (heptad) repeats, and linker positions similar to other type III intermediate filament proteins. It shares high homology with other type III intermediate filament proteins, which include vimentin, glial fibrillary acidic protein, and peripherin. The head domain of desmin has several putative phosphorylation sites while the rod domain is characterized by the conservation of hydrophobic amino acids [18, 19]. Desmin has a central rod domain with four helical regions (1A, 1B, 2A, and 2B), separated by linker regions (L1, L12, and L2) and globular end domain, a structure conserved for the other intermediate filament proteins as well (Fig. 19.2, [20]). The central rod domains function in polymerization by lateral association as intermediate filaments associate laterally in pairs, which then associate with each other to form a protofilament [20, 21]. The standard 10-nm filament is formed by eight individual protofilaments [20]. As noted above, desmin is a type III intermediate filament, with the ability to form homopolymers with itself or heteropolymers with other type III intermediate filaments defining this subset. In smooth muscle cells, desmin is ubiquitous but is concentrated around and between dense bodies. Dense bodies are contractile centers that contain α-actinin and actin in close association with desmin. In adult skeletal muscle, desmin is located throughout the cell but is concentrated at the Z-lines and in the costameres. Costameres are composed of multiprotein complexes that link the intermediate filaments and actin microfilaments to the cell’s membrane [22]. Thus, desmin connects myofibrils to the costameres, forming an extramyofibrillar cytoskeletal network that is also linked to the cell’s periphery [7, 23]. Immunofluorescent staining with the appropriate antibodies showed that desmin not only forms interconnections between the Z-discs in series within each sarcomere, but also forms a link between the

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Fig. 19.2 Desmin protein is organized with a highly conserved α-helical rod domain of 303 amino acids flanked by globular N- and C-terminal structures (known as the head and tail domains). The helical rod structure is interrupted by linker regions (L1, L12, and L2), resulting in four α-helical segments 1A, 1B, 2A, and 2B. Desmin intermediate filaments form dimers through the interaction of rod domains in parallel Fig. 19.3 Desmin (red) shows a striated pattern of Z-line staining and more dense bands of staining at the intercalated discs in healthy heart tissue. Cytoplasmic CryAB counterstaining shown in green

terminal Z-disc and junctional membrane. This suggests that desmin comprises a mechanotransduction connection between the Z-disc and membrane. In cardiac cells, desmosomes tether cardiomyocytes laterally while desmin intermediate filaments surround the Z-discs, tethering together adjacent cells and forming an intercalated disk (Fig. 19.3, [24, 25]). Thus, desmin forms mechanical integration centers in all three muscle types.

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Desmin associates with a variety of other cellular structures including the sarcoplasmic reticulum, nucleus, and mitochondria [24, 26, 27]. These associations have led to the hypothesis that desmin may play a role in conserving a defined spatial distribution of interacting organelles and that this distribution has functional but as of yet undefined consequences. As mislocalization of organelles is correlated with abnormal morphology and function, desmin may contribute to the cellular integrity of muscle cells by functioning as a scaffolding network, keeping the contractile apparatus in register and even functioning in the defined interactions with the mitochondria such that the energy production is positioned in proximity to the sarcomeres that are consuming the ATP [28]. As part of a dynamic and responsive network, it is not surprising that desmin is subject to posttranslational modification. Higher levels of desmin phosphorylation are observed with cardiomyopathy [29]. Desmin is a substrate for several protein kinases including PKA, PKC, Cdc2 kinase, PAK, Aurora-B, and Rho-associated kinase [18, 30–35]. Phosphorylation assays done in vitro and on skinned fibers showed that desmin can be phosphorylated by PKCa at 4 sites: Ser-12, Ser-29, Ser-38, and Ser-56 [29, 32]. Use of the PKC-activating phorbol ester, 12-O-tetradecanylphorbol13-acetate (TPA) caused desmin filament disassembly in hamster cardiac cells, suggesting that PKC-mediated phosphorylation might contribute to the desmin disarray characteristic for a number of cardiomyopathies [29]. Similarly, PAK and Cdc2 kinases phosphorylate desmin at Ser-6, Ser-22, and Thr-64, which resulted in desmin depolymerization [33, 36]. Rho-kinase phosphorylates desmin at Thr-16, Thr-75, and Thr-76 and the phosphorylated desmin was compromised in its ability to form filaments in vitro [33]. Aurora-B phosphorylates desmin in vitro at Thr-16 and Ser59 and phospho-ablated desmin mutants caused defects in filament separation [35]. While desmin phosphorylation is generally associated with filament disassembly or depolymerization, it has been proposed that desmin phosphorylation and dephosphorylation by protein phosphatase 1 may antagonize each other in constant competition to regulate desmin polymerization and function [30]. However, to causally establish the role of desmin phosphorylation in cardiac disease, transgenic or knock-in mice, in which the relevant sites are either rendered nonphosphorylatable or replaced with a charged amino acid to create a phosphomimetic, need to be created and studied.

19.4

Desminopathies

The first realization that desmin was involved in cardiac disease came from the study of a familial cardiomyopathy, in which the heart contained deposits of filaments [37]. Desmin-related myopathy (DRM) was initially described as a skeletal and cardiac myopathy characterized by accumulations containing desmin within myofibers [38]. Defining desminopathy this way linked a number of clinically and pathologically similar muscle disorders. Genetic analyses of these disorders revealed that a portion were due to mutations in the desmin gene [39, 40]. A second form of DRM is caused by mutations in the molecular chaperone αB-crystallin (CryAB, [41, 42]). CryAB

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functions as a chaperone that, under stressed conditions, binds to and stabilizes desmin, preventing its misfolding and subsequent aggregation. In the human diseases, pathological manifestations are indistinguishable between the mutations in desmin or CryAB, although the relevant mouse models show clear differences in the severity, onset, and detailed pathological paths elicited by the desmin versus CryAB mutations [43–45]. Disease presentation is quite heterogeneous, depending on the type of inheritance and specific mutation, with some patients presenting only in skeletal muscle while, in others, clinical presentation of the disease is restricted to the heart [46, 47]. Eighty percent of desminopathy mutations are autosomal dominant, 6% are autosomal recessive, and 2–3% arise as de novo mutations in desmin [48, 49]. There are 45 known DRM-causing mutations in desmin (reviewed by Goldfarb et al. in 2009, [48]). Three mutations in the CryAB gene have been shown to cause DRM [48]. Mutations in desmin and CryAB genes account for approximately one-third of all DRM cases, suggesting mutations in other genes encoding desmin-interacting proteins may exist. The clinical traits of desminopathy patients are characterized by progressive skeletal myopathy, which develops in about one-quarter of patients [48]. Muscle weakness initially presents in the distal leg muscles and progresses to more proximal muscles. In advanced cases, all four limbs are affected, with weakness and atrophy in the trunk, neck, and facial muscles. Skeletal muscle biopsies show cytoplasmic accumulation of amorphous deposits. Loss of respiratory muscle function is a major cause of disability and mortality [48]. Atrioventricular conduction abnormalities are common with desminopathy patients routinely requiring pacemaker implantation [49]. Nearly 25% of desminopathy patients present with cardiomyopathy, with no skeletal involvement [49]. Heart disease is observed in > 60% of patients overall [48]. Desmin mutations show a wide range of pathogenic effects and the mechanisms leading to variable disease expression requires further study with new models and technologies.

19.5

Desmin Loss of Function

Loss of function can oftentimes illuminate cryptic functions of the protein in question. Because of its presumed importance in muscle stability, DES was a relatively early target for gene ablation. In 1996, two groups generated desmin null mice by homologous recombination [50, 51]. The desmin knockout mice were viable and fertile. Desmin-deficient skeletal muscle showed several architectural and ultrastructural defects, with regions of myofibrillar disorganization, fibrosis, and degeneration [52]. However, the loss of desmin was most phenotypically pronounced in the heart, with evidence of calcification and fibrosis of the myocardium at 10 weeks of age by Von Kossa and Masson’s trichrome staining, respectively [50]. Ultrastructural examination of the desmin-ablated hearts showed myofibril disorganization, with abnormal nuclear and mitochondrial morphology and positioning [52]. The desmin null hearts were characterized by impaired exercise performance, mild hypertrophy (∼20%), and age-dependent loss of systolic function [53, 54]. Using the Langendorff technique, the desmin knockout hearts were found to have increased diastolic pressure

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with reduced developed pressure, suggesting impairment in myocardial force generation [55]. Further interrogation of the mitochondrial defects in desmin-deficient mice found mitochondrial clumping and proliferation, accompanied by reduced ADPstimulated respiration in situ using saponin skinned fibers. However, no differences were detected in isolated mitochondria from mice with or without desmin [56]. Desmin-deficient hearts also showed a threefold increase in creatine kinase activity as compared to controlled hearts, but this may simply reflect a compensatory response to other mitochondrial defects [57]. To explore functional correlates between the altered mitochondrial distribution and cell death through the intrinsic apoptotic pathway, Weisleder et al. crossed desmin knockout mice with Bcl-2 overexpressing transgenic mice [58]. Bcl-2 is an integral outer mitochondrial membrane protein that protects some cell types from apoptosis without affecting mitochondrial function. With high levels of Bcl-2 expression, desmin-deficient hearts failed to develop calcification or fibrosis. Bcl-2 overexpression also restored mitochondrial organization and morphology. Desmin null mitochondria display increased sensitivity to calcium induced evidenced by mitochondrial swelling, which was reversed by Bcl-2 overexpression [58]. These data suggest that the lack of desmin leads to mitochondrial sensitivity to calcium, which can be ameliorated by Bcl-2 expression through some undefined mechanism.

19.6

Desmin Transgenic Mice

Desmin is upregulated in a number of cardiac disorders including cardiac hypertrophy and congestive heart failure [59–61]. To test whether desmin overexpression contributed to cardiac disease, Wang et al. created transgenic mice with cardiacspecific expression of wild-type desmin [45]. Hearts expressing desmin at threefold over endogenous protein levels were phenotypically normal and had normal life spans [62]. These data support the hypothesis that excess desmin production is not deleterious to the heart. Because the majority of the desmin- and CryAB-based DRM mutations are autosomal dominant, they can be modeled using tissue-directed transgenic expression of the mutated protein(s). Overall, two overarching facts are widely accepted: (1) mutations result in a loss of desmin function and (2) misfolded proteins, including desmin, progressively accumulate in intracellular aggregates, which impair contractile function and predispose myocytes to death. It is this accumulation and pathological aggregation that is thought to underlie the basic disease process and not a simple, loss of function of the protein’s activity. A human patient with severe myopathy and intrasarcoplasmic inclusions of desmin filaments was found to have a seven amino acid deletion (R173-E179, [40]). This deletion corresponded to segment 1B of the rod domain of desmin, which is believed critical for desmin self-polymerization. To test whether this deletion was sufficient to cause DRM cardiomyopathy, Wang et al. created transgenic mice with cardiacspecific expression of DesminR173−E179 (Des7 mice, [45]). It is important to note

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that in these mice the heart continued to express normal amounts of normal desmin protein as well, with the transgenic expression of the mutant desmin superimposed upon the normal protein. Des7 mice developed aberrant intrasarcoplasmic desmin aggregates, with abnormal myofibril alignment and impaired cardiomyocyte contractile function, recapitulating characteristics of human DRM [45]. By 8 months of age, Des7 mice showed impaired cardiac function with decreased rates of contractility and relaxation. These data prove that cardiomyocyte-autonomous expression of the desmin mutation is sufficient to cause DRM. In many cases, as the heart disease progresses, the desmin organization and localization visibly changes and desmin-containing aggregates accumulate intracellularly. These processes are often accompanied by the upregulation of apoptosis leading to the hypothesis that desmin may be an apoptotic target. Chen et al. found that using TNFα to induce apoptosis in myogenic cells, desmin was specifically cleaved by caspase-6 at Asp-263 [63]. Caspase-6 cleavage abolished desmin’s ability to assemble into intermediate filaments and resulted in the accumulation of large intracellular aggregates that contained the misfolded protein. These findings were further validated in vivo, when transgenic mice with cardiac-specific expression of TNFα were crossed with mice expressing mutant desmin D263E in a desmin-null background. DesD263E mice were resistant to caspase cleavage, failed to form aggregates and rescued TNFα-induced cardiac dysfunction. The data, while limited, are intriguing and imply that desmin cleavage may be sufficient to cause aggregation and contribute to cardiac dysfunction under conditions where apoptosis is actively occurring [64]. In the heart, the CryAB functions as a molecular chaperone and promotes proper folding of desmin and actin as well as maintenance of their conformation during stress [65]. CryAB upregulation and accumulation is observed in a number of cardiac disorders including desminopathy, dilated cardiomyopathy, and familial hypertrophic cardiomyopathy [66, 67]. A mutation in the αB-crystallin gene is linked to DRM. Patients from a large French family—presenting with proximal and distal muscle weakness, hypertrophic cardiomyopathy, and lens opacity—carried a missense Arg120Gly mutation in αB-crystallin (CryABR120G , [41, 42]). To determine whether CryAB or CryABR120G expression was sufficient to cause desmin-related cardiomyopathy, Wang et al. created transgenic mice with cardiac-specific expression of CryABWT and CryABR120G [44]. Mice expressing high levels of wild-type CryAB were normal while even modest levels of CryABR120G expression led to the formation of intracellular aggregates of desmin and CryAB protein. CryABR120G hearts also displayed aberrant desmin networks, myofibril disorganization, progressive cardiac hypertrophy, and loss of cardiac function leading to dilation and heart failure [44]. These data showed that the CryABR120G mutation is sufficient to cause DRM. Subsequent publications have better characterized the CryABR120G pathology and begun to unravel the biological mechanisms contributing to the associated cardiomyopathy. CryABR120G expression leads to an upregulation and accumulation of ubiquitinated proteins and, at least initially, is accompanied by an increase in proteasomal activity, suggesting a misfolded protein overload of the ubiquitin-proteasomal

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system. Maloyan et al. demonstrated that the mitochondrial dysfunction and apoptosis are increased with CryABR120G expression [68–70]. Additional publications demonstrated that CryABR120G expression leads to the formation and accumulation of preamyloid oligomers, which are toxic, soluble, conformer-specific protein intermediates associated with neurodegenerative disease [43, 71–74]. Rajasekaran et al. suggest a role for reductive stress in CryABR120G -induced cardiac hypertrophy and aggregate accumulation. Recent data suggest a significant role for autophagy regulation in development of CryABR120G pathology [70, 75]. In total, these data suggest that CryABR120G expression initiates a multifaceted series of pathological processes, whose absolute and relative contributions may differ along the longitudinal course of the disease.

19.7

Conclusions

Intermediate filament proteins have a broad range of functions and interactions. The muscle-specific intermediate filament protein, desmin, plays an integral role in myocyte structural and functional integrity. Mutations in the desmin and αB-crystallin genes are associated with pathology of the skeletal and cardiac muscle, characterized by the accumulation of intracellular aggregates and loss of skeletal muscle and cardiac function, called desminopathy. Loss of desmin function demonstrates the critical role desmin plays in myofibril organization as well as mitochondrial distribution and function. Transgenic mouse models recapitulating the human pathology show that numerous pathological mechanisms are activated in desminopathy. Due to the complex and multifocal nature of the pathological processes underlying desminopathy, continuing research must focus on defining the early/primary mechanisms causing the disease so that therapeutic targets can be identified. Acknowledgments The research in the authors’ laboratory is supported by National Institutes of health (NIH) grants P01HL69799, P50HL074728, P50HL077101, P01HL059408, and R01HL087862 (to J.R.). National Institutes of Health fellowship awards T32 HL07752 and F32 HL087478; and an American Heart Association Postdoctoral Fellowship (to JSP).

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Chapter 20

Neurodegenerative Diseases and Intermediate Filaments Rodolphe Perrot and Jean-Pierre Julien Abstract Intermediate filaments (IFs) represent the most abundant cytoskeletal constituent in mature neurons. Their mutations and/or accumulations are associated with many human neurodegenerative disorders and it is now well established that disorganization of the intermediate filament network may be directly involved in neurodegeneration. Diseases caused by intermediate filament abnormalities show a wide range of phenotypes, depending on many factors, including the class of affected intermediate filaments and the type of mutation. Various mouse models were extensively used to provide a better understanding of the role played by the disorganization of intermediate filaments in the pathogenesis of neurodegenerative disorders. However, the mechanisms leading to the formation of these aggregates often remain elusive. Multiple factors can potentially induce the accumulation of neuronal intermediate filaments, including dysregulation of intermediate filament gene expression, intermediate filament mutations, defective axonal transport, abnormal posttranslational modifications, and/or proteolysis. Here, we review some neurodegenerative diseases involving intermediate filament abnormalities and possible mechanisms susceptible to provoke them.

20.1

Introduction

Similar to most eukaryotic cells, neuronal and glial cytoskeletons are composed of three interconnected structures: actin microfilaments (MFs), microtubules (MTs), and intermediate filaments (IFs). IF proteins are classified into six types and neurons express differentially several IF proteins depending on their developing stage or their localization in the nervous system: neurofilament (NF) triplet proteins (called NFL (light, 68 kDa), NFM (medium, 160 kDa) and NFH (heavy, 205 kDa); type IV), α-internexin (66 kDa; type IV), peripherin (57 kDa; type III), nestin (200 kDa; type IV), vimentin (57 kDa; type III) syncoilin isoforms (Sync1 (64 kDa), Sync2 (64 kDa); type III) and synemin isoforms (Low synemin (41 kDa), Middle or beta synemin (150 kDa), and High or α-synemin (180 kDa); type IV). In adult central (CNS) and peripheral nervous system (PNS), IFs are the most abundant cytoskeletal R. Perrot () · J.-P. Julien Institut de Biologie en Santé, Service Commun d’Imageries et d’Analyses Microscopiques, Angers University, CHU, 4 Rue Larrey, 49933 ANGERS Cedex 09, France e-mail: [email protected]

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components of large myelinated axons [1]. They are principally composed of the NF triplet proteins and also α-internexin in CNS and peripherin in PNS [2, 3]. The main role recognized for NFs is to increase the axonal caliber of myelinated axons and consequently their conduction velocity [4]. They also contribute to the dynamic properties of the axonal cytoskeleton during neuronal differentiation, axon outgrowth, regeneration, and guidance [5]. The IF proteins share a common tripartite structure with nonhelical amino and carboxy-terminal regions (called the head and tail domains) flanking a central rod domain of approximately 310 amino acids that contains four α-helical regions and which is involved in the assembly of 10-nm filaments [6]. NFM and NFH subunits differ from other neuronal IF proteins by their long tail domains containing numerous repeats of phosphorylation sites Lys-Ser-Pro (KSP) [4]. An increasing body of evidence supports the view that the most common mechanism of chronic neurodegenerative disorders involves abnormal protein production, processing or misfolding, and the subsequent accumulation in nervous system. Alterations in the metabolism and/or organization of neuronal IFs are frequently associated, directly or indirectly, with various neurodegenerative diseases, including amyotrophic lateral sclerosis (ALS), Charcot-Marie-Tooth disease (CMT), giant axonal neuropathy (GAN), neuronal intermediate filament inclusion disease (NIFID), Parkinson’s disease (PD), diabetic neuropathy, dementia with Lewy bodies, and spinal muscular atrophy [7]. While IF abnormalities in neurodegenerative disorders could simply reflect a pathological consequence of neuronal dysfunction, recent studies using transgenic mouse models suggested that IF disorganization itself can also produce deleterious effects and therefore could contribute to the neurodegeneration process. Glial IF, and more particularly glial fibrillary acidic protein (GFAP) in astrocytes, is also the target of mutations leading to neurodegenerative diseases. Astrocytes express various IF proteins—including nestin, vimentin, and synemin—but GFAP is the most abundant. GFAP is a type III IF protein existing under different spliced forms. The relative abundance of these GFAP transcripts is variable and can be dependent upon astrocyte location or pathological states [8]. GFAP mutations lead to accumulations of GFAP protein and causeAlexander disease (AXD), a rare leukodystrophy. Here, we attempted to cover the current knowledge related to neuronal and glial IF involvement in human neurodegenerative diseases.

20.2

Neuronal Intermediate Filaments and Neurodegenerative Diseases

20.2.1 Amyotrophic Lateral Sclerosis Amyotrophic lateral sclerosis (ALS), often referred to as Lou Gehrig’s disease, is a lethal disease characterized by a progressive degeneration of upper and lower motor neurons leading to weakness and atrophy of the muscles followed by progressive

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paralysis while the cognitive functions of the patients remain usually unaffected. ALS has a worldwide prevalence of 1–2 per 100,000. There is no cure and death usually occurs within 3–5 years from the onset of symptoms. Approximately 90% of ALS cases are sporadic while 10% are inherited in an autosomal dominant pattern. Twenty percent of all the familial cases are due to mutations in Cu/Zn superoxide dismutase 1 (SOD1), the most abundant cytosolic enzyme. The discovery of NF gene variants in ALS patients suggested the involvement of NFs in the pathogenesis of the disease. Indeed, codon deletions or insertions in the KSP repeat motifs of NFH have been identified in a small number of patients with sporadic ALS [9–11]. However, two other studies failed to identify variants in the NF genes linked to sporadic and familial ALS [12, 13], suggesting that mutations in the NF genes are not a systematic common cause of ALS but could be a risk factor for sporadic ALS. Peripherin mutations have also been identified in three sporadic ALS patients [14–16], including a frame-shift mutation in the PRPH gene able to disrupt the NF network assembly in vitro, reinforcing the view that NF disorganization may contribute to pathogenesis. These results suggest that peripherin mutations may be responsible for a small percentage of ALS cases. Two peripherin isoforms are also associated with ALS: aggregate-inducing Per28 is upregulated in patients with ALS, at both the mRNA and protein levels, and is associated with round inclusions in disease pathology [17]. The Per61 splice variant is neurotoxic and has been observed in ALS mouse models and human patients [18]. These observations raise the possibility that missplicing of peripherin could lead to disease. It is also of interest to note the presence of high NFL and NFH levels and autoantibodies against NFL in cerebrospinal fluid of ALS patients [19–21]. Both sporadic and familial ALS are characterized by the presence in motor neurons of axonal spheroids and perikaryal accumulations composed of NFs and/or peripherin [22]. NFs in perikaryal aggregates are extensively phosphorylated, a process that occurs normally only within the axon [23]. The mechanisms governing the formation of IF aggregates are still not clearly established. There is evidence that IF accumulations could results from defects of axonal transport or from abnormal stoichiometry of IF proteins. Perturbations of the axonal transport of NFs and organelles is one of the earliest pathological changes seen in several transgenic mouse models of ALS [24–27]. The premature phosphorylation of NF tail domains in motor neuron cell bodies could directly mediate their accumulation in this region. Glutamate excitotoxicity, another pathogenic process in ALS, may induce abnormal phosphorylation of NFs. Treatment of primary neurons with glutamate activates members of the mitogen-activated protein kinase family which phosphorylate NFs with ensuing slowing of their axonal transport [28]. In addition, glutamate leads to caspase cleavage and activation of protein kinase N1 (PKN1), an NF head-rod domain kinase [29]. This cleaved form of PKN1 disrupts NF organization and axonal transport. Excitotoxicity mediated by non-N-methyl-D-aspartic acid (NMDA) receptor is also associated with the aberrant colocalization of phosphorylated and dephosphorylated NF proteins [30]. Inhibition of Pin1, a prolyl isomerase, was suggested as a possible therapeutic target to reduce pathological accumulation of phosphorylated NFs. Pin1 associates with

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phosphorylated NFH in neurons and is colocalized in aggregates found in the spinal cord of patients with ALS [31]. Its inhibition reduces glutamate-induced perikaryal accumulation of phosphorylated NFH. Finally, it was recently reported that riluzole protects against glutamate-induced slowing of NF axonal transport by decreasing perikaryal NF side-arm phosphorylation [32], probably via the inhibition of ERK and p38 activities, two NF kinases activated in ALS. Alterations of the anterograde or retrograde molecular motors may also be responsible for aggregation of IFs. Mutation of dynein or p150glued [33], overexpression of dynamitin [34] and absence of kinesin heavy chain isoform 5A (KIF5A, [35]) induce NF accumulations in mice. Recent studies suggest that inhibition of retrograde transport is more susceptible to cause accumulation of NFs than inhibition of anterograde transport. The inhibition of dynein by increasing the level of dynamitin induces aberrant focal accumulation of NFs within axonal neurites whereas inhibition of kinesin inhibits anterograde transport but does not induce similar focal aggregations [36]. Similarly, the neuronspecific expression of Bicaudal D2 N-terminus (BICD2-N), a motor-adaptor protein, impairs dynein-dynactin function, causing the appearance of giant NF swellings in the proximal axons [37]. However, these mice did not develop signs of motor neuron degeneration and motor abnormalities. Modification in NF stoichiometry was also proposed to induce accumulation of NFs. The overexpression of different NF subunits in mice provokes the formation of NF aggregates [38, 39, 40]. Remarkably, the motor neuron disease caused by excess human NFH (hNFH) can be rescued by overexpression of hNFL in a dosage– dependent fashion [41]. Overexpression of peripherin in mice also provokes the formation of cytoplasmic protein aggregates and the subsequent selective loss of motor neurons during aging [42, 43]. This loss is preceded by axonal transport defects and formation of axonal spheroids [44]. Because NFL mRNA levels are reduced in cases of ALS, Beaulieu et al. [43] generated double transgenic mice overexpressing peripherin and deficient for NFL (Per; NFL-/- mice). Here, the onset of peripherin-mediated disease is accelerated by the deficiency of NFL. Without NFL, peripherin interacts with NFM and NFH to form disorganized IF structures. This could explain why the number of IF inclusion bodies is increased in Per; NFL-/mice, leading to an earlier neuronal death and to defects of fast axonal transport in cultured Per; NFL-/- neurons [45]. In contrast, peripherin toxicity can be attenuated by coexpression of NFL or NFH [46, 47], illustrating once again the importance of IF protein stoichiometry. NFH overexpression shifted the intracellular localization of inclusion bodies from the axonal to the perikaryal compartment of motor neurons, suggesting that the toxicity of peripherin inclusions may be related to their axonal localization, possibly by altering the axonal transport. However, it should be noted that peripherin is not a key contributing factor to the neuronal death in disease caused by SOD1 mutations because absence or overexpression of peripherin in SOD1G37R mice do not affect the onset and progression of motor neuron disease [48]. ALS motor neurons also show a selective decrease in the levels of NFL, αinternexin, and peripherin mRNA, while in familial ALS the levels of peripherin mRNA appear to be abnormally elevated [49–51] This suggests a change in the stoichiometry of cytoskeletal protein expression which could be conducive to the

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formation of neurofilamentous aggregates in ALS. This decrease of IF mRNA could be due in part to modification in their stability. Several NFL mRNA-binding proteins have been identified in human, including 14-3-3 proteins [52], TAR DNA-binding protein (TDP43) [53], both mutant and wild-type SOD1 [54] and Rho guanine nucleotide exchange factor (RGNEF) [55]. These proteins are incorporated in ALS intraneuronal aggregates and affect the stability of NFL mRNA. In agreement with this, mice expressing human TDP-43 displayed reduced NF mRNAs and protein content, inducing a decrease of caliber of their motor axons [56]. The involvement of TDP-43 in ALS pathogenesis was reinforced by the recent discovery of several mutant forms of this protein in familial and sporadic ALS [57]. Neuronal IF abnormalities in ALS may also occur as a result of posttranslational protein modifications. Indeed, advanced glycation endproducts were detected in NF aggregates of motor neurons in familial and sporadic ALS [58]. Moreover, it was showed that SOD1 can catalyze nitration of tyrosines by peroxynitrite in the rod and head domains of NFL [59]. However, no significant changes were detected in the nitration of NFL isolated from cervical spinal cord tissue of sporadic ALS cases [60]. Finally, it seems that nonneuronal cells could be directly involved in the formation of cytoskeletal aggregates within proximal axon from motor neurons. Indeed, cultured mouse spinal motor neurons in contact with nonneuronal cells displayed swellings that were morphologically and neurochemically comparable to axonal spheroids that develop in vivo in ALS transgenic mouse models [61]. These swellings contained NFL, NFM, NFH, α-internexin and peripherin, and induced the accumulation of mitochondria and vesicle-like structures, suggesting a disruption of the axonal transport. Moreover, the severity of this axonopathy correlated with the phenotype of the glial cells, with a significant increase being induced by a glial feeder layer expressing mutant SOD1 or that was preaged prior to plating the motor neurons [61]. To further determine whether NFs are directly involved in SOD1-mediated disease, mice expressing mutant SOD1 were mated with transgenic mice deficient for axonal NFs. The withdrawal of NFs from the axonal compartment and their perikaryal accumulation induced by the expression of NFH-β-galactosidase fusion protein conferred no beneficial effect to SOD1G37R mice [62], indicating that axonal NFs are not necessary for SOD1-mediated disease. This was also observed in SOD1G85R mice deprived of NFL, but the absence of axonal NFs in these animals prolongs their life span by approximately 15% [63]. Surprisingly, overexpression of mouse NFL or mouse NFH in SOD1G93A mice [64] and overexpression of hNFH in SOD1G37R mice [65] also increase their life span by, respectively, 15 and 65%. This suggests a protective effect of NF perikaryal accumulation in motor neuron disease caused by mutant SOD1. While the mechanism of protection is unclear, it seems that perikaryal accumulation of NFs rather than their axonal deficiency is responsible for slowing disease in these models. Indeed, the formation of large perikaryal aggregates and a massive depletion of axonal NFs due to the expression of the human NFH43 allele cause more positive effects than human NFH44 allele which induces smaller aggregates and more axonal NFs [65]. Moreover, the disruption of one allele for each NF gene induces a 40% decrease of axonal NF proteins content and an important axonal atrophy without perikaryal accumulation of NFs in SOD1G37R mice, but it does not

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extend their life span nor does it alleviate the loss of motor axons [66]. Several hypotheses were proposed to explain this protective effect of perikaryal aggregates in SOD1-mediated disease. Through their multiple calcium-binding sites NFs may act as calcium chelators. Supporting this hypothesis, a significant neuroprotection was obtained by overexpressing the calcium-binding protein calbindin-D28k in cultured motor neurons [67]. It was also proposed that perikaryal accumulations of NFs in motor neurons may alleviate ALS pathogenesis by acting as a phosphorylation sink for cyclin-dependent kinase 5 dysregulation induced by mutant SOD1, thereby reducing the excessive phosphorylation of tau and other neuronal substrates [66]. This was supported by the fact that NF accumulations contain hyperphosphorylated NFM and NFH subunits in ALS patients [23] and in SOD1 mutant mice [68]. However, removal of NFM and NFH sidearms led to a delay of disease in SOD1 mutant mice rather than the acceleration predicted by a kinase dysregulation model [69] indicating that perikaryal phosphorylation of NFs is not an essential contributor to reduced toxicity of SOD1 mutants. Alternatively, axonal removal of NFs could enhance axonal transport, which is impaired in SOD1 mice by providing a more flexible axoplasm. Finally, it was shown that NFs are involved in the localization of NMDA receptors in the neuronal plasma membrane by interacting with the NMDA NR1 subunit [70]. Thus, accumulation of NFs could interfere with glutamate receptor function and prevent glutamate excitotoxicity. However, NF aggregate-bearing neurons demonstrate increased intracellular calcium levels and enhanced cell death in response to NMDA receptor activation without increased NMDA receptor expression. These results suggest that the presence of NF aggregates renders motor neurons more susceptible to NMDA-mediated excitotoxicity [71].

20.2.2

Charcot-Marie-Tooth Disease

Charcot-Marie-Tooth disease (CMT) represents a heterogeneous group of inherited peripheral neuropathies affecting both motor and sensory nerves. With an estimated prevalence of 1 in 2,500 individuals, CMT is the most common inherited disorder of the PNS. CMT is characterized by an insidious onset and slowly progressive weakness and atrophy of the distal limb muscles. First signs typically appear in the first or second decade of life, although it may be detected in infancy. On the basis of electrophysiological properties and histopathology, CMT was originally subclassified into CMT1 and CMT2. CMT1 is a demyelinating disease with reduced nerve conduction velocity whereas CMT2 is an axonal neuropathy with relatively normal nerve conduction velocity. CMT patients show a high degree of heterogeneity, due to mutations in multiple genes. This led to the distinction of other subtypes of CMT, including CMT3 (or Dejerine-Sottas disease, a particularly severe demyelinating form of CMT), CMT4 (autosomal recessive form of demyelinating CMT), and CMTX (X-linked form of CMT with both demyelinating and axonal features). Vogel et al. [72] reported the presence of NF accumulations in CMT. Evidence for the involvement of IFs in the pathogenesis of CMT was provided by the identification of 20 mutations in gene coding for NFL in patients with CMT1F and CMT2E.

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Fig. 20.1 Location of CMT-associated mutations in NFL. Schematic representation of human NFL subunit. The gray boxes correspond to the four α-helical subdomains within the central rod domain, separated by nonhelical linkers. Twenty mutations identified in CMT patients, including deletion, missense, and frame-shift mutations, were located. They are distributed throughout the NFL coding sequence, with a more frequent occurrence in head and rod domains. C carboxy-terminal, N aminoterminal. (Reference UniProtKB/Swiss-Prot database P07196)

Mutations in NEFL gene are responsible for approximately 2% of CMT cases and a high percentage of CMT2 cases. These mutations are located throughout the three functional domains of this protein (head, rod, and tail) and consist of substitutions, deletions, and frame-shift mutations (Fig. 20.1). The first two CMT-associated NEFL mutations, NFLP8R and NFLQ333P , were identified in, respectively, a Belgian and a Russian family. They disrupt NF assembly and axonal transport in vitro [73, 74]. This effect was dominant, since wild-type NFL could not rescue the assembly defect. These mutations also cause the sequestration of mitochondria in cell bodies and proximal axons, the fragmentation of the Golgi apparatus, and the degeneration of neuritic processes in cultured neurons. Filament formation was also abolished in SW13 cells by the rod domain A148V mutation [75]. These data provide possible mechanisms by which these mutants could be involved in axonal degeneration and CMT pathogenesis. The Pro-22 residue of NFL is also the target of several mutations: P22R, P22S, and P22T. The P22R mutation, identified in a Korean family, is associated with demyelinating neuropathy features of CMT1F [76]. The P22S substitution was first described in a Slovenian CMT2 family [77], then in an Italian family developing a primary axonopathy characterized by giant axons with swellings composed essentially of aggregated NFs [78]. Interestingly, clinical and electrophysiological studies from patients with P22S mutation revealed a mixed axonal and demyelinating neuropathy [79], emphasizing the complexity of genotype-phenotype correlations in CMT. Finally, the P22T mutation was detected in unrelated Japanese patients with CMT disease [80]. The formation of NF aggregates in patients expressing NFLP22S and NFLP22T mutant proteins could be explained by the ability of these mutations to abolish the phosphorylation of the adjacent Thr21 which normally inhibits filament assembly [81]. The phosphorylation of NFL head domain by PKA alleviated aggregates in cortical neurons, providing a potential therapeutic approach to dissociate NF aggregates in CMT disease [81].

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The screening of 323 patients with CMT or related peripheral neuropathies allowed the identification of six disease-associated missense mutations and one 3-bp in-frame deletion in the NEFL gene [82]. Other mutations were also detected in Korean CMT patients [83], in a German family [84], and four mutations in the head and rod domains of NFL, including an L268P substitution and a del322Cys_326 Asn deletion, were identified by the screening of 177 patients [85]. Most of these mutated proteins (except E7K and D469N) form aggregates, and thus could alter the axonal transport following their abnormal aggregation in cell bodies and axons. A duplication-insertion mutation of NFL in a patient with CMT was also reported [86], which probably provoked neuronal degeneration through both aggregation and destabilization of the IF network. Finally, new mutations in the NEFL gene were identified following the screening of 223 Japanese CMT patients [87]. Four heterozygous missense mutations (P8L, E90K, N98S, and E396K) were detected in five unrelated patients as well as a homozygous nonsense mutation (E140Stop) in one patient. All these patients displayed moderate delayed nerve conduction velocities, possibly caused by a loss of large diameter fibers. This study suggested that nonsense NEFL mutations probably cause a recessive phenotype, while missense mutations cause a dominant phenotype [87]. The majority of NFL mutations are linked to axonal forms of CMT but their implication in demyelinating CMT cannot be excluded since nerves from patients expressing NFLL268P or NFLE90K showed evidence of Schwann cell abnormalities [82, 85]. The generation of the first mouse model of the CMT2E disease was recently reported [88]. These mice express the hNFLP22S mutant protein specifically in the nervous system and develop a key feature of CMT2E disease at 9 months of age, including aberrant hindlimb posture, motor deficits, hypertrophy of muscle fibres, and loss of muscle innervation without neuronal loss. To address whether CMT2E disease is potentially reversible, this mouse model was based on the tetracycline-responsive gene system that allows the suppression of mutant hNFLP22S expression in mature neurons through administration of doxycycline. Remarkably, a 3-month treatment of these mice with doxycycline after disease onset efficiently downregulated expression of hNFLP22S and reversed the neurological phenotype [88], providing hope that future therapeutic strategies might not only stop progress of CMT2E disease but also reverse it. Mutations of myotubularin-related protein 2 (MTMR2, CMT4B), heat-shock protein B1 (HSPB1, CMT2F), or HSPB8 (CMT2L) can also cause NFL aggregation [89, 90, 91, 92], indicating that mutation of NFs is not the only mechanism inducing their accumulation in CMT. Coexpression of Wt HSPB1 with P8R or Q333P CMT mutant NFL reduced their aggregation, induced reversal of mutant NFL aggregates, and decreased mutant NFL-induced loss of motor neuron viability [93]. On the opposite, mutant HSPB1 has dominant effect on disruption of NF assembly and aggregation of NFL protein. In the same way, mutant MTMR2 induces abnormal NFL assembly in transfected cells [91]. Zhai et al. [93] showed that deletion of NFL markedly reduces degeneration and loss of motor neurons induced by mutant HSPB1. Another study recently showed that expression of NFLP8R or NFLQ333P in cultured motor neurons causes the rounding of mitochondria and reduction of axonal

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Fig. 20.2 Mapping of gigaxonin mutations identified in GAN patients. Schematic representation of human gigaxonin protein. BTB, BACK, and C-terminal Kelch domains are represented. Thirty-three mutations associated with GAN and found in each functional domain were located. C carboxyterminal, N amino-terminal. (Reference UniProtKB/Swiss-Prot database: Q9H2C0)

diameter before disruption of the NF network [94]. Cotransfection of HSPB1 helped to maintain normal NF network, axonal caliber, and mitochondrial morphology. On the other hand, the cotransfection of HSPA1 was effective in neurons expressing NFLQ333P , but not NFLP8R , suggesting that chaperone-based therapies have potential for the treatment of CMT2E but their efficacy would depend on the profile of HSPs induced and the type of NEFL mutation.

20.2.3

Giant Axonal Neuropathy

Giant axonal neuropathy (GAN) is a rare progressive neurodegenerative disorder affecting both PNS and CNS which generally appears in infancy or early childhood. Phenotypic variability has been reported but typical clinical features include distal limb weakness, areflexia, and a marked gait disturbance. As the disorder progresses, CNS involvement includes electroencephalographic abnormalities, mental retardation, seizures, and defective upper motor neuron function. GAN is caused by mutations in the GAN gene encoding the ubiquitously expressed protein gigaxonin. Gigaxonin belongs to a protein family that is characterized by an N-terminal BTB (broad-complex, Tramtrack, and Bric a brac) domain and six kelch repeats [95]. BTB/kelch proteins are organizers of the cytoskeletal network and closely linked to the ubiquitin degradation pathway. More than 35 mutations of the gigaxonin have been identified to date (Fig. 20.2). They are localized throughout the GAN gene and are thought to lead a loss of function of the encoded protein. The major cytopathological hallmark of GAN is the presence of masses of neuronal IFs producing focal enlargements in the distal regions of axons associated with a reduced number of MTs [96]. In contrast, axonal segments proximal to the swellings exhibit a reduced number of NFs [97]. Disorganization and accumulation of other types of IFs are also found in skin fibroblasts, Schwann cells, and muscle fibers [98–100], suggesting a critical role of gigaxonin in IF organization. A decreased inter-NF distance was observed in sural nerve axons of a GAN patient and, more

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surprisingly, the mean diameter of NFs was increased (12.4 nm in GAN compared with 10.1 nm in controls) [101]. Although the mechanism leading to the distal axonal accumulation of NFs is still unclear, an acceleration of their axonal transport was observed in optic nerve from experimentally induced GAN rat model, concomitant with a proximal decreased content of NFs and their distal accumulation [102]. The authors proposed that acceleration of NF transport in the presence of a normal rate of NF protein synthesis and insertion into transport system would lead to the formation of distal axonal swellings with packed NFs. Mice deleted in exons 3–5 of the GAN gene were produced in order to determine how loss of gigaxonin’s function leads to GAN [103]. Despite the development of a progressive deterioration for motor function, these animals displayed normal life span and fertility, and giant axons were never seen. Nevertheless, these mice exhibited enlarged axons with densely packed NF, leading to the segregation of axonal organelles, a feature characteristic of human GAN pathology. This was accompanied by an axonal loss at the age of 9–12 months. However, it should be noted that some null mice showed no overt neurological phenotypes, suggesting that some genetic modifiers may exist [104]. Another mouse model with deletion of exon 1 of the GAN gene was generated [105], which exhibited no overt phenotype and no giant axons but developed accumulations of IF proteins. These aggregates were composed of nonphosphorylated NFH and α-internexin and formed preferentially in the cerebral cortex and thalamus. Small aggregates of NFL and peripherin also formed in cell bodies of dorsal root ganglion neurons. Moreover, increased levels of neuronal IF proteins were detected in various regions of the nervous system, confirming the importance of gigaxonin in modulating the levels and organization of IF proteins. Gigaxonin was proposed to promote the ubiquitin-mediated degradation of microtubule-associated protein 1B (MAP1B, [106]), tubulin-folding cofactor B (TBCB, [107]), and MAP8 [103]. Disease-associated gigaxonin mutations perturb its association with these partners while gigaxonin ablation results in their accumulation [103, 107, 108]. This raised the possibility that IF accumulation in GAN results from an MT reorganization/destabilization. However, it is intriguing to note that these proteins have opposite effects on the MT network: MAP1B is an MTstabilizing phosphoprotein, whereas overexpression of TBCB depolymerizes MTs. Using primary fibroblasts derived from skin biopsies of GAN patients with aberrant aggregates of vimentin, Cleveland et al. [109] demonstrated that gigaxonin mutations do not affect MT density or TBCB levels. Moreover, the prolonged depletion of the MT network did not induce GAN-like aggregates of vimentin in normal fibroblasts. These results indicated that the generalized disorganization of IFs in GAN patients may not involve TBCB-mediated MT disassembly and must be regulated by a yet unidentified mechanism [109].

20.2.4

Neuronal Intermediate Filament Inclusion Disease

Neuronal intermediate filament inclusion disease (NIFID) is a recently described uncommon neurological disorder of early onset with a heterogeneous clinical

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phenotype, including sporadic frontotemporal dementia associated with a pyramidal and/or extrapyramidal movement disorder. The symptoms comprise behavioral and personality changes, which can be associated to memory loss, cognitive impairment, language deficits, and motor weakness. The cytopathological characteristics consist of neuronal loss, gliosis, swollen neurons, and presence of large inclusions in the cell body of many neurons that are immunoreactive for all of the class IV neuronal IFs and especially in α-internexin [110, 111]. These inclusions of α-internexin but negative for tau or synuclein distinguish NIFID from most cases of frontotemporal lobar degeneration. This raises the question whether α-internexin-positive neuronal inclusions in NIFID reflect any selective neuronal dysfunction, and as such if they are associated with some specific clinical symptoms. Genetic screening revealed no pathogenic variants for all type IV neuronal IFs, SOD1, NUDEL, and gigaxonin [112, 113]. Interestingly, the number of IF aggregates is high in areas with reduced neuronal loss and low in sites of intense neuronal degeneration. Cairns et al. [110] proposed that the formation of these inclusions is an early event in the pathogenesis of NIFID, and these aggregates are then released and degraded into the extracellular space following degeneration of the neuron. The mechanism of IF aggregation and the role they play in neuronal dysfunction and cell death are still unclear. Although immunoreactivity for IFs was initially described as the defining pathological feature of NIFID, not all the inclusions in NIFID are IF-positive. It now appears that aggregates of FUS (fused in sarcoma) protein, is a more consistent feature of NIFID. Indeed, intracellular accumulations of FUS are more often encountered than IF inclusions and all neurons that contained abnormal IF aggregates also contained FUS inclusions [114]. The authors interpreted this finding as suggesting that FUS plays a more central role in the pathogenesis of NIFID and that the abnormal accumulation of IFs is likely a secondary phenomenon.

20.2.5

Diabetic Neuropathy

Diabetes is the leading cause of peripheral neuropathy worldwide. Diabetic neuropathies are complex, heterogeneous disorders that affect dorsal root ganglia and sensory axons more so than motor fibers. Although its pathogenesis has not been fully elucidated, diabetic neuropathy is characterized by slower conduction velocity, impairment of axonal transport, axonal atrophy, and reduced capacity for nerve regeneration. All these features of nerve function depend on the integrity of the axonal cytoskeleton and particularly on NFs. In agreement with this, multiple abnormalities of NF biology were identified in models of diabetes. An impairment of the axonal transport of NFs, actin, and tubulin concomitant with a proximal increase and a distal decrease of axonal calibers were observed in rats with streptozotocin-induced diabetes and in BioBreeding rats (a model of spontaneous type I diabetes, [115, 116]). The distal axonal atrophy is accompanied by a concomitant NF loss in this region [117], and accumulations of highly phosphorylated NF epitopes are present in proximal axonal segments of dorsal root ganglia sensory neurons from diabetic patients [118]. An increase of NF phosphorylation, correlated with activation of JNK, was also

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detected in lumbar dorsal root ganglia from rat models [119]. Finally, there were substantial declines in the mRNA levels of all three NF subunits as well as reduced NF numbers and densities within large myelinated sensory of long-term diabetic models [120]. All these results suggest that NF abnormalities may contribute to the development of diabetic neuropathy, or may be affected by this disease. However, slowing of conduction velocity in diabetic models occurs much earlier than loss of NF investment or axonal atrophy [120]. To further elucidate the contribution of NFs to diabetic neuropathy pathogenesis, the effect of streptozotocin-induced diabetes was analyzed in NFH-LacZ transgenic mice characterized by axons completely lacking NFs [121]. Interestingly, diabetic mice lacking NFs developed progressive slowing of conduction velocity in their motor and sensory fibres and displayed decreased nerve action potential amplitudes earlier than diabetic mice with normal IF cytoskeleton. Moreover, superimposing diabetes on axons without NFs also accentuated axonal atrophy. Administration of insulin that restored normoglycemia reversed conduction slowing and restored sensory axon caliber. These findings indicate that changes in NF expression, transport, or posttranslational modifications cannot account alone for neurological features of diabetic neuropathy, but these IFs may help axons to better resist the negative effects of diabetes [121].

20.2.6

Parkinson’s Disease

A neuropathological hallmark of Parkinson’s Disease (PD) is the formation of filamentous neuronal inclusions named Lewy bodies, composed of α-synuclein, NF proteins, ubiquitin, and proteasome subunits. Various features distinguish NFs in PD, including inappropriate phosphorylation and proteolysis in Lewy bodies [122, 123], and decreased NFL and NFH mRNA levels [124]. A point mutation in the NEFM gene was reported in a case of PD with early onset [125]. This mutation consisted in a substitution of Ser for Gly at residue 336, a highly conserved region in the rod domain 2B of NFM. Although three other unaffected family members also carried this mutation, the authors had then proposed that aberrations in neuronal IFs could lead to the development of the pathology seen in PD. However, the G336S mutation does not disrupt the assembly and the distribution of NFs in vitro [126] and the screenings of PD patients of similar or different ethnic background failed to identify this mutations [127, 128], arguing against the implication of this NFM mutation in pathogenesis of PD. Nevertheless, the discovery of two other mutations located in highly conserved sites of NFM in PD patients suggests that NF gene mutations could act as susceptibility factors.

20.3

Glial Intermediate Filament GFAP and Alexander Disease

Neuronal IFs are not the only class of IF to be responsible for the development of neurological disorders. Glial IF can also be the primary cause of a CNS disorder. Indeed, GFAP, the major constituent of astrocytic IFs, is directly involved in the

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development of the Alexander disease (AXD). This disease is a fatal, progressive white matter disorder that has been classified into three types based on the age of onset: infantile, juvenile, and adult. The infantile type, with onset between birth and about 2 years of age, is the most frequent form of the disease and is fatal either within that period or by around the age of 10 years. Clinical symptoms comprise progressive megalencephaly, seizures, and impaired cognitive function, which may be associated with ataxia and hydrocephalus. Such phenotypes become progressively less common for the juvenile and adult forms (for recent reviews, see [129, 130]). Both the infantile and juvenile forms usually appear to be sporadic while the adult form is often familial. AXD is a primary astrocytic disease and its manifestations are the result of astrocyte dysfunctions leading to both myelin damage and neuron dysfunction. Neuronal loss is often reported but axons are relatively well preserved in demyelinated areas. The pathological hallmark of AXD is the presence of protein aggregates known as Rosenthal fibers within the cytoplasm of astrocytes throughout the CNS, but especially those located in the subpial, periventricular, and subependymal zones. Different constituents were identified in Rosenthal fibers: GFAP, αB-crystallin, HSP27, and ubiquitin [131, 132, 133]. Although GFAP is also expressed in glial cells of the PNS and in several other organs, Rosenthal fibers were not reported outside the CNS of AXD patients. To examine the function of GFAP in vivo, GFAP knock-out mice were generated [134, 135, 136, 137]. These studies showed that mice lacking GFAP displayed astrocytes devoid of the IF, but still developed and reproduced normally. Only subtle phenotypes emerged with age, arguing for a role of GFAP in the white matter architecture, blood-brain barrier integrity, astrocyte-neuronal interactions, and in modulating synaptic efficacy in the CNS [136, 137]. This is consistent with the known roles of astrocytes that help to form blood-brain barrier, promote synaptic plasticity, and coordinate neuronal activity. To determine the influence of increased GFAP expression on astrocyte function, mice overexpressing the human GFAP gene were produced [138]. Mice in the highest expressing lines developed a phenotype close to that observed in AXD. Indeed, their brains contain many inclusion bodies indistinguishable from human Rosenthal fibers, astrocytes are hypertrophic and these animals died from an encephalopathy at an age that is inversely correlated with the level of expression of the transgene. However, no myelin abnormalities were observed. Microarray analysis performed on olfactory bulbs of these animals recently highlighted the appearance of an initial stress response by astrocytes which results in the activation of microglia and compromised neuronal function [139]. All these results suggested that a primary alteration in GFAP may be responsible for AXD. Sequence analysis of DNA samples from AXD patients was thus performed and revealed that most cases are associated with mutations in the GFAP gene [140]. Since then, numerous mutations of this gene were identified; many of them being located in highly conserved domains of the encoded protein that play specific roles in the assembly of IF network [8, 130]. It was estimated that more than 95% of AXD cases are due to GFAP mutation. To date, all the identified mutations are heterozygous and none leads to an absence of GFAP or truncated protein. Nearly all

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of them involve amino acid substitutions, but several insertion or deletion/insertion alterations have also been reported (a continually updated list of all published mutations is maintained at the Waisman Center of the University of Wisconsin-Madison; www.waisman.wisc.edu/alexander). Numerous mutations cluster in the coils 1A and 2B of GFAP and two sites (R79 and R239) account for approximately half of all patients affected. The comparison of mutations occurring in the various IF proteins revealed that frequent mutations lying in the 2A segment seem to be unique to GFAP. It is possible that molecular partners specifically interact with this region of GFAP but not with the equivalent region of other IFs. The calcium-binding protein S100B binds to the N-terminal part of GFAP-coil 2A [141]. As S100B prevents GFAP assembly [142], mutations in this domain could impair GFAP-S100B interactions, resulting in the accumulation of GFAP polymers and possibly aggregates. It seems that a correlation exists between the different mutations and the severity of the disease. However, there also exists significant phenotypic variability and age of onset for the same mutation [143], suggesting that epigenetic and environmental factors influence the appearance and timing of disease symptoms. It should also be noted that in rare cases of AXD, no mutations in the GFAP gene have been found [144], indicating that there may be additional causes of the disease. The discovery of GFAP mutations led to the generation of knock-in mice with missense mutations homologous to those found in humans (R76H and R236H, which correspond to the R79H and R239H mutations in human) [145, 146]. If the presence of mutant GFAP per se seemed insufficient for aggregate formation, a 30% increase in GFAP content over that in wild type induced the formation of Rosenthal fibers in multiple sites throughout the CNS [146]. These animals were also more susceptible to kainate-induced seizures. Nevertheless, they had a normal lifespan, showed no overt behavioral defects and general white matter architecture and myelination appeared normal. These features resemble those found in the adult form of AXD rather than in the infantile form. This indicates that the presence of GFAP aggregates-containing mutant GFAP is not sufficient to induce a major phenotype of AXD, even though it causes some abnormalities in the mouse. Interestingly, further elevation of GFAP via crosses to GFAP transgenic animals led to a shift in GFAP solubility, an increased stress response, and ultimately death [145]. This correlates GFAP protein levels to the severity of the disease. While the genetic basis for AXD is now firmly recognized, there is little information concerning the mechanisms by which GFAP mutations lead to disease. Several studies showed that mutations of GFAP alters the normal solubility and organization of GFAP networks [143, 147]. When expressed alone, these mutant proteins lost their ability to form filaments in vitro. However, in the presence of assembly partners, such as wild-type GFAP or vimentin, they were still capable of incorporation into filament networks in transfected cells. If wild-type GFAP is prone to aggregate, mutations of GFAP exacerbates this accumulation [148]. Insufficient amounts of plectin, due to R239C GFAP expression, were also proposed to promote GFAP aggregation and Rosenthal fibers formation in AXD [149]. Both inhibited proteasome activity and activated stress pathways seemed to be important consequences of GFAP accumulation [148]. As a positive feedback response, both the proteasome hypofunction

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and JNK activation exacerbated GFAP accumulation, increasing susceptibility of the cell to stressful stimuli. It thus appeared that accumulations of GFAP protein would be more deleterious to the astrocytes than the mutant protein itself. However, as a positive consequence, upregulation of αB-crystallin and HSP27 were also associated with the aggregation of GFAP in AXD patients [133, 150] as well as in cell and animal models [139, 145, 148]. Increased αB-crystallin levels would contribute to the disaggregation of GFAP aggregates and could protect cells from apoptotic events [151]. Moreover, a recent study demonstrated that AXD mutant GFAP accumulation stimulates autophagy which in turn contributes to decrease GFAP levels [152]. The balance between the positive and negative effects of GFAP accumulation might define the survival or death of the cell. Compounds known to reduce GFAP expression in vitro, such as quercetin, might be useful as therapeutics. For instance, treatment with the antibiotic ceftriaxone alleviates intracytoplasmic aggregates of mutant GFAP by inducing the upregulation of HSP27 and αB-crystallin, polyubiquitination and autophagy, and by reducing the GFAP promoter transcriptional regulation [153]. The GFAP gene is known to generate different splice variants, including the most abundant isoform GFAP-α, and seven other differentially expressed transcripts including GFAP-δ (human homologous GFAP-ε). GFAP-δ is incapable of self-assembly into IF per se, but can incorporate into a filament network composed of GFAP-α if the proportion of GFAP-δ to GFAP-α remains < 10% [154]. However, elevating the proportion of GFAP-δ perturbs association of αB-crystallin with the IF fraction and induced IF bundling and aggregation in transiently transfected cells. Interestingly, GFAP-δ isoform is preferentially expressed in the same populations of astrocytes that contain the most Rosenthal fibers in AXD. This raises the possibility that GFAP-δ may play a key role in aggregate formation in combination with mutated GFAP. It remains to determine whether GFAP-α:GFAP-δ ratio is perturbed in AXD tissues.

20.4

Conclusion

Intermediate filament abnormalities are reminiscent of many human neurodegenerative disorders. Despite extensive efforts over the past 40 years, processes leading to these abnormalities as well as their precise contribution to disease pathogenesis often remain poorly understood. For instance, if it is clearly established that mutation in IF genes can be a primary cause of neurodegenerative disorders, the question as to how they induce neurodegeneration frequently remains unsolved. Although transgenic mouse models have been somewhat helpful in understanding some mechanisms, most of these animals displayed a much less severe phenotype than patients. A growing body of evidence suggests that perturbation of IF axonal transport and/or stoichiometry are directly involved in the formation of intracellular IF aggregates. Destabilization of IFs’ mRNA could be responsible for alterations in IFs’

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stoichiometry whereas aberrant posttranslational modifications could affect their transport. The identification of compounds able to alleviate these IFs aggregates is crucial for the development of new therapeutic approaches. More investigations are also necessary to identify IF partners. Indeed, protein kinase Cε (PKCε) interacts with peripherin in vitro and causes its aggregation, leading to an increase of apoptosis [155]. The importance of IF associated proteins in the development of neurodegenerative disorders was also highlighted by the identification of mutations in genes encoding IF partners that mimic intermediate filaments-related disease. This is particularly the case of gigaxonin in GAN. Finally, it will be important to elucidate why certain types of intermediate accumulations appear more toxic than others. While perikaryal accumulations are generally well tolerated, axonal inclusions are often noxious. The more deleterious effect of axonal aggregates on axonal transport could be a promising avenue to explore in the future.

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Index

A Acetylation, 37, 38, 242, 247, 248, 252 ACTA1, 125–127 ACTG1, 172, 174, 177 Actin, 4, 6, 10, 78, 103, 104, 110, 171, 182, 183, 186, 290 β−Actin, 18, 171, 172, 176, 177, 189 Actin-binding proteins, 7, 150, 169, 175, 176, 187, 190 Actin cytoskeleton, 84, 100, 114, 165, 176, 184, 192, 194 Actin isoforms, functions, 4, 8, 11, 12, 15, 20 Actin mutation, 12, 128 Actin polymerization, 6, 7, 19, 164, 171 Acto-myosin complex, 4 Adenomatous Polyposis Coli, 45 Adhesion, 17, 83, 109, 148, 182, 184, 186, 190, 193, 334, 3363 ADNSHL, 172 Adriamycin, see Doxorubicin Agents (VDA), see Vascular disrupting agents Anoikis, 189 Autosomal dominant nonsyndromic hearing loss, see ADNSHL Aging, 377, 386, 387, 395–398, 428 Agyria–pachygyria spectrum, 321, 322 Alexander disease, 426, 436, 437 Alzheimer’s disease, 41, 63, 66, 285, 361 Alzheimer’s syndrome, 62 Amyotrophic lateral sclerosis (ALS), 63, 426 Annealing, 6 Annexin 6, 114 Anticancer drugs, 63–65, 204, 214 Apoptosis, 64, 186–189, 207, 209–211, 230, 265, 275, 393, 395, 418–420, 440 Arginylation, 4, 11, 12, 40, 251 Atelosteogenesis, 151 Atypical Werner syndrome, 378, 381, 385, 386

ATPase, 41, 42, 60, 62, 75, 78, 79, 130, 390 Autoimmune disorders, 113 Axonal transport, 41, 57, 60–62, 361, 362, 427–430, 435 Axonal fibers, 332, 333 B Breast carcinoma, 37, 165, 271 Buschke–Ollendorff syndrome (BOS), 390 C Cabazitaxel, 210 Calponin homology (CH), 44, 141, 150, 350, 360 Cancer, 34, 37, 66, 87, 154, 182, 186, 188, 204, 214, 225, 228, 236, 268, 273, 276 Cancer metastasis, 162, 164–166 Cancer therapy, biomarkers, 64 Cap disease, 124, 127 Carboplatin, 229, 230, 234 Cardio binding domains, 81 Cardiomyopathy, 78, 128–130, 153, 378, 384, 397, 399 Cardiac muscle, 9, 12, 13, 83, 124, 131, 133, 143, 182, 378, 388, 411 CCNU, 274 Ceftriaxone, 439 Cell invasion, 164–166, 273 Cell–cell interactions, 109 Cell–ECM interactions, 16 Central nervous system (CNS), 146, 207, 234, 386 Cerebellar hypoplasia, moderate-to-severe, 322 Cerebral spinal fluid (CSF), 299 Cervical kyphosis, see Larsen syndrome Charcot-Marie-Tooth disease, 378, 426, 430 Chromatin associated proteins, 392 Ciliar dysfunction, 63 Cisplatin, 230, 274

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449

450 Clostridium difficile toxin A, 184 Clubfoot, 127 CM mutations, mouse, 129 Cofilin, 7, 124, 160, 164, 166, 187, 190, 191, 193 Cofilin activity cycle, 166, 191 Colchicine, 207, 214, 215, 263 Colon cancer, 64 Combretastatins, 214, 215 Congenital fiber type disproportion (CFTD), 124, 126 Congenital muscle diseases, 124 Contraction, 18, 20, 75, 81, 83, 123, 128, 190, 193, 380 Cryoelectron microscopy, 76 C-terminal tail (CTT), 31, 39 Cuticular plate, 169, 171, 177 Cyclostreptin, 211 Cytochalasin, 182 Cytoskeleton, 8, 40, 103, 165, 178, 186, 326, 393, 411, 425 Cytotoxicity, 182, 207, 211, 213, 215 D DCM mutations, 130 Deafness, 84, 151, 172, 174, 363 Deiters’ cells, 171 Desmin, 153, 411, 412, 414, 416–418, 420 DFNA20/A26, 172, 173, 175–177 Dictyostatin, 211 Dictyostelium, 9 Dilated cardiomyopathy (DCM), see Cardiomyopathy Discodermia dissoluta, 211 Disease phenotypes, 130, 377, 378, 391 Distal arthrogryposis, 124, 127 Distribution, 10–13, 15, 30, 35, 44, 83, 150, 171, 294, 348, 352, 420, 436 DNA-alkylating agent, see CCNU DNA break repair, 395, 401 Docetaxel, 65, 208–210, 215, 231, 234 Dolabella auricularia, 22 Doxorubicin, 164, 274 Drug-refractory cancers, 226 Drug resistance, 7, 210, 213, 215, 224, 228–231, 247, 252, 273 Drug sensitivity, 209, 229, 231, 233, 252 Drug–tubulin interactions, 276 Dynamic, 4, 17, 40, 64, 109, 163, 225 E EB homology (EBH), 44, 45 Ehlers-Danlos syndrome, 327

Index Emery-Dreifuss muscular dystrophy (EDMD), 377, 379, 389 End-to-end annealing, 6 Epidermal growth factor (EGF), 85, 165, 183 Epothilones, 210, 211, 237 Eribulin, 207, 237 ERM family, 106, 171 ERM proteins, 107, 109 Espin, 171 F Familial cardiomyophies, 128–130, 410 Farnesyl transferase, 386–388, 395, 397, 400, 401 Fibroblast growth factor (FGF), 183 Filamin, 7, 11, 141, 144, 148 Filamins role, in cancer, 154 Fimbrin, 176 FLNA germline mutations XL-PH, neurological clinical manifestations, 146 depression, 146 dyslexia, 146 epilepsy, 146 psychosis, 146 FLNA germline mutations (male) cardiovascular defects, 147 congenital lobar emphysema, 147 cortical dysgenesis, 147 small intestine, shortening of, 147 tracheobronchomalacia, 147 Fluorescence resonance energy transfer (FRET), 272 5-Fluorouracil, 274 Function, 18, 19, 63, 123, 128, 131, 362, 418 G Gain-of-function (FLNA), see FLNA germline mutations Glioblastoma, 37, 234, 235 Gamma-actin, 11, 171, 174–177 Gamma-actinin, 176 Gelsolin, 7, 176, 188, 189 Gene transcription, 392–394 Genetic disorders, 345 Giant axonal neuropathy, 426, 433 Gigaxonin, 433–435, 440 Glial fibrillary acidic protein, 414, 426 H HCM loop, 78, see also Cardiomyopathy HCM mutations, 130 Hearing loss, 149, 171–173, 175–177 Heart failure, 130, 418, 419

Index Heart-hand syndrome (HHS), 384 Heterochromatin, 379, 387, 389, 390, 393, 401 Hemolytic anemia, 346, 351–353, 355–357, 364 Hereditary elliptocytosis, 346, 352, 363 Hereditary spherocytosis, 356 Histones, 245, 248, 414 HRD syndrome, 329, 330 Human actin genes, 8 Hutchinson–Gilford progeria syndrome (HGPS), 378, 386 Hypertrophic cardiomyopathy (HCM), see Cardiomyopathy I IHCs, inner hair cells, see Inner hair cell (IHC) Immunodeficiency, 104, 111, 113 Inner hair cell (IHC), 169, 176 Integral membrane proteins of the INM, 346, 389, 393 Intermediate filament, 41, 84, 375, 376, 386, 411, 412, 414, 419, 425 Isoforms, 18, 81, 86, 131, 160, 193, 265, 297, 348, 351, 427 Ixabepilone, 210, 211, 247 J Jasplakinolide, 182, 183, 187 K Kaplan-Meier curves, 235 Katanin, 40, 42, 247 Kinesin spindle protein (KSP), 289 Knock-out mice, 17, 19, 275, 437 L Lamins, 365, 376–378, 389–394, 396–398 Larsen syndrome, 151–153 Latrunculin, 182, 187, 189 Laulimalide, 211, 213 LIM kinase 1 (LIMK1), 159–166, 191 LIM kinase 2 (LIMK2), 159–164 Lipid rafts, 109, 110, 113–115 Lipodystrophy, 378, 385–388, 396 Lissencephaly, 316, 321–323, 327 LMNA gene mutations, 378, 387, 391 M Magnetic resonance imaging (MRI), 146, 323, 327, 332 Major histocompability complex (MHC), 103, 104, 115 Major histocompability complex (MHC) class II, 103

451 Mandibuloacral dysplasia type A (MADA), 378, 387, 390, 395, 399, 401 type B (MADB), 387 MAPK, 106, 183, 185, 186, 264, 265, 288 Mechanotransduction, 63, 145, 169, 412, 415 Mental retardation, moderate-to-severe, 316, 328 Mesenchymal stem cells, 397, 398 Metastatic melanoma median survival time, 234 2-Methoxyestradiol, 215 Metastasis, 7, 64, 115, 154, 164, 165, 189, 191–194, 248, 272, 273 Microclusters, 106, 109, 110 Microtubule destabilizer, 206, 207 Microtubule dynamics, 30, 43, 64, 204, 206, 209, 230, 247, 334 Microtubule nucleation, 30, 36, 37, 41, 45 Microtubule regulatory proteins, 30, 40, 41 Microtubule stabilizer, 208, 211, 213, 215 Microtubule structure, 29–31, 206 Microtubule-associated proteins (MAPs), 31, 224, 234, 320 Microtubule-organizing centers (MTOC), 30 Microtubules, 30, 37–40, 43, 59, 63, 65, 205, 207, 330 Milataxel, 210 Miro-Milton complex, 62 Membrane skeleton, 346, 349 Microtubule destabilize, 206, 207 Miller–Dieker syndrome, 328 Motility, 4, 57, 59, 78, 164, 189, 203, 268 MRI scans, brain, 146 Muscular dystrophy, 132, 378, 380, 386 Muscularis propria, 148 Myosin, 6, 9, 15, 74–85, 123, 176, 194, 414 N NEB, 125, 127 Nemaline myopathy, 124, 125, 129 Néstor-Guillermo progeria syndrome (NPGS), 390 Neurodegenerative diseases, 41, 247, 295, 361, 426 Neurofibrillary tangles (NFTs), 288, 296, 297 Neurofilament, 289, 425, 429 Neuronal migration, 146, 147, 316, 317, 319, 323, 327, 331–334 Neuronal polarity, 287, 288, 291, 292, 297, 298, 319, 333 Neuropathy, 206, 207, 210, 388, 433, 435 NM mutations, 127, 129

452 Nocodazole, 164, 247, 248, 261, 233 Nomenclature, 57, 59, 73, 318 Non-small cell lung cancer (NSCLC) cells, 85, 235 Nuclear dysmorphisms, 381 Nuclear envelope, 375, 377, 388–390, 392–394 Nuclear fragility, 392 Nuclear lamina, 375–377, 379, 387, 392 Nuclear membranes, 375, 393 O OHCs, 169 Otopalatodigital syndrome spectrum disorders, 149, 150, 153 Outer hair cells, see OHCs OPDSDs frontometaphyseal dysplasia (FMD), 149 Melnick-Needles syndrome (MNS), 149 otopalatodigital syndrome types 1 and 2(OPD1, OPD2), 149 pathogenesis, 149, 150 terminal osseous dysplasia (TOD), 149 Ovarian cancer, 87, 230, 231, 236, 247, 250, 273 progression-free survival, 229, 231 P Paclitaxel, 57, 63, 64, 208–211, 213–215, 227, 229–235, 273–275 Palmitoylation, 39, 242, 248, 249 Pancreatic cancer TBAs, 223 Pathogenic mechanisms, 144, 148 Parkinson’s disease, 249, 250, 291, 299, 426, 436 Peloruside A, 211, 213 Peripherin, 414, 425, 427, 429, 434 Pick’s disease, 293, 294 Pharmacological treatments, 85, 165, 166, 185, 193, 399 Phosphorylation, 39, 161, 250, 263, 289, 416 Point mutation, 62, 230, 293, 363, 436 Polycystic kidney disease, 63 Polyglutamylation, 37, 39, 40, 242, 245–247 Polyglycylation, 37, 39, 242, 245, 246 Polymerization, 4–7, 32, 43, 64, 164, 176, 225, 241, 249, 332, 414, 416 Post-translational modifications (PTMs) acetylaction, 37 detyrosination, 37 polyglutamylation, see Polyglutamylation polyglycylation, see Polyglycylation

Index Polysorbate-80, 209 Periventricular nodular heterotopia, 146, 149, 327 Prelamin A, 385–388, 390, 391, 394–396, 400 Premature aging, 377, 379, 386, 397, 398 Profilin, 6, 9, 35, 176, 185, 268 Progeroid syndromes, 385 Proliferation, 15, 106, 109, 165, 183, 209, 271, 274, 320, 331, 364 Protein aggregates, 126, 153, 428, 437 Protein interaction, 113, 266, 350, 351, 356 Protein–protein interactions, 109, 231, 265, 290, 346 Protein trafficking, 235, 361 R Radixin, 171 Red cell membrane, 347, 352 Refractory epilepsy, 316, 328 Restrictive cardiomyopathy (RCM), see Cardiomyopathy Restrictive dermopathy (RD), 378, 388 Rho, 19, 84, 110, 143, 147, 161, 182, 185, 429 RT-PCR, 35, 164, 229 S Sanjad-Sakati syndrome, 329 Saccharomyces cerevisiae, 249 Scarf-Bloom-Richardson (SBR) grade, 245 Schwann cells, defects in, 62 Scoliosis, see Larsen syndrome Septin, 114, 115 Serum response factor (SRF), 10 Signaling pathways, 11, 154, 165, 209, 216, 317, 320, 395, 396, 398 Single alpha helix (SAH), 81, 82 Spondylocarpotarsal syndrome (SCTS), see FLNA mutations Skeletal muscle, 13, 82, 123, 124, 128, 131, 414 Sorangium cellulosum, 210 Spectrin–actin interaction, 347, 353 Spinocerebellar ataxia, 359 Stress fibers, 9, 16, 18, 20, 162 Stathmin, 7, 42, 43, 259, 267 Stereocilia, 18, 81, 83–85, 169, 171, 176, 177 Strain-stiffening, 8 Structure, 30, 31, 43, 73, 77, 79, 142, 144, 206, 350, 351 T Taccalonolides, 213, 214 Tail–motor interactions, 82 Tau–actin interaction, 290, 292

Index Tau–tau interactions, 290 Tauopathy mutations, 290, 293, 295, 296, 299 Taxane, 208–211, 213, 224, 230, 247, 273 Taxus brevifolia, 208 T cell activation,104, 106, 109, 110, 113, 115, 116 T cell migration, 110, 111 T cell receptor (TCR), 104–108, 110, 111, 113–115 +TIPs, 40, 43–45 TNAP, 289, 290 Torsion dystonia, 390 Toxoplamsa gondii, 251 TPM3, 125–132 Traumatic brain injury, 298 TRIOBP, 171 Tropomyosin, 9, 15, 124, 127, 176, 185, 186, 189, 194, 347 Tubulin, 31, 33, 39, 241, 262, 321 γ−Tubulin, 36, 37, 234, 249, 250 βIII-Tubulin, 34, 36, 228–237, 250, 252 Tubulin acetylation, see PTMs αβ-Tubulin dimers, 31 Tubulin binding agents, 216, 224, 226–228, 230, 232 Tubulin code, 40, 245, 247 Tubulin detyrosination, 38, 40 Tubulin isotypes, 30, 33, 35, 226, 242, 334 Tubulin phosphorylation, see PTMs Tubulin post-translational modifications, 30, 37 Tubulin tyrosine ligase (TTL), 38, 244, see also Tubulin detyrosination Tubulin tyrosine ligase-like proteins (TTLL), 39, 245, 246 Tumour, 115, 181, 182, 186, 187, 189, 191, 192, 266, 268, 270–272, 274–276

453 Twinfilin 2, 172, 177 Tyrosination, 38, 242, 244 V Vascular disrupting agents (VDA), 214, 215 Vasodilator-stimulated phosphoprotein (VASP), 84, 191 VAV1, 106, 108, 110, 111, 113 Vinblastine, 57, 63, 205–207, 247–249, 273–275 Vinca alkaloids, 204–209, 214, 223, 226, 227, 230, 241, 247, 273, 275 Vincristine, 164, 205, 206, 215, 227, 230, 273 Vinflunine, 205–207 Voltage-dependent anion channel (VDAC), 188 W WASP, 111, 113, 191 Whirlin, 84, 172, 176, 177 Wiskott-Aldrich syndrome, 104, 110, 111, 113 X Xenograft model, 271, 273 Xenopus FRET analysis, 272 X-Linked neutropenia, 104, 111, 113 XL-PH, vascular anomalies cardiac valvular anomalies, 146 focal glomeruloid malformations, 146 hemmorrhagic complications, rare in male, 146 megathrombocytopenia, 146 patentd ductus arteriosus, 146 Z Zebrafish, 63, 362, 364, 412