Neuromuscular Diseases

............................ Neuromuscular Diseases: From Basic Mechanisms to Clinical Management

..

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

Monographs in Clinical Neuroscience Vol. 18

Series Editor

M. Fisher, Worcester, Mass.

Advisory Board

W.G. Bradley, Miami, Fla. H.P. Hartung, Wu¨rzburg M.A. Moskowitz, Charlestown, Mass.

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

Neuromuscular Diseases: From Basic Mechanisms to Clinical Management

Volume Editor

F. Deymeer, Istanbul

16 figures and 21 tables, 2000

............................ Dr. Feza Deymeer Professor of Neurology Department of Neurology University of Istanbul C ¸ apa, Istanbul Turkey

Library of Congress Cataloging-in-Publication Data Neuromuscular diseases: From basic mechanisms to clinical management / volume editor, Feza Deymeer. p. ; cm. – (Monographs in clinical neuroscience, ISSN 1420-2441; vol. 18) Includes bibliographical references and indexes. ISBN 3805570562 (hardcover : alk. paper) 1. Neuromuscular diseases. I. Deymeer, Feza. II. Series. [DNLM: 1. Neuromuscular Diseases. WE 550 N4935 2000] RC925.5 .N473 2000 616.744–dc21 00-030965

Bibliographic Indices. This publication is listed in bibliographic services, including Current ContentsÔ and Index Medicus. Drug Dosage. The authors and the publisher have exerted every effort to ensure that drug selection and dosage set forth in this text are in accord with current recommendations and practice at the time of publication. However, in view of ongoing research, changes in government regulations, and the constant flow of information relating to drug therapy and drug reactions, the reader is urged to check the package insert for each drug for any change in indications and dosage and for added warnings and precautions. This is particularly important when the recommended agent is a new and/or infrequently employed drug. All rights reserved. No part of this publication may be translated into other languages, reproduced or utilized in any form or by any means electronic or mechanical, including photocopying, recording, microcopying, or by any information storage and retrieval system, without permission in writing from the publisher. Ó Copyright 2000 by S. Karger AG, P.O. Box, CH–4009 Basel (Switzerland) www.karger.com Printed in Switzerland on acid-free paper by Reinhardt Druck, Basel ISSN 1420–2441 ISBN 3–8055–7056–2

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

Contents

VII Preface 1 Novel Approaches to Therapeutics of the Muscular Dystrophies Hoffman, E.P. (Washington, D.C.); Buyse G.M. (Leuven) 12 Use of Normal and Genetically Modified Myoblasts for the Treatment

of Myopathies Skuk, D.; Tremblay J.P. (Ste-Foy, Que.)

26 Disorders of the Sarcoglycan Complex (Sarcoglycanopathies) Bo¨nnemann, C.G. (Go¨ttingen) 44 Facioscapulohumeral Muscular Dystrophy: Diagnostic and Molecular

Aspects Lunt, P.W. (Bristol)

61 Myotonic Dystrophy Moxley, R.T. (Rochester, N.Y.); Meola, G. (Milan) 79 Muscle Ion Channel Diseases Ru¨del, R.; Jurkat-Rott, K.; Lehmann-Horn, F. (Ulm) 96 Congenital Myasthenic Syndromes Engel, A.G.; Ohno, K.; Stans, A.A. (Rochester, Minn.) 113 Juvenile and Late-Onset Myasthenia gravis ¨ zdemir, C. (Istanbul) Deymeer, F.; Serdarog˘lu, P.; O 128 Hereditary Peripheral Neuropathies De Jonghe, P.; Timmerman, V.; Nelis, E. (Antwerp) 147 Acute Inflammatory Demyelinating Polyradiculoneuropathy Bella, I.R. (Worcester, Mass.)

163 Proximal Spinal Muscular Atrophy of Childhood Hausmanowa-Petrusewicz, I.; Zaremba, J. (Warsaw) 177 Familial Amyotrophic Lateral Sclerosis: A Review Morita, M.; Brown, R.H., Jr. (Charlestown, Mass.) 190 Author Index 191 Subject Index

Contents

VI

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

Preface

This volume of Monographs in Clinical Neuroscience is devoted to neuromuscular disorders, a field where much progress has been made in the last two decades, particularly in genetics. Mainly genetic, but also some immunological disorders, with a wide spectrum ranging from those affecting the muscle to those affecting the motor neuron, are presented in detail, including present and potential future therapeutic modalities. The first two chapters discuss different approaches which can lead the way to successful treatment of muscle diseases, particularly Duchenne muscular dystrophy. One approach is screening pharmacological agents in animal models with the same genetic defect as in human muscular dystrophy. Another important advancement is new gene therapy vectors based upon adenovirus and adeno-associated virus. Much research is aimed at improving a different potential therapeutic modality, myoblast transfer. Major areas of concern are suppressing the immune response to the myoblasts, promoting the migration of myoblasts into the target muscle and improving their survival in the muscle. Three major dystrophies are the subjects of the next three chapters: sarcoglycanopathies, facioscapulohumeral muscular dystrophy and myotonic dystrophy. Clinical features, protein expression pattern and mutational spectrum of a-, b-, c- and d-sarcoglycanopathies, a distinct group within recessive limbgirdle muscular dystrophies, are elucidated together with animal models and gene transfer. Facioscapulohumeral muscular dystrophy where the DNA mutation is known but the gene affected is unknown has a unique mutation. The intricacies of the mutational analysis and genotype-phenotype correlations are addressed and hypotheses for possible mechanisms of action of the muta-

VII

tion are suggested. Possible therapeutic options are also touched upon. In the chapter on myotonic dystrophy, molecular genetics of the disease is discussed including the phenomenon of anticipation and questions are raised about the reliability of predicting phenotype from genotype. Management issues and therapeutic trials are reviewed. The chapter on muscle ion channel diseases elaborates on the pathogenesis and the molecular genetics of sodium, calcium and chloride channelopathies. While the electrophysiological basis of some of the symptoms is explained, attention is drawn to phenomena which remain unexplained such as the temperature sensitivity of paramyotonia congenita patients as opposed to its absence in other sodium channelopathies. In the chapters on neuromuscular junction and peripheral nerves, both genetic and immunological diseases are included. A detailed discussion of the electrophysiological and molecular biological characteristics of congenital myasthenic syndromes, classified into presynaptic, synaptic and postsynaptic defects, is presented. Juvenile and late-onset myasthenia gravis stand out as distinct subgroups within myasthenia gravis. The recognition of epidemiological, clinical and immunological characteristics of the disease in these age groups helps in diagnosis and treatment. The chapter on hereditary peripheral nerve disorders discusses clinical and genetic characteristics of a wide range of disorders including CharcotMarie-Tooth 1 and 2, distal hereditary motor neuropathies, hereditary neuropathy with liability to pressure palsies and hereditary neuralgic amyotrophy. The next chapter deals with acute inflammatory demyelinating polyradiculoneuropathy and its axonal variants including AMAN and AMSAN. Clinical features, pathogenesis, significance of autoantibodies to glycoconjugates and approach to treatment are some of the topics analyzed. The chapter on proximal spinal muscular atrophy of childhood discusses its classification and its electrophysiological and pathological features. The role of the SMN1 gene and other candidate genes as well as that of the SMN protein and epigenetic factors including gender influence and incomplete penetrance of the mutated genes are considered. The last chapter concentrates on the pathophysiology of familial amyotrophic lateral sclerosis linked to the mutant SOD1 and on the neurotoxicity of the mutant SOD1 protein and gives a review of therapeutic trials. I hope that this monograph will contribute to efforts linking basic science to clinical medicine and that it will provide the reader with new insights and fresh perspectives in neuromuscular disorders. I thank all the authors who have made this monograph possible. Feza Deymeer

Preface

VIII

Deymeer F (ed): Neuromuscular Diseases: From Basic Mechanisms to Clinical Management. Monogr Clin Neurosci. Basel, Karger, 2000, vol 18, pp 1–11

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

Novel Approaches to Therapeutics of the Muscular Dystrophies Eric P. Hoffman a, Gunnar M. Buyse b a

b

Research Center for Genetic Medicine, Children’s National Medical Center, George Washington University, Washington, D.C., USA; Department of Pediatrics, Division of Child Neurology, University Hospital Gasthuisberg, KU, Leuven, Belgium

Recessively inherited muscular dystrophies are diseases caused by a lack of a specific protein component of muscle tissue. The identification of the dystrophin gene and protein initiated an impressive series of studies which have led to the characterization of nearly a dozen different muscular dystrophy genes over the last 12 years. The dramatically increased knowledge of the molecular basis of the muscular dystrophies has led to improved diagnosis of patients, accurate genetic counseling of their families, and an understanding of the pathophysiology of many of these disorders. However, the advances in molecular technology and understanding of the disease process have not yet led to improvements in patient treatment. Two recent approaches have shown particular promise in developing novel therapeutics. First, the identification of animal models with the same molecular genetic defect as human muscular dystrophy patients has allowed large-scale screening of pharmacological agents capable of increasing the strength of dystrophic muscle. Second, new developments in gene therapy vectors based upon adenovirus and adeno-associated virus have led to functional recovery of dystrophic muscle via delivery of the normal gene. This review focuses on these two approaches.

Animal Models as an Experimental Platform for Therapeutic Studies in Duchenne Muscular Dystrophy The identification of the dystrophin gene and protein as causative of Duchenne muscular dystrophy (DMD) made it possible to screen for dys-

trophic animals with the same primary biochemical defect [Koenig et al., 1987; Hoffman et al., 1987]. A number of mouse, dog and cat models were quickly identified which showed dystrophin deficiency of muscle and mutations of the dystrophin gene [Hoffman et al., 1987; Sicinski et al., 1988; Kornegay et al., 1988; Cooper et al., 1988; Gaschen et al., 1992; Winand et al., 1994; Carpenter et al., 1989]. Subsequently, the cardiomyopathic hamster was found to show primary delta-sarcoglycan deficiency [Nigro et al., 1997; Sakamoto et al., 1997], and additional ‘knockouts’ of sarcoglycan genes were accomplished [Hack et al., 1998; Duclos et al., 1998; Coral-Vazquez et al., 1999]. These ‘homologous’ animal models show clinical presentations and progressions which were discordant with their human counterpart. Dystrophindeficient cats (HFMD) show dramatic muscle hypertrophy, with lingual hypertrophy leading to an inability to lap water, and diaphragmatic hypertrophy leading to a constriction of the esophagus [Gaschen et al., 1992, 1998, 1999]. Dystrophin-deficient dogs (CXMD) show rapid muscle weakness and wasting, with some puppies dying as neonates, while most show substantial physical disability by just 6 months of age [Kornegay et al., 1988; Cooper et al., 1988]. Dystrophin-deficient mice (mdx) show asymptomatic hyperCKemia and muscle hypertrophy, with weakness only relatively late in their normal life span [Lefaucheur et al., 1995; Pastoret and Sebille, 1995]. Sarcoglycan-deficient hamsters present with cardiomyopathy, while the human counterpart shows no overt heart involvement [Nigro et al., 1997]. The clinical discordancy between the human disease and homologous animal counterparts led to the perception that these were not ‘good’ animal models. Indeed, a series of studies were conducted to ‘improve’ the mdx mouse model, by adding additional gene defects to the mouse to increase the clinical severity: ‘double mouse knockouts’ for dystrophin and myoD [Megeney et al., 1996], and dystrophin and utrophin [Grady et al., 1997] were done to hasten the disease presentation and progression. In some instances, these confirmed existing models of pathophysiology of the disease. For example, it is known that dystrophin deficiency leads to secondary loss of neuronal nitric oxide synthase (nNOS) localization at the myofiber membrane [Brenman et al., 1995]. This loss of nNOS localization leads to abnormal vascularization of muscle during exercise [Thomas et al., 1998]. A double knockout of nNOS and dystrophin showed no effect on the mdx phenotype [Chao et al., 1998]: this is to be expected, as genetic loss of nNOS (via knockout) is not likely to be different from biochemical loss of nNOS (via mislocalization). In other instances, the double knockouts clarified functional roles of proteins involved in the pathophysiology of dystrophin deficiency. For example, utrophin had been hypothesized to be capable of functionally replacing dystrophin [Tinsley et al., 1996, 1998], and the creation of double knockouts for dystrophin and

Hoffman/Buyse

2

utrophin confirmed this hypothesis [Grady et al., 1997; Rafael et al., 1998; Gilbert et al., 1999]. Finally, in yet other instances, double knockouts simply clouded the ‘pathophysiological picture’. For example, deficiency of both myoD and dystrophin showed a more severe phenotype, however the mechanism(s) underlying this exacerbation of symptoms is not at all clear [Megeney et al., 1999]. An alternative approach to ‘improving’ the animal models is to rest assured that they do indeed represent ‘good’ genetic and biochemical homologues of their human counterparts, and to utilize them to develop novel therapeutics. Gene and/or protein replacement through gene therapy is one clear and rational approach, which is discussed later in this review. A second approach is to develop experimental systems through which pharmacological agents can be screened for possible efficacy in human patients. This latter approach has been utilized in a recent series of reports using the dystrophin-deficient mdx model.

Drug Screening in Animal Models Drug screening of animal models of muscular dystrophy has been pursued since the early 1970s. In these ‘premolecular era’ studies, specific inherited muscular dystrophies in animals were argued to be a valid model for human disease based upon clinical findings. One such popular model, the dystrophic chicken, was used to screen various drugs which targeted specific pathophysiological processes (immune system, calcium traffic, muscle growth and homeostasis) [Barnard et al., 1976; Hudecki et al., 1981]. These studies led to the identification of the glucocorticosteroid prednisone as a compound able to slow the dystrophic process, a drug which has since been applied to large numbers of DMD patients with clear success [Fenichel et al., 1991; Griggs et al., 1991]. The rationale of these studies was that many of the pathophysiological processes (myofiber degeneration, regeneration, connective tissue proliferation, muscle wasting) were probably shared by the different muscular dystrophies, and that pharmacological agents able to target these processes could show efficacy independent of the specific (unknown) primary biochemical defect. The molecular identification of homologous animal models has greatly facilitated their use as an experimental model for their human counterpart. A number of investigators have shown that exercise exacerbates the mdx mouse phenotype [Hudecki et al., 1993]. This knowledge has been further developed into a drug screening protocol using exercised mdx mice, with a relatively crude yet effective whole-body strength assay for drug efficacy [Granchelli et al., 1999]. The initial screen of 18 drugs has shown promising agents which target specific aspects of the pathophysiology of dystrophin deficiency. Specifi-

Novel Approaches to Therapeutics of the Muscular Dystrophies

3

cally, considerable gains in muscle strength have been shown using glutamine, pentoxifylline, creatinine, IGF-I, oxatomide, and prednisone. Oxatomide is a mast cell stabilizer, which inhibits degranulation of mast cells, thereby protecting myofibers from damage via proteases and immune mediators [Gorospe et al., 1994a, b, 1996]. IGF-I is a muscle growth factor, and likely improves muscle regeneration [Barton-Davis et al., 1998]. Glutamine is a ‘conditionally essential’ amino acid, which is largely derived from skeletal muscle stores. Supplementation of this amino acid presumably replenishes depleted stores due to diseased muscle, and consequently may prevent muscle necrosis through decreased protein degradation and protection against oxidative stress. Indeed, preliminary data on DMD patients appears to confirm the beneficial effect of this compound on protein homeostasis in dystrophic muscle [Hankard et al., 1998]. Pentoxifylline is a hemorheologic agent which is used to enhance peripheral vascularization of tissues; in DMD it likely functions by counteracting the nNOS-deficiency-induced vascular insufficiency of dystrophin-deficient muscle [Thomas et al., 1998]. Alternatively, by inhibiting phosphodiesterase activity, it might be beneficial in DMD by decreasing the production of pro-inflammatory cytokines. Creatine is a well-documented ‘performance enhancer’, and likely functions through improvement of muscle energy metabolism, stimulation of protein synthesis, and/or decreasing intracellular calcium overloading in dystrophic muscle [Pulido et al., 1998]. Creatinine is a metabolite of creatine that presumably functions in an analogous manner. Additional recent studies have suggested that gentamycin, an aminoglycoside antibiotic which may permit the reading-through of genetic stop codons, may show benefit in mdx mice [Barton-Davis et al., 1999]. Human clinical trial networks are being established internationally to test these drugs, and other agents able to improve the strength of dystrophin-deficient mice.

Gene Delivery to Dystrophic Muscle All pharmacological approaches to treatment of muscular dystrophy hinge on altering the pathophysiology of the disease, and, with a few exceptions, do not address the primary biochemical defect. Gene delivery is an alternative approach which should halt disease onset and/or progression through biochemical replacement of the missing protein. Two viral gene delivery systems have achieved considerable success in gene therapy of the muscular dystrophies: third generation adenovirus (aka ‘gutted’ adenovirus), and adeno-associated virus (AAV). It should be pointed out that these two types of viruses are fundamentally different in their biology, despite the similar names. In a word, AAV can be viewed as a ‘parasite’ of more harmful viruses; it borrows ma-

Hoffman/Buyse

4

chinery for the production of its viral progeny from another virus which has coinfected the same cell (such as adenovirus and herpes-type viruses). AAV is not ‘replication-competent’; it is unable to produce viral progeny without the assistance of these other, larger viruses, and instead relies on its ‘helper’ virus for completion of its life cycle. As most of the helper viruses of AAV are themselves quite toxic to cells, AAV seems to be able to confer some beneficial effect to its host; reports have suggested that AAV can confer protection against human papilloma virus-induced cervical carcinoma [Hermonat, 1994] and inflammatory muscle disease [Tezak et al., 1999]. Thus, harnessing AAV as a gene delivery vehicle has intrinsic advantages, relative to other toxic viruses; these will be pointed out below. Gutted adenovirus is the only gene therapy vehicle that has been shown to be able to deliver full-length dystrophin protein, at levels sufficient to rescue significant amounts of dystrophin-deficient muscle [Kochanek et al., 1996; Clemens et al., 1996]. This viral vector is based upon adenovirus, but has been modified so as to remove all adenoviral genes. This ‘gutting’ of the viral genetic backbone has two important advantages relative to previous versions of adenoviral vectors. First, the removal of adenoviral genes releases considerable ‘space’ in the virus, so that constructs of up to 30 kb can be used. This ‘carrying capacity’ is much, much larger than any other viral delivery system used to date. Second, the removal of the adenoviral genes reduces the immunogenicity of the virus, at least with regard to the cellular immune response against infected cells. This is due to the lack of virally encoded antigens expressed by the gutted adenoviral backbone. Using this gutted adenovirus, a series of reports have shown that the fulllength dystrophin cDNA can be delivered, driven by a muscle-specific (creatine kinase) gene promoter [Kochanek et al., 1996; Clemens et al., 1996; Haecker et al., 1996]. Normal levels of dystrophin can be produced in an entire mdx mouse muscle, providing that the injections are done in neonatal mice. The recombinant viral genome persists as an episome in mature myofibers, and expression has been seen for as much as 1 year from the time of injection [Chen et al., 1997, 1999]. However, recent systematic studies have shown that the degree of persistence of dystrophin expression from the recombinant virus largely depends on the level of immune response against infected cells [Lochmuller et al., 1996; Howell et al., 1998]. There are remaining issues which must be solved before the gutted adenovirus can be considered a therapeutic approach with likely efficacy. First and foremost, mature myofibers lack viral attachment receptors for adenovirus, and thus significant infection can only be achieved in neonatal muscle, or in actively regenerating muscle [Feero et al., 1997b]. This ‘maturation-dependent’ infectivity of muscle may be overcome by altering the tropism of the virus;


5

recent experiments have been published where the protein coat of the virus was altered to target the virus towards heparin proteoglycan receptors [Bouri et al., 1999]. This increased infectivity of adult muscle, although not to the extent needed. Further research on altering the topism of the virus for adult muscle is needed. A second issue is that of sensitivity of the immune system for this virus. This problem was recently underscored by the report of a number of deaths due to acute immune response against recombinant adenovirus, in different human clinical trials. An additional concern is the need to separate recombinant virus from first generation adenovirus which is used as a ‘helper virus’ during production of infectious virions. A recent report has shown that this can be circumvented using cre/lox recombination sites in the helper virus, which are able to prevent the helper from packaging into infectious virions [Aoki et al., 1999]. Third, adenovirus may show lingering problems with ‘persistence’, due to its existence as an episome, which may be subject to loss from the myofiber nucleus. Finally, adenovirus remains an immunogenic virus, even with all viral genes eliminated. A recent publication suggests that infection of professional antigen-presenting cells (dendritic cells) in muscle may be a major reason for the antigenicity of adenovirus [Jooss et al., 1998]. AAV is a relatively recently characterized human parvovirus, which shows proclivity for mature muscle [Tezak et al., 1999]. The normal life cycle of the wild-type virus involves infection of nonmitotic cells, with subsequent integration into human chromosome 19 using a virally encoded recombinase protein (rep). It remains as a silent (latent) integrated virus, until the cell is infected with a helper virus (adenovirus or herpes-type viruses). It seems to confer some protection against the immune system; a recent study found that muscle may be protected from inflammatory attacks when infected with AAV. The virus is quite small in size, and appears able to pass through the myofiber basal lamina, unlike larger viruses (such as adenovirus and HSV). The ability of AAV to infect mature myofibers, to integrate stably into host chromosomes, and to evade immune system recognition makes AAV a seemingly ideal vector for muscle gene delivery. Its only apparent drawback is its limited packaging capacity: AAV can only contain transgene constructs of up to 5 kb, which is far too small for the 11-kb dystrophin coding sequence. Due to the constraints on transgene size, all published data to date has been centered on the sarcoglycan genes, which are much smaller in size and easily accommodated into recombinant AAV vectors. A spontaneously occurring hamster model of delta sarcoglycan deficiency has been characterized [Nigro et al., 1997; Sakamoto et al., 1997], and most reports have utilized this hamster to demonstrate the feasibility and efficacy of AAV-mediated gene therapy for muscle [Fisher et al., 1997]. To date, impressive data on biochemical, histological and functional rescue of the hamster tibialis anterior muscle has been

Hoffman/Buyse

6

published [Li et al., 1999], as well as some data on large-scale delivery to the hamster leg after permeabilization of the vasculature [Greelish et al., 1999]. These studies have shown persistence, high level expression and lack of immune response against the delta sarcoglycan protein, as was observed earlier with marker transgene studies [Xiao et al., 1996]. Human clinical studies have been approved by regulatory agencies, with injection of small foot muscles in alphasarcoglycan-deficient patients through a collaborative study of the University of Ohio in Columbus and the University of Pennsylvania in Philadelphia. Additional clinical trials on larger muscle groups are planned by our center with the University of Florida in Gainesville. Long-term toxicity trials are underway in primates, to ensure that there is no late adverse effect of sarcoglycan overexpression in muscle. It is important to note that only a few studies have been published regarding AAV transgene delivery in muscle. For example, while delta sarcoglycan overexpression in muscle does not appear to be toxic to myofibers, it is not unlikely that overexpression of alpha sarcoglycan or other proteins is in fact more deleterious. It will be critical to conduct a series of studies using many different transgenes to determine the general sensitivity of the myofiber to overexpression of proteins delivered by AAV. It is also important to determine whether recombinant AAV integrates into myofiber nuclear DNA, or whether it is able to persist long-term as an extrachromosomal episome. There are remaining hurdles before AAV vectors can be expected to achieve ‘therapeutic’ levels of gene expression in muscular dystrophy patients. First and foremost, widescale delivery must be accomplished, including gene delivery (AAV infection) of the heart and respiratory muscles. This will almost certainly entail systemic delivery through the vasculature. Such delivery routes raise problems of ‘targeting’ of the virus to only myogenic cells, permeabilization of the capillary bed, and production of the very large amount of virus needed to achieve genetic rescue of sufficient muscle. If the recombinant virus is injected into the vascular system, then it will likely infect endothelial cells, neurons, and other cell types. It is currently not known whether overexpression of a muscle protein such as delta sarcoglycan in nonmuscle cells will result in dysfunction of those cells. Two approaches to circumvent this (likely) problem include use of muscle-specific gene promoters, and development of targeting ligands which allow AAV to infect only myogenic cells [Feero et al., 1997a]. Successful permeabilization of the capillary bed has recently been reported [Greelish et al., 1999], however no studies of toxicity of such approaches in dystrophic muscle have been reported. With regard to large-scale production of recombinant AAV, the three-plasmid cotransfection method developed by Xiao et al. [1996] has proven highly successful in producing large amounts of recombinant virus from tissue culture cells, with no contamination of helper


7

virus. However, most studies to date have used virus preparations with a relatively low proportion of infectious particles, often with only 1 in 1,000 or less of virions able to successfully infect cells. Recently developed chromatography methods have been reported which seem able to increase the proportion of infectious virus particles in laboratory preparations [Clark et al., 1999], and this field promises to continue to quickly evolve.

References Aoki K, Barker C, Danthinne X, Imperiale MJ, Nabel GJ: Efficient generation of recombinant adenoviral vectors by Cre-lox recombination in vitro. Mol Med 1999;5:224–231. Barnard EA, Bhargava AK, Hudecki MS: Postponement of symptoms of hereditary muscular dystrophy in chickens by 5-hydroxytryptamine antagonists. Nature 1976;263:422–424. Barton-Davis ER, Cordier L, Shoturma DI, Leland SE, Sweeney HL: Aminoglycoside antibiotics restore dystrophin function to skeletal muscles of mdx mice. J Clin Invest 1999;104:375–381. Barton-Davis ER, Shoturma DI, Musaro A, Rosenthal N, Sweeney HL: Viral mediated expression of insulin-like growth factor I blocks the aging-related loss of skeletal muscle function. Proc Natl Acad Sci USA 1998;95:15603–15607. Bouri K, Feero WG, Myerburg MM, Wickham TJ, Kovesdi I, Hoffman EP, Clemens PR: Poly-lysine modification of adenoviral fiber protein enhances muscle cell transduction. Hum Gene Ther 1999; 10:1633–1640. Brenman JE, Chao DS, Xia H, Aldape K, Bredt DS: Nitric oxide synthase complexed with dystrophin and absent from skeletal muscle sarcolemma in Duchenne muscular dystrophy. Cell 1995;82:743–752. Carpenter JL, Hoffman EP, Romanul FCA, Kunkel LM, Rosales RK, Ma NSF, Dasbach JJ, Rae JF, Moore FM, McAfee MB, Pearce LK: Feline muscular dystrophy with dystrophin deficiency. Am J Pathol 1989;135:909–919. Chao DS, Silvagno F, Bredt DS: Muscular dystrophy in mdx mice despite lack of neuronal nitric oxide synthase. J Neurochem 1998;71:784–789. Chen HH, Mack LM, Choi SY, Ontell M, Kochanek S, Clemens PR: DNA from both high-capacity and first-generation adenoviral vectors remains intact in skeletal muscle. Hum Gene Ther 1999;10:365–373. Chen HH, Mack LM, Kelly R, Ontell M, Kochanek S, Clemens PR: Persistence in muscle of an adenoviral vector that lacks all viral genes. Proc Natl Acad Sci USA 1997;94:1645–1650. Clark KR, Liu X, McGrath JP, Johnson PR: Highly purified recombinant adeno-associated virus vectors are biologically active and free of detectable helper and wild-type viruses. Hum Gene Ther 1999; 10:1031–1039. Clemens PR, Kochanek S, Sunada Y, Chan S, Chen HH, Campbell KP, Caskey CT: In vivo muscle gene transfer of full-length dystrophin with an adenoviral vector that lacks all viral genes. Gene Ther 1996;3:965–972. Cooper BJ, Winand NJ, Stedman H, Valentine BA, Hoffman EP, Kunkel LM, Scott MO, Fischbeck KH, Kornegay JN, Avery RH, Williams JR, Schmickel RD, Sylvester JE: The homologue of the Duchenne locus is defective in X-linked muscular dystrophy of dogs. Nature 1988;334:154–156. Coral-Vazquez R, Cohn RD, Moore SA, Hill JA, Weiss RM, Davisson RL, Straub V, Barresi R, Bansal D, Hrstka RF, Williamson R, Campbell KP: Disruption of the sarcoglycan-sarcospan complex in vascular smooth muscle: A novel mechanism for cardiomyopathy and muscular dystrophy. Cell 1999;98:465–474. Duclos F, Straub V, Moore SA, Venzke DP, Hrstka RF, Crosbie RH, Durbeej M, Lebakken CS, Ettinger AJ, van der Meulen J, Holt KH, Lim LE, Sanes JR, Davidson BL, Faulkner JA, Williamson R, Campbell KP: Progressive muscular dystrophy in alpha-sarcoglycan-deficient mice. J Cell Biol 1998; 142:1461–1471. Feero WG, Li S, Rosenblatt JD, Sirianni N, Morgan JE, Partridge TA, Huang L, Hoffman EP: Selection and use of ligands for receptor-mediated gene delivery to myogenic cells. Gene Ther 1997a;4:664–674.

Hoffman/Buyse

8

Feero WG, Rosenblatt JD, Huard J, Watkins SC, Epperly M, Clemens PC, Kochanek S, Glorioso JC, Partridge TA, Hoffman EP: Single fibers as a model system for viral gene delivery to skeletal muscle: Insights on maturation-dependent loss of fiber infectivity for adenovirus and herpes simplex type I viral vectors. Human Gene Therapy 1997b;8:371–380. Fenichel GM, Florence JM, Pestronk A, Mendell JR, Moxley RT, Griggs RC, Brooke MH, Miller JP, Robison J, King W, et al: Long-term benefit from prednisone therapy in Duchenne muscular dystrophy. Neurology 1991;41:1874–1877. Fisher KJ, Jooss K, Alston J, Yang Y, Haecker SE, High K, Pathak R, Roger SE, Wilson JM: Recombinant adeno-associated virus for muscle directed gene therapy. Nat Med 1997;3:306–312. Gaschen F, Gaschen L, Seiler G, Welle M, Jaunin VB, Jmaa DG, Neiger-Aeschbacher G, Ade-Damilano M: Lethal peracute rhabdomyolysis associated with stress and general anesthesia in three dystrophindeficient cats. Vet Pathol 1998;35:117–123. Gaschen FP, Hoffman EP, Gorospe JRM, Cardinet GH, Ulh EW, Senior DF, Pearce LK: Dystrophin deficiency causes lethal muscle hypertrophy in cats. J Neurol Sci 1992;110:149–159. Gaschen L, Lang J, Lin S, Ade-Damilano M, Busato A, Lombard CW, Gaschen FP: Cardiomyopathy in dystrophin-deficient hypertrophic feline muscular dystrophy. J Vet Intern Med 1999;13:346–356. Gilbert R, Nalbantoglu J, Petrof BJ, Ebihara S, Guibinga GN, Tinsley JM, Kamen A, Massie B, Davies KE, Karpatı´ G: Adenovirus-mediated utrophin gene transfer mitigates the dystrophic phenotype of mdx mouse muscles. Hum Gene Ther 1999;10:1299–1310. Gorospe JRM, Nishikawa BK, Hoffman EP: Recruitment of mast cells to muscle after mild damage. J Neurol Sci 1996;135:10–17. Gorospe JRM, Tharp MD, Demitsu T, Hoffman EP: Dystrophin-deficient myofibers are vulnerable to mast cell granule-induced necrosis. Neuromuscul Disord 1994a;4:325–334. Gorospe JRM, Tharp MD, Hinckley J, Kornegay JN, Hoffman EP: A role for mast cells in the progression of Duchenne muscular dystrophy? Correlations in dystrophin-deficient humans, dogs, and mice. J Neurol Sci 1994b;122:44–56. Grady RM, Teng H, Nichol MC, Cunningham JC, Wilkinson RS, Sanes JR: Skeletal and cardiac myopathies in mice lacking utrophin and dystrophin: A model for Duchenne muscular dystrophy. Cell 1997;90:729–738. Granchelli JA, Pollina C, Hudecki M: Therapeutic evaluation of drugs using the mdx mouse. Neuromuscul Disord, in press. Greelish JP, Su LT, Lankford EB, Burkman JM, Chen H, Konig SK, Mercier IM, Desjardins PR, Mitchell MA, Zheng XG, Leferovich J, Gao GP, Balice-Gordon RJ, Wilson JM, Stedman HH: Stable restoration of the sarcoglycan complex in dystrophic muscle perfused with histamine and a recombinant adeno-associated viral vector. Nat Med 1999;5:439–443. Griggs RC, Moxley RT, Mendell JR, Fenichel GM, Brooke MH, Pestronk A, Miller JP: Prednisone in Duchenne dystrophy. A randomized, controlled trial defining the time course and dose response. Clinical Investigation of Duchenne Dystrophy Group. Arch Neurol 1991;48:383–388. Hack AA, Ly CT, Jiang F, Clendenin CJ, Sigrist KS, Wollmann RL, McNally EM: Gamma-sarcoglycan deficiency leads to muscle membrane defects and apoptosis independent of dystrophin. J Cell Biol 1998;142:1279–1287. Haecker SE, Stedman HH, Balice-Gordon RJ, Smith DB, Greelish JP, Mitchell MA, Wells A, Sweeney HL, Wilson JM: In vivo expression of full-length human dystrophin from adenoviral vectors deleted of all viral genes. Hum Gene Ther 1996;7:1907–1914. Hankard RG, Hammond D, Haymond MW, Darmann D: Oral glutamine slows down whole body protein breakdown in Duchenne muscular dystrophy. Pediatr Res 1998;43:222–226. Hermonat PL: Adeno-associated virus inhibits human papillomavirus type 16: A viral interaction implicated in cervical cancer. Cancer Res 1994;54:2278–2281. Hoffman EP, Brown RH, Kunkel LM: Dystrophin: The protein product of the Duchenne muscular dystrophy locus. Cell 1987;51:919–928. Howell JM, Lochmuller H, O’Hara A, Fletcher S, Kakulas BA, Massie B, Nalbantoglu J, Karpati G: High-level dystrophin expression after adenovirus-mediated dystrophin minigene transfer to skeletal muscle of dystrophic dogs: Prolongation of expression with immunosuppression. Hum Gene Ther 1998;9:629–634.


9

Huard J, Feero WG, Watkins SC, Hoffman EP, Rosenblatt D, Glorioso JC: The basal lamina is a physical barrier to viral mediated gene delivery to mature muscle fibers. J Virol 1996;70:8117– 8123. Hudecki MS, Pollina CM, Granchelli JA, Daly MK, Byrnes T, Wong JC, Hsiao JC: Strength and endurance in the therapeutic evaluation of prenisolone-treated MDX mice. Res Commun Chem Pathol Pharmacol 1993;79:45–60. Hudecki MS, Pollina CM, Heffner RR, Bhargava AK: Enhanced functional ability in drug-treated dystrophic chickens: Trial results with indomethacin, diphenylhydantoin, and prednisolone. Exp Neurol 1981;73:173–185. Jooss K, Yang Y, Fisher KJ, Wilson JM: Transduction of dendritic cells by DNA viral vectors directs the immune response to transgene products in muscle fibers. J Virol 1998;72:4212–4223. Kochanek S, Clemens PR, Mitani K, Chen HH, Chan S, Caskey CT: A new adenoviral vector: Replacement of all viral coding sequences with 28 kb of DNA independently expressing both full-length dystrophin and beta-galactosidase. Proc Natl Acad Sci USA 1996;93:5731–5736. Koenig M, Hoffman EP, Bertelson CJ, Monaco AP, Feener C, Kunkel LM: Complete cloning of the Duchenne muscular dystrophy (DMD) cDNA and preliminary genomic organization of the DMD gene in normal and affected individuals. Cell 1987;50:509–517. Kornegay JN, Tuler SM, Miller DM, Levesque DC: Muscular dystrophy in a litter of golden retriever dogs. Muscle Nerve 1988;11:1056–1064. Lefaucheur JP, Pastoret C, Sebille A: Phenotype of dystrophinopathy in old mdx mice. Anat Rec 1995; 242:70–76. Li J, Dressman D, Tsao YP, Toyo-oka T, Hoffman EP, Xiao X: rAAV vector mediated sarcoglycan gene transfer in a hamster model for limb girdle muscular dystrophy. Gene Ther 1999;6:74–82. Lochmuller H, Petrof BJ, Pari G, Larochelle N, Dodelet V, Wang Q, Allen C, Prescott S, Massie B, Nalbantoglu J, Karpati G: Transient immunosuppression by FK506 permits a sustained high-level dystrophin expression after adenovirus-mediated dystrophin minigene transfer to skeletal muscles of adult dystrophic (mdx) mice. Gene Ther 1996;3:706–716. Megeney LA, Kablar B, Garrett K, Anderson JE, Rudnicki MA: MyoD is required for myogenic stem cell function in adult skeletal muscle. Genes Dev 1996;10:1173–1183. Megeney LA, Kablar B, Perry RL, Ying C, May L, Rudnicki MA: Severe cardiomyopathy in mice lacking dystrophin and MyoD. Proc Natl Acad Sci USA 1999;96:220–225. Nigro V, Okazaki Y, Belsito A, Piluso G, Matsuda Y, Politano L, Nigro G, Ventura C, Abbondanza C, Molinari AM, Acampora D, Nishimura M, Hayashizaki Y, Puca GA: Identification of the Syrian hamster cardiomyopathy gene. Hum Mol Genet 1997;6:601–607. Pastoret C, Sebille A: mdx mice show progressive weakness and muscle deterioration with age. J Neurol Sci 1995;129:97–105. Pulido SM, Passaquin AC, Leijendekker WJ, Challet C, Wallimann T, Ruegg UT: Creatine supplementation improves intracellular Ca2+ handling and survival in mdx skeletal muscle cells. FEBS Lett 1998; 439:357–362. Rafael JA, Tinsley JM, Potter AC, Deconinck AE, Davies KE: Skeletal muscle-specific expression of a utrophin transgene rescues utrophin-dystrophin deficient mice. Nat Genet 1998;19:79–82. Sakamoto A, Ono K, Abe M, Jasmin G, Eki T, Murakami Y, Masaki T, Toyo-oka T, Hanaoka F: Both hypertrophic and dilated cardiomyopathies are caused by mutation of the same gene, deltasarcoglycan, in hamster: An animal model of disrupted dystrophin-associated glycoprotein complex. Proc Natl Acad Sci USA 1997;94:13873–13878. Sicinski P, Geng Y, Ryder-Cook AS, Barnard EA, Darlison MG, Barnard PJ: The molecular basis of muscular dystrophy in the mdx mouse: A point mutation. Science 1989;244:1578–1580. Tezak Z, Kanneboyina N, Plotz P, Hoffman EP: Adeno-associated virus genomes in human skeletal muscle: High frequency in muscle, and possible protection from polymyositis. Submitted. Thomas GD, Sander M, Lau KS, Huang PL, Stull JT, Victor RG: Impaired metabolic modulation of alpha-adrenergic vasoconstriction in dystrophin-deficient skeletal muscle. Proc Natl Acad Sci USA 1998;95:15090–15095. Tinsley J, Deconinck N, Fisher R, Kahn D, Phelps S, Gillis JM, Davies K: Expression of full-length utrophin prevents muscular dystrophy in mdx mice. Nat Med 1998;4:1441–1444.

Hoffman/Buyse

10

Tinsley JM, Potter AC, Phelps SR, Fisher R, Trickett JI, Davies KE: Amelioration of the dystrophic phenotype of mdx mice using a truncated utrophin transgene. Nature 1996;384:349–353. Watchko JF, Li J, Hoffman EP, Xiao X: Contractile properties of Bio 14.6 delta-sarcoglycan-deficient hamster tibialis anterior muscle. Muscle Nerve, in press. Winand NJ, Edwards M, Pradhan D, Berian CA, Cooper BJ: Deletion of the dystrophin muscle promoter in feline muscular dystrophy. Neuromuscul Disord 1994;4:433–445. Xiao X, Li J, Samulski RJ: Efficient long-term gene transfer into muscle tissue of immunocompetent mice by adeno-associated virus vector. Virology 1996;70:8098–8108. Xiao X, Li J, Samulski RJ: Production of high-titer recombinant adeno-associated virus vectors in the absence of helper adenovirus. J Virol 1998;72:2224–2232. Xiao X, Li J, Tsao YP, Dressman D, Hoffman EP, Watchko JF: Full functional rescue of a complete muscle (TA) in dystrophic hamsters by AAV vector-directed gene therapy. J Virol 2000;74:1436–1442.

Dr. Eric P. Hoffman, Research Center for Genetic Medicine, Children’s National Medical Center, George Washington University, 111 Michigan Ave. NW, Washington, DC 20010 (USA) Tel. +1 202 884 6011, Fax +1 202 884 6014, E-Mail [email protected]


11


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

Use of Normal and Genetically Modified Myoblasts for the Treatment of Myopathies Daniel Skuk, Jacques P. Tremblay Unite´ de Recherche en Geńe´tique Humaine, Centre de Recherche de Pavillon Centre Hospitalier de l’Universite´ Laval, CHUQ, et Faculte´ de Me´decine de l’Universite´ Laval, Ste-Foy, Que´., Canada

The Rationale of Myoblast Transplantation Genetic myopathies are characterized by different degrees of muscle weakness determined by intrinsic muscular defects of genetic origin. Although the underlying genetic defect of most genetic myopathies was unknown a decade ago, efforts in molecular biology led to the identification of the genetic basis of many diseases [1]. The first step in understanding the molecular basis of genetic myopathies was the discovery of the molecular cause of Duchenne muscular dystrophy (DMD). The severity and frequency of this disease made DMD a stereotype for all severe genetic myopathies, and most of the efforts in neuromuscular research were directed towards this disease. The discovery of the molecular basis of a disease must be followed by the development of its treatment. For many genetic diseases, the solution is to replace the defective gene with a normal one, and this concept is the basis of gene therapy [2]. Nevertheless, considering the muscle as a target for a genetic treatment, its particularities provide other possibilities to incorporate the affected protein into the cells. Skeletal muscles are composed of long syncytial elements (myofibres) that cannot enter the mitotic cycle. During adult life, precursor myogenic cells (called satellite cells) are quiescent in the periphery of mature myofibres [3]. When a myofibre is damaged, satellite cells proliferate and fuse to repair the damaged segment or replace it altogether [4]. These satellite cells can be proliferated in vitro, where they are called myoblasts. The existence of these cells has led to the idea that myoblast

transplantation (MT) into a diseased muscle could contribute to form new and healthy fibres [5].

First Experiments in MT As early as in 1978, Partridge et al. [5] suggested that the intramuscular injection of muscle precursor cells could provide a treatment for recessive inherited myopathies. However, some of the first experiments involving intramuscular injection of satellite cells were not performed as a therapeutic approach but, rather, as an experimental tool for basic studies of muscle biology [6, 7]. In 1979, Lipton and Schultz [6] implanted autologous cloned myoblasts into muscles and demonstrated that these cells participated in the regeneration of muscle fibres. The developmental role of myoblasts during embryogenesis was also investigated through MT [7]. Watt [8] and Watt et al. [9] showed, in the early 1980s, that muscle precursor cells obtained from enzymatically disaggregated neonatal muscles fused and formed hybrid fibres after their injection into regenerating mouse muscles. This group investigated cyclosporin A as the first immunosuppressive treatment for MT in mouse [10]. The transplantation of mesenchymal cells from mouse embryos into dystrophic mice was also studied as an experimental approach for the treatment of dystrophic muscles [11]. Thereafter, myoblasts injected into completely cryodamaged muscles were shown to repopulate almost 70% of these muscles [12]. Moreover, MT was tested with variable success in mouse models of different myopathies, such as phosphorylase kinase deficiency [13], the mdx mice [14, 15] and the dy/dy mice [16].

Early Clinical Trials Inspired by these first experiments, several clinical MT trials were soon conducted by various research teams [17–24]. Unfortunately, the clinical trials obtained very limited results. Functional tests showed, at best, a transient strength increase in a few patients [18]. The presence of donor dystrophin RNA was demonstrated in the transplanted muscles [17] as well as a significant but small increase in dystrophin-positive fibres [18, 22, 23]. The presence of 10.3% of dystrophin-positive fibres formed by the donor myoblasts in the transplanted muscle of a patient is currently the best histological result obtained from these clinical trials [22]. Donor nuclei in some of the transplanted patients were also confirmed by fluorescence in situ hybridization, using a probe specific for a dystrophin exon, which was absent in the patient [25].

Use of Normal and Genetically Modified Myoblasts for the Treatment of Myopathies

13

The only group claiming to have had a modest success with their clinical trials was the Cell Therapy Research Foundation [26, 27]. This clinic reported slight functional improvement and the presence of dystrophin in the transplanted muscles of some patients. Nonetheless, the functional improvement was not correctly evaluated since it failed to exclude a placebo effect or the beneficial effect of immunosuppression [28]. On the other hand, it was recently demonstrated that the dystrophin in one of these patients did not correspond to the transplanted myoblasts but was, in fact, the product of a back-mutation [29]. Some criticisms over the clinical trials came as a result of insufficient prior animal research [30]. Thereafter, certain research groups reexamined the MT-related problems using experimental models.

Immunobiology of MT The role of the immune system in MT was underestimated in some of the clinical trials. An immune response was considered unlikely because it was previously shown that major histocompatibility complex (MHC) molecules were not present on normal mature myofibres [31–33]. Nevertheless, MHC was expressed by muscle fibres under certain conditions, such as inflammatory reaction, muscle regeneration, and in DMD patients [32–34]. Moreover, it was later observed that myoblasts express MHC [35–38]. Evidence of cellular rejection following allo- and xeno-MTs was observed in unimmunosuppressed mice [39–41]. Muscles injected with myoblasts were infiltrated by CD4+ and CD8+ lymphocytes [39, 40, 42]. These lymphocytes expressed granzyme B, an enzyme released to damage the target cells [43]. Moreover, transplantation of human myoblasts into severe combined immunodeficient (SCID) mice led to many myofibres that contained human dystrophin [44–46]. Human transplanted myoblasts could then form muscle fibres when the specific immune response is avoided. Myoblasts obtained from MHC-compatible mice, but from a strain different from the host, were also rejected by immunocompetent hosts [47]. Rejection due to minor antigens was also observed after syngeneic MT from males into females, although slower than that observed following MHC-incompatible MTs [47]. Moreover, syngeneic transplantation of normal myoblasts in mdx mice led to specific immune responses against dystrophin, as explained later. There is cumulating evidence that myoblasts themselves are antigen-presenting cells. Following stimulation with c-interferon, a cytokine present during the inflammatory reaction, myoblasts express MHC molecules class I and II [36, 48, 49] and cell adhesion molecules involved in the activation of lymphocytes [50–52]. Myoblasts can process and present the same antigens as antigen-

Skuk/Tremblay

14

presenting cells and do increase secretion of IL-2 by lymphocytes [53]. It is, therefore, not surprising that they are rapidly rejected, following transplantation of myoblasts incompatible with major or minor antigens.

Immunosuppression in MT Following the demonstration of the immune response after MT, attention was focused on identifying an adequate immunosuppressive treatment. It was observed that the degree of success of MT depended on the immunosuppressive treatment [54]. The immunosuppressive agents used in clinical trials were cyclosporin A [17, 22] and cyclophosphamide [21]. However, in some animal experiments, cyclosporin A could not suppress the cellular immune reaction after MT [44]. Cyclophosphamide-immunosuppressed animals did not exhibit hybrid myofibres after MT, and it was proposed that this drug killed the transplanted myoblasts by its antiproliferative properties [55]. Rapamycin was shown to control the immune rejection after MT [56], but the best results were obtained with FK506 [57]. Using FK506 following transplantation of normal myoblasts expressing the gene of b-galactosidase, up to 95% of the muscle fibres in mdx mouse muscles expressed both b-galactosidase and dystrophin [57]. FK506 was also found very effective in controlling the immune response after MT in non-human primates [42, 58, 59]. In monkeys, the immunosuppressive benefits of FK506 were observed up to 1 year following MT [59a]. Successful MTs were also obtained using a combination of monoclonal antibodies directed against lymphocyte adhesion molecules [52].

Transplantation of Genetically Modified Autologous Myoblasts Although used clinically for immunosuppression, the administration of FK506 is limited by adverse effects such as nephrotoxicity [60], neurotoxicity [61], pancreatic toxicity [62], and posttransplantation lymphoproliferative disorders [63]. An alternative approach to avoid long-term immunosuppression is to transplant the patient’s own genetically corrected myoblasts. For this approach, the dystrophin gene must be introduced into the dystrophic myoblasts before their autologous transplantation. This has recently been done with success in mdx mice using adenoviral vectors containing the human dystrophin minigene [64] or the full-length dystrophin [65]. Adenoviral-mediated dystrophin transfer was successfully done in human DMD myoblasts, and these corrected myoblasts transplanted in SCID mice formed humandystrophin-positive muscle fibres [66]. However, the autotransplantation of


15

myoblasts infected with an adenoviral vector could trigger possible immune reactions against the viral vector itself, as observed following direct gene therapy with this vector [67, 68]. This immune reaction may eventually be controlled by a transient immunosuppression with FK506 [68]. Although this autologous MT is presumed to have no specific immune reactions, the introduction of dystrophin into DMD patients could be potentially immunogenic. Antibodies reacting with dystrophin were observed in some of the transplanted patients [18, 69]. There is also some evidence of dystrophin immunogenicity after normal MT into mdx mice. Syngeneic transplantation of normal myoblasts into mdx mice triggered the production of antibodies against dystrophin [70]. These antibodies were detected 1 month following MT, but dystrophin-positive fibres were present up to 9 months after MT and CD4+ and CD8+ lymphocyte infiltration were not observed [47, 70]. In another study, however, the same kind of transplantation led to a slow cellular rejection of the dystrophin-formed fibres, mediated by cytotoxic T lymphocytes and restricted to H-2Kb [71]. Three epitopes were identified as the principal antigenic targets of dystrophin in this study. Another problem of autotransplanting myoblasts from the DMD patient itself is the low proliferative capacity of these myoblasts [72]. The muscle fibres in the DMD are submitted to recurrent cycles of degeneration-regeneration exhausting the proliferative capacity of satellite cells. Some strategies to increase the proliferative capacity of DMD myoblasts were proposed. One such strategy is the immortalization of cells with the large tumor (T) antigen of simian virus 40 (SV40) [73]. The introduction of the SV40 large T antigen under the control of the human vimentin promoter increased the proliferative capacity of myoblasts, retaining their capacity to differentiate [74]. However, the success of transplanting myoblasts expressing the T antigen was significantly reduced compared with that observed with control myoblasts [unpubl. data]. This was attributed to an increased death of the T antigen-positive myoblasts following transplantation. An alternative approach to increase the proliferative capacity of cells is the introduction of the telomerase gene [75]. In recent experiments of our research group, the introduction of the telomerase gene in myoblasts led to better transplantation success than with the introduction of the T antigen [unpubl. data]. Another strategy to circumvent the low proliferative capacity of DMD myoblasts may be the transformation of other cell types into myoblasts. Intramuscular injection of dermal fibroblasts, genetically modified by introducing MyoD1 (a master regulator gene for myogenesis), was tested but provided limited results so far [76]. Recently, evidence has been presented suggesting that bone marrow cells could contain myogenic progenitors [77]. The use of these bone marrow-derived myogenic progenitors was suggested as a potential treatment of muscular dystrophies.

Skuk/Tremblay

16

Early Survival of Transplanted Myoblasts Most of the observations on myoblast survival after intramuscular injection demonstrated a rapid loss of these cells after transplantation. The use of myoblasts labeled by introducing the b-galactosidase gene showed a loss of 50–80% of the transplanted cells during the first 6 days that followed MT [78–82]. After labelling myoblasts with [14C]thymidine, instead of b-galactosidase, 99% of cell loss was reported after 4 days [83]. These different results may depend on the method used to evaluate cell death, since b-galactosidase detection does not only account for cell death, because the transgene will increase with the proliferating surviving cells. In spite of this, different results were obtained even with the same method, suggesting that early cell death may depend on the cell type used for transplantation [84]. Since the rapid death of injected myoblasts was detected following autotransplantation and transplantation in SCID and immunosuppressed mice, a specific immune reaction as the reason for this early cell death is ruled out [81]. Up until now, three explanations regarding the early death of the transplanted myoblasts have been suggested: the conditions of cell storage before transplantation [79], the specific survival of a subpopulation of myoblasts with stem cell-like characteristics [83], and the damage induced by the invasion of inflammatory cells [80–82]. The last hypothesis proposed that the muscle damage induced by the injections and the presence of some necrosed cells trigger the influx of inflammatory cells that may produce a ‘bystander’ damage on the graft [80, 81]. This hypothesis was supported by the findings that an antiLFA-1 antibody decreased cellular mortality [80, 81], that myoblast loss is reduced in hosts that are deficient in neutrophils or in macrophages [82], and that myoblasts that are genetically engineered to express an inhibitor of IL-1 showed improved survival rates [84]. Complement was observed to be fixed spontaneously in vitro by human myoblasts [85] and was suspected to be one of the reasons of the cell death. Nevertheless, in vivo observations showed that complement was fixed only intracellularly by necrosed cells, and the immediate cell death observed after MT did not decrease in complement-depleted mice [86].

Diffusion of Transplanted Myoblasts MT is also confronted with the limited capacity of myoblasts to migrate into the muscle tissue where they are injected. In monkeys, injections of myoblasts resuspended in a saline solution led to limited bands of hybrid fibres in the transplanted tissue, showing that the myoblasts fused only with


17

the fibres located near the injection trajectories but not so far [42]. It was reported that the migration of myoblasts was only 1.6 mm in mice [87] and unique intramuscular injections of these cells showed that they spread on a limited surface of 2.6–9.5¶105 lm2 [88]. The only current possibility for a successful MT in large muscles, therefore, is to perform multiple injections very close to each other [59a]. Some authors have shown that myoblasts can migrate under some experimental conditions. Myoblasts were capable of migrating into fatty connective tissue implanted into mouse muscles [89]. Myoblast injections into irradiated muscles of mdx mice showed the presence of donor dystrophin-positive myofibres in the adjacent muscle at the later stages (250 days) following MT [90]. Myoblasts from neighbouring normal muscles were also capable of invading and repopulating freeze-killed muscles [12, 91]. Cells with the capacity to migrate into different tissues, such as leucocytes and tumour cells, secrete metalloproteinases, a group of enzymes that degrade the extracellular matrix [92]. The migration of myoblasts may be improved by increasing the secretion of metalloproteinases. Incubation of myoblasts with a high dose of basic fibroblast growth factor, a factor that can trigger the secretion of metalloproteinases, increased by 4-fold the success of MT [93]. The addition of concanavalin A to the myoblast culture increased by more than 3-fold the dispersion of these cells in a muscle [88]. This effect was attributed to the secretion of metalloproteinases by fibroblasts in the primary culture. To avoid multiple cell injections into the muscles, intra-arterial administration of myoblasts could be used [94]. Although this technique could have the potential advantage of reaching important muscles that are relatively inaccessible to injection, such as the diaphragm, few hybrid fibres were detected in previously damaged muscles only.

MT in Large Animals MT must prove to be effective under conditions approaching the clinical situation and not only in the small muscle of a mouse. Therefore, studies of MT in large animals are of crucial importance. Muscular-dystrophic dogs represent a good model for the study of MT due to an absence of dystrophin [95]. However, only one article exists on MT in dystrophic dogs, where the absence of positive results can be attributed to the lack of immunosuppression [96]. Better results of MT were obtained in normal dogs treated with FK506 at low doses combined with cyclosporin A and mycophenolate mofetil [97].

Skuk/Tremblay

18

Monkeys are anatomically and immunologically closer to humans, and it can be expected that the results obtained with this model could be extrapolated to patients. Experiments in monkeys demonstrated that the specific immune response is developed against the transplanted myoblasts and against the myofibres created by their fusion, and that this rejection can be controlled by FK506 [42, 58, 59]. Recently, our group demonstrated that it is possible to achieve successful MTs in the whole biceps of monkeys [59a]. Up to 75% of hybrid fibres were obtained in the transplanted muscles 1 month after MT, and the hybrid fibres were present in the transplanted muscles up to 1 year after MT. Considering the size of a monkey biceps, and the immunological similitude with humans, these experiments provide some parameters that could make this technique applicable to humans.

Could MT Improve Patients? The effectiveness of MT must not only be demonstrated by the presence of the reporter gene in many muscle fibres, but by its capacity to stop histological degeneration and progressive strength loss in the disease muscles as well. There is already some evidence that supports potential beneficial effects of MT. In mdx mice, the lack of dystrophin makes muscle fibres more vulnerable to damage resulting from eccentric exercise [98, 99]. When MT was performed 1 month before submitting the mdx mice to eccentric exercise, myofibre damage was only observed in dystrophin-negative fibres [100]. These results suggest that MT protected the muscle tissue of mdx mice from mechanical stress, the mechanism considered responsible for triggering myofibre necrosis in DMD. It was also observed that the number of dystrophin-positive fibres remained unchanged, 35–250 days following MT, while the number of dystrophin-negative fibres progressively decreased [90]. This was attributed to the protective role of the donor dystrophin. Most of MT experiments demonstrated that myoblasts fuse with previously existing myofibres. Nevertheless, if the transplanted myoblasts could not regenerate new fibres when the muscles are replaced by fibrosis and fat tissue, MT would be a potential treatment only during the first years of DMD patients. Can MT restore the muscular tissue and strength of patients with advanced disease? Although few studies on this aspect of MT exist, preliminary results are encouraging. It was demonstrated that myoblasts could invade the fatty connective tissue to form myotubes and new muscle fibres [89]. Moreover, myoblasts could form ectopic and functional muscles without the need of a previous tissular support when they were implanted under the skin [101]. MT was shown to restore both muscle mass and force following irreversible muscle


19

damage [102, 103], as well as increase twitch and tetanic tension in normal regenerating muscles [104]. Some evidence suggests that some transplanted myoblasts survive as muscle precursor cells, which can participate in subsequent muscle regeneration [105, 106]. This is important because it means that these cells can provide a permanent source of normal cells to replace the damaged muscle fibres in the dystrophic patients.

MT in Other Myopathies Although most of the attention on MT was given to the treatment of DMD, other genetic myopathies could benefit from this technique as well. Congenital Dystrophy. Merosin is a component of the skeletal muscle basal lamina and its absence determines a type of usually severe congenital dystrophy [107]. An animal model for this disease is the dy/dy mouse, which shows an absence of merosin and a severe and progressive dystrophy [108]. Transplantation of normal myoblasts into dy/dy mice was shown to restore merosin expression in the muscle fibres [109]. Metabolic Myopathies. Hybrid fibres formed in vitro by the fusion of normal myoblasts with those from a glucose-6-phosphate dehydrogenase deficient mouse contained high levels of this enzyme, suggesting that MT could restore normal enzymatic activity in metabolic myopathies [110]. Similar in vitro experiments suggested that MT could restore acid a-glucosidase in the type II glycogen storage disease [111]. MT was also tested in phosphorylase kinase-deficient mice, but results were limited [13].

Conclusion MT remains the only treatment that could eventually not only introduce the dystrophin gene into the already existing muscle fibres but increase their size and perhaps their number as well, thereby increasing the strength of myopathic patients. Although initial trials have not produced clinically acceptable results, ongoing research during of the last decade provided a better understanding of the problems associated with MT. This cumulative knowledge will permit a more effective MT, applicable to myopathic patients. Research is still being pursued on many potential further improvements of MT, such as avoiding sustained immunosuppression and improving the migration as well as the survival of myoblasts in the host muscle.

Skuk/Tremblay

20

References 1 2 3 4 5 6 7 8 9 10 11 12 13 14

15 16 17

18

19

20

21

22

23

Chance PF, Ashizawa T, Hoffman EP, Crawford TO: Molecular basis of neuromuscular diseases. Phys Med Rehabil Clin North Am 1998;9:49–81. Mulligan RC: The basic science of gene therapy. Science 1993;260:926–932. Campion DR: The muscle satellite cell: A review. Int Rev Cytol 1984;87:225–251. Carlson BM, Faulkner JA: The regeneration of skeletal muscle fibers following injury: A review. Med Sci Sports Exerc 1983;15:187–198. Partridge TA, Grounds M, Sloper JC: Evidence of fusion between host and donor myoblasts in skeletal muscle grafts. Nature 1978;273:306–308. Lipton BH, Schultz E: Developmental fate of skeletal muscle satellite cells. Science 1979;205: 1292–1294. Womble MD, Bonner PH: Developmental fate of a distinct class of chick myoblasts after transplantation of cloned cells into quail embryos. J Embryol Exp Morphol 1980;58:119–130. Watt DJ: Factors which affect the fusion of allogeneic muscle precursor in vivo. Neuropathol Appl Neurobiol 1982;8:135–147. Watt DJ, Morgan JE, Partridge TA: Use of mononuclear precursor cells to insert allogeneic genes into growing mouse muscles. Muscle Nerve 1984;7:741–750. Watt DJ, Partridge TA, Sloper JG: Cyclosporin a as a means of preventing rejection of skeletal muscle allografts in mice. Transplantation 1981;31:266–271. Law PK: Beneficial effects of transplanting normal limb-bud mesenchyme into dystrophic mouse muscles. Muscle Nerve 1982;5:619–627. Morgan JE, Coulton GR, Partridge TA: Muscle precursor cells invade and repopulate freeze-killed muscles. J Muscle Res Cell Motil 1987;8:386–396. Morgan JE, Watt DJ, Sloper JC, Partridge TA: Partial correction of an inherited biochemical defect of skeletal muscle by grafts of normal muscle precursor cells. J Neurol Sci 1988;86:137–147. Partridge TA, Morgan JE, Coulton GR, Hoffman EP, Kunkel LM: Conversion of mdx myofibres from dystrophin-negative to -positive by injection of normal myoblasts. Nature 1989;337: 176–179. Morgan JE, Hoffman EP, Partridge TA: Normal myogenic cells from newborn mice restore normal histology to degenerating muscles of the mdx mouse. J Cell Biol 1990;111:2437–2449. Law PK, Goodwin TG, Wang MG: Normal myoblast injections provide genetic treatment for murine dystrophy. Muscle Nerve 1988;11:525–533. Gussoni E, Pavlath GK, Lanctot VK, Sharma RG, Miller L, Steinman L, Blau HM: Normal dystrophin transcripts detected in DMD patients after myoblast transplantation. Nature 1992;356: 435–438. Huard J, Bouchard JP, Roy R, Malouin F, Dansereau G, Labrecque C, Albert N, Richards CL, Lemieux B, Tremblay JP: Human myoblast transplantation: Preliminary results of four cases. Muscle Nerve 1992;15:550–560. Tremblay JP, Bouchard JP, Theáu D, Cottrell F, Collin H, Tremblay M, Richards CL, Malouin F, Tome´ F, Fardeau M: Myoblast transplantation between monozygotic twin girls which are carriers of Duchenne muscular dystrophy. Neuromuscul Disord 1993;3:583–593. Tremblay JP, Malouin F, Roy R, Huard J, Bouchard JP, Satoh A, Richards CL: Results of a triple blind clinical study of myoblast transplantations without immunosuppressive treatment in young boys with Duchenne muscular dystrophy. Cell Transplant 1993;2:99–112. Karpati G, Ajdukovic D, Arnold D, Glendhill RB, Guttmann R, Holland P, Koch PA, Shoubridge E, Spence D, Vanasse M, Watters GV, Abrahamowicz M, Duff C, Worton RG: Myoblast transfer in Duchenne muscular dystrophy. Ann Neurol 1993;34:8–17. Mendell JR, Kissel JT, Amato AA, King W, Signore L, Prior TW, Sahenk S, Besson S, McAndrew PE, Rice R, Nagaraja H, Stephens R, Lantry L, Morris GE, Burghes AHM: Myoblast transfer in the treatment of Duchenne’s muscular dystrophy. N Engl J Med 1995;333:832–838. Miller RG, Sharma KR, Pavlath GK, Gussoni E, Mynhier M, Yu P, Lanctot AM, Greco CM, Steiman L, Blau HM: Myoblast implantation in Duchenne muscular dystrophy: The San Francisco Study. Muscle Nerve 1997;20:469–478.


21

24

25 26

27

28 29 30 31 32

33 34

35

36 37 38 39 40 41 42

43 44 45 46

47

Neumeyer AM, Cros D, McKenna-Yasek D, Zawadzka A, Hoffman EP, Pegoraro E, Hunter RG, Munsat TL, Brown RH: Pilot study of myoblast transfer in the treatment of Becker muscular dystrophy. Neurology 1998;51:589–592. Gussoni E, Blau HM, Kunkel LM: The fate of individual myoblasts after transplantation into muscles of DMD patients. Nat Med 1997;3:970–977. Law PK, Goodwin TG, Fang Q, Deering MB, Duggirala V, Larkin C, Florendo JA, Kirby DS, Li HJ, Chen M, Cornett J, Li LM, Shirzad A, Quinley T, Yoo TJ, Holcomb R: Cell transplantation as an experimental treatment for Duchenne muscular dystrophy. Cell Transplant 1993;2:485–505. Law PK, Goodwin TG, Fang Q, Hall TL, Quinley T, Vastagh G, Duggirala V, Larkin C, Florendo JA, Li L, Jackson T, Yoo TJ, Chase N, Neel M, Krahn T, Holcomb R: First human myoblast transfer therapy continues to show dystrophin after 6 years. Cell Transplant 1997;6:95–100. Thompson L: Cell-transplant results under fire. Science 1992;257:472–474. Partridge T, Lu QL, Morris G, Hoffman E: Is myoblast transplantation effective? Nat Med 1998; 4:1208–1209. Thompson L: Researchers call for time out on cell-transplant research. Science 1992;257:738. Ponder BAJ, Wilkinson MM, Wood M, Westwood JH: Immunohistochemical demonstration of H2 antigens in mouse tissue sections. J Histochem Cytochem 1983;31:911–919. Appleyard ST, Dunn MJ, Dubowitz V, Rose ML: Increased expression of HLA ABC class I antigens by muscle fibers in Duchenne muscular dystrophy, inflammatory myopathy, and other neuromuscular disorders. Lancet 1985;i:361–363. Karpati G, Pouilot Y, Carpenter S: Expression of immunoreactive major histocompatibility complex products in human skeletal muscles. Ann Neurol 1988;23:64–72. Emslie-Smith AM, Arahata K, Engel AG: Major histocompatibility complex class I antigen expression, immunolocalization of interferon subtypes, and T cell-mediated cytotoxicity in myopathies. Hum Pathol 1989;20:224–231. Mantegazza R, Hughes SM, Mitchell D, Travis M, Blau HM, Steinman L: Modulation of MHC class II antigen expression in human myoblasts after treatment with IFN-gamma. Neurology 1991; 41:1128–1132. Roy R, Dansereau G, Tremblay JP, Belles-Isles M, Labrecque C, Bouchard JP: Expression of major histocompatibility complex (MHC) antigens on human myoblasts. Transplant Proc 1991;23:799–801. Cifuentes-Diaz C, Delaporte C, Dautreáux B, Charron D, Fardeau M: Class II MHC antigens in normal human skeletal muscle. Muscle Nerve 1992;15:295–302. Bao S, Dos Remedios CG, King NJC: Ontogeny of major histocompatibility complex antigen expression on cultured human embryonic skeletal myoblasts. Transplantation 1994;58:585–591. Gue´rette B, Asselin I, Roy R, Tremblay JP: Lymphocyte infiltration following allo- and xenomyoblast transplantation in mdx mice. Muscle Nerve 1995;18:39–51. Irintchev A, Zweyer M, Wernig A: Cellular and molecular reactions in mouse muscles after myoblast implantation. J Neurocytol 1995;24:319–331. Wernig A, Irintchev A, Lange G: Functional effects of myoblast implantation into histoincompatible mice with or without immunosuppression. J Physiol (Lond) 1995;484:493–504. Skuk D, Roy B, Goulet M, Tremblay JP: Successful myoblast transplantation in primates depends on appropriate cell delivery and induction of regeneration in the host muscle. Exp Neurol 1999; 155:1–9. Gue´rette B, Roy R, Tremblay M, Asselin I, Kinoshita I, Tremblay JP: Increased granzyme B mRNA following alloincompatible myoblast transplantation. Transplantation 1995;60:1011–1016. Huard J, Roy R, Gue´rette B, Verreault S, Tremblay G, Tremblay JP: Human myoblast transplantation in immunodeficient and immunosuppressed mice: Evidence of rejection. Muscle Nerve 1994;17:224–234. Huard J, Verreault S, Roy R, Tremblay JP: High efficacy of muscle regeneration following human myoblast transplantation in SCID mice. J Clin Invest 1994;93:586–599. Skuk D, Furling D, Bouchard J-P, Goulet M, Roy B, Lacroix Y, Vilquin J-T, Tremblay JP, Puymirat J: Transplantation of human myoblasts in SCID mice as a potential muscular model for myotonic dystrophy. J Neuropathol Exp Neurol 1999;58:921–931. Boulanger A, Asselin I, Roy R, Tremblay JP: Role of non-major histocompatibility complex antigens in the rejection of transplanted myoblasts. Transplantation 1997;63:893–899.

Skuk/Tremblay

22

48 49 50 51 52 53 54

55 56 57 58 59

59a 60 61

62

63

64 65

66

67 68

69

Hohlfeld R, Engel AG: Induction of HLA-DR expression on human myoblasts with interferongamma. Am J Pathol 1990;136:503–508. Hohlfeld R, Engel AG: HLA expression in myoblasts. Neurology 1991;41:2015. Goebels N, Michaelis D, Wekerle H, Hohlfeld R: Human myoblasts as antigen-presenting cells. J Immunol 1992;149:661–667. Michaelis D, Goebels N, Hohlfeld R: Constitutive and cytokine-induced expression of human leukocyte antigens and cell adhesion molecules by human myotubes. Am J Pathol 1993;143:1142–1149. Gue´rette B, Wood K, Roy R, Tremblay JP: Efficient myoblast transplantation in mice immunosuppressed with monoclonal antibodies and CTLA4 Ig. Transplant Proc 1997;29:1932–1934. Garlepp MJ, Chen W, Tabarias H, Baines M, Brooks A, McCluskey J: Antigen processing and presentation by a murine myoblast cell line. Clin Exp Immunol 1995;102:614–619. Vilquin J-T, Asselin I, Gue´rette B, Kinoshita I, Lille S, Roy R, Tremblay JP: Myoblast allotransplantation in mice: The degree of success varies depending on the efficacy of various immunosuppressive treatments. Transplant Proc 1994;26:3372–3373. Vilquin J-T, Kinoshita I, Roy R, Tremblay JP: Cyclophosphamide immunosuppression does not permit successful myoblast allotransplantation in mouse. Neuromuscul Disord 1995;5:511–517. Vilquin J-T, Asselin I, Gue´rette B, Kinoshita I, Lille S, Roy R, Tremblay JP: Successful myoblast allotransplantation in mdx mice using rapamycin. Transplantation 1995;59:422–449. Kinoshita I, Vilquin J-T, Gue´rette B, Asselin I, Roy R, Tremblay JP: Very efficient myoblast allotransplantation in mice under FK506 immunosuppression. Muscle Nerve 1994;17:1407–1415. Kinoshita I, Vilquin J-T, Gravel C, Roy R, Tremblay JP: Myoblast allotransplantation in primates. Muscle Nerve 1995;18:1217–1218. Kinoshita I, Roy R, Dugre´ FJ, Gravel C, Vilquin J-T, Roy B, Goulet M, Asselin I, Tremblay JP: Myoblast transplantation in monkeys: Control of immune response by FK506. J Neuropathol Exp Neurol 1996;55:687–697. Skuk D, Goulet M, Roy B, Tremblay JP: Myoblast transplantation in whole muscle of nonhuman primates. J Neuropathol Exp Neurol, in press. Klintmalm GB, Gonwa TA: Nephrotoxicity associated with cyclosporine and FK506. Liver Transplant Surg 1995;1(suppl 1):11–19. Neu AM, Furth SL, Case BW, Wise B, Colombani PM, Fivush BA: Evaluation of neurotoxicity in pediatric renal transplant recipients treated with tacrolimus (FK506). Clin Transplant 1997;11: 412–414. Uchida K, Tominaga Y, Haba T, Katayama A, Ichimori T, Yamada K, Hibi Y, Uemura O, Morozumi K, Takagi H: Decreasing pancreatic toxicity of tacrolimus by dosage reduction. Transplant Proc 1998;30:1276–1278. Cacciarelli TV, Green M, Jaffe R, Mazariegos GV, Jain A, Fung JJ, Reyes J: Management of posttransplant lymphoproliferative disease in pediatric liver transplant recipients receiving primary tacrolimus (FK506) therapy. Transplantation 1998;66:1047–1052. Moisset P-A, Gagnon Y, Karpati G, Tremblay JP: Expression of human dystrophin following the transplantation of genetically modified mdx myoblasts. Gene Ther 1998;5:1340–1346. Floyd SS, Clemens PR, Ontell MR, Kochanek S, Day CS, Yang J, Hauschka SD, Balkir L, Morgan J, Moreland MS, Feero GW, Epperly M, Huard J: Ex vivo gene transfer using adenovirus-mediated full-length dystrophin delivery to dystrophic muscles. Gene Ther 1998;5:19–30. Moisset P-A, Skuk D, Asselin I, Goulet M, Roy B, Karpati G, Tremblay JP: Successful transplantation of genetically corrected DMD myoblasts following ex vivo transduction with the dystrophin minigene. Biochem Biophys Res Commun 1998;247:94–99. Yang Y, Nunes FA, Berencsi K, Furth EE, Gonczol E, Wilson JM: Cellular immunity to viral antigens limits E1-deleted adenoviruses for gene therapy. Proc Natl Acad Sci USA 1994;91:4407–4441. Vilquin J-T, Gue´rette B, Kinoshita I, Roy B, Goulet M, Gravel C, Roy R, Tremblay JP: FK506 immunosuppression to control the immune reactions triggered by the first generation adenovirusmediated gene transfer. Hum Gene Ther 1995;6:1391–1401. Roy R, Tremblay JP, Huard J, Richards CL, Malouin F, Bouchard JP: Antibody formation after myoblast transplantation in Duchenne-dystrophic patients, donor HLA compatible. Transplant Proc 1993;25:995–997.


23

70

71

72

73 74

75

76

77 78

79 80 81

82 83

84 85 86 87 88 89 90 91 92

Vilquin J-T, Wagner E, Kinoshita I, Roy R, Tremblay JP: Successful histocompatible myoblast transplantation in dystrophin-deficient mdx mouse despite the production of antibodies against dystrophin. J Cell Biol 1995;131:975–988. Ohtsuka Y, Udaka K, Yamashiro Y, Yagita H, Okumura K: Dystrophin acts as a transplantation rejection antigen in dystrophin-deficient mice: Implications for gene therapy. J Immunol 1998;160: 4635–4640. Webster C, Blau HM: Accelerated age-related decline in replicative life-span of Duchenne muscular dystrophy myoblasts: Implications for cell and gene therapy. Somat Cell Mol Genet 1990;16:557– 565. Bryan TM, Reddel RR: SV40–induced immortalization of human cells. Crit Rev Oncog 1994;5: 331–357. Descheńes I, Chahine M, Tremblay J, Paulin D, Puymirat J: Increase in the proliferative capacity of human myoblasts by using the T antigen under the vimentin promoter control. Muscle Nerve 97;20:437–445. Bodnar AG, Ouellette M, Frolkis M, Holt SE, Chiu CP, Morin GB, Harley CB, Shay JW, Lichtsteiner S, Wright WE: Extension of life-span by introduction of telomerase into normal human cells. Science 1998;279:349–352. Huard C, Moisset P-A, Dicaire A, Merly F, Tardif F, Asselin I, Tremblay JP: Transplantation of dermal fibroblasts expressing MyoD1 in mouse muscles. Biochem Biophys Res Commun 1998;248: 648–654. Ferrari G, Cusella-De Angelis G, Coletta M, Paolucci E, Stornaiuolo A, Cossu G, Mavilio F: Muscle regeneration by bone marrow-derived myogenic progenitors. Science 1998;279:1528–1530. Huard J, Acsadi A, Jani B, Massie B, Karpati G: Comparison of the efficiency of gene transfer by genetically marked co-isogenic myoblasts vs uncultured satellite cells in normal and mdx mouse muscles. Muscle Nerve 1994;(suppl 1):S261. Rando TA, Pavlath GK, Blau HM: The fate of myoblasts following transplantation into mature muscle. Exp Cell Res 1995;220:383–389. Gue´rette B, Asselin I, Skuk D, Entman M, Tremblay JP: Control of inflammatory damage by antiLFA-1: Increase success of myoblast transplantation. Cell Transplant 1997;6:101–107. Gue´rette B, Skuk D, Ce´lestin F, Entman M, Tardif F, Asselin I, Roy B, Goulet M, Huard C, Tremblay JP: Prevention by anti-LFA-1 of acute myoblast death following transplantation. J Immunol 1997; 159:2522–2531. Merly F, Huard C, Asselin I, Robbins PD, Tremblay JP: Anti-inflammatory effect of transforming growth factor-beta1 in myoblast transplantation. Transplantation 1998;65:793–799. Beauchamp JR, Morgan JE, Pagel CN, Partridge TA: Dynamics of myoblast transplantation reveal a discrete minority of precursors with stem cell-like properties as the myogenic source. J Cell Biol 1999;144:1113–1122. Qu Z, Balkir L, van Deutekom JCT, Robbins PD, Pruchnic R, Huard J: Development of approaches to improve cell survival in myoblast transfer therapy. J Cell Biol 1998;142:1257–1267. Gasque P, Morgan BP, Legoedec J, Chan P, Fontaine M: Human skeletal myoblasts spontaneously activate allogeneic complement but are resistant to killing. J Immunol 1996;156:3402–3411. Skuk D, Tremblay JP: Complement deposition and cell death after myoblast transplantation. Cell Transplant 1998;7:427–434. Fan Y, Beilharz MW, Grounds M: A potential alternative strategy for myoblast transfer therapy: The use of sliced muscle grafts. Cell Transplant 1996;5:421–429. Ito H, Hallauer PL, Hastings KEM, Tremblay JP: Prior culture with concanavalin A increases intramuscular migration of transplanted myoblast. Muscle Nerve 1998;21:291–297. Satoh A, Labrecque C, Tremblay JP: Myotubes can be formed within implanted adipose tissue. Transplant Proc 1992;24:3017–3019. Morgan JE, Pagel CN, Sherratt T, Partridge TA: Long-term persistence and migration of myogenic cells injected into pre-irradiated muscles of mdx mice. J Neurol Sci 1993;115:191–200. Phillips GD, Hoffman JR, Knighton DR: Migration of myogenic cells in the rat extensor digitorum longus muscle studied with a split autograft model. Cell Tissue Res 1990;262:81–88. Ennis BW, Matrilisian LM: Matrix degrading metalloproteinases. J Neurooncol 1994;18:105–109.

Skuk/Tremblay

24

93

94 95 96

97 98 99 100 101 102 103 104

105 106 107

108 109

110

111

Kinoshita I, Vilquin J-T, Tremblay JP: Pretreatment of myoblast cultures with basic fibroblast growth factor increases the efficacy of their transplantation in mdx mice. Muscle Nerve 1995;18: 834–841. Neumeyer AM, DiGregorio DM, Brown RH Jr: Arterial delivery of myoblasts to skeletal muscle. Neurology 1992;42:2258–2262. Cooper BJ: Animal models of Duchenne and Becker muscular dystrophy. Br Med Bull 1989;45: 703–718. Kornegay JN, Sharp NJH, Bartlett RJ, Van Camp SD, Burt CT, Hung WY, Kwock L, Roses AD: Golden retriever muscular dystrophy: Monitoring for success; in Eastwood AB, Karpati G, Griggs R (eds): Myoblast Transfer Therapy. New York, Plenum Publishing, 1990, pp 267–272. Ito H, Vilquin J-T, Skuk D, Roy B, Goulet M, Lille S, Dugre´ FJ, Asselin I, Roy R, Fardeau M, Tremblay JP: Myoblast transplantation in non-dystrophic dog. Neuromuscul Disord 1998;8:95–110. Brussee V, Tardif F, Tremblay JP: Muscle fibers of mdx mice are more vulnerable to exercise than those of normal mice. Neuromuscul Disord 1997;7:487–492. Vilquin J-T, Brussee V, Asselin I, Kinoshita I, Gingras M, Tremblay JP: Evidence of mdx mouse skeletal muscle fragility in vivo by eccentric running exercise. Muscle Nerve 1998;21:567–576. Brussee V, Merly F, Tardif F, Tremblay JP: Normal myoblast implantation in mdx mice prevents muscle damage by exercise. Biochem Biophys Res Commun 1998;250:321–327. Irintchev A, Rosenblatt JD, Cullen MJ, Zweyer M, Wernig A: Ectopic skeletal muscles derived from myoblasts implanted under the skin. J Cell Sci 1998;111:3287–3297. Alameddine HS, Louboutin JP, Dehaupas M, Se´bille A, Fardeau M: Functional recovery induced by satellite cell grafts in irreversibly injured muscles. Cell Transplant 1994;3:3–14. Irintchev A, Langer M, Zweyer M, Theisen R, Wernig A: Functional improvement of damaged adult mouse muscle by implantation of primary myoblasts. J Physiol (Lond) 1997;500:775–785. Arcila ME, Ameredes BT, DeRosimo JF, Washabaugh CH, Yang J, Johnson PC, Ontell M: Mass and functional capacity of regenerating muscle is enhanced by myoblast transfer. J Neurobiol 1997; 33:185–198. Yao SN, Kurachi K: Implanted myoblasts not only fuse with myofibers but also survive as muscle precursor cells. J Cell Sci 1993;105:957–963. Gross JG, Morgan JE: Muscle precursor cells injected into irradiated mdx mouse muscle persist after serial injury. Muscle Nerve 1999;22:174–185. Tome´ FMS, Evangelista T, Leclerc A, Sunada Y, Manole E, Estournet B, Barois A, Campbell KP, Fardeau M: Congenital muscular dystrophy with merosin deficiency. CR Acad Sci III 1994;317: 351–357. Sunada Y, Bernier SM, Kozak CA, Yamada Y, Campbell KP: Deficiency of Merosin in dystrophic dy mice and genetic linkage of laminin M chain gene to dy locus. J Biol Chem 1994;269:13729–13732. Vilquin J-T, Kinoshita I, Roy B, Goulet M, Engvall E, Tom F, Fardeau M, Tremblay JP: Partial laminin alpha2 chain restoration in alpha2 chain-deficient dy/dy mouse by primary muscle cell culture transplantation. J Cell Biol 1996;133:185–197. Meola G, Tremblay JP, Sansone V, Rotondo G, Radice S, Bresolin N, Huard J, Scarlato G: Muscle glucose-6-phosphate dehydrogenase deficiency: Restoration of enzymatic activity in hybrid myotubes. Muscle Nerve 1993;16:594–600. Zaretsky JZ, Candotti F, Boerkoel C, Adams EM, Yewdell JW, Blaese RM, Plotz PH: Retroviral transfer of acid alpha-glucosidase cDNA to enzyme-deficient myoblasts results in phenotypic spread of the genotypic correction by both secretion and fusion. Hum Gene Ther 1997;8:1555–1563.

Dr. Jacques P. Tremblay, Unite´ Recherche en Geńe´tique Humaine, Rc 9300, CHUQ-Pavillon CHUL, 2705 Boulevard Laurier, Ste-Foy, Que´. G1V 4G2 (Canada) E-Mail [email protected]


25


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

Disorders of the Sarcoglycan Complex (Sarcoglycanopathies) Carsten G. Bo¨nnemann Department of Neuropediatrics, University Childrens’ Hospital, Go¨ttingen, Germany

Limb-Girdle Muscular Dystrophies After years of inconsistent use, the diagnostic category of limb-girdle muscular dystrophy (LGMD) more recently has undergone a reevaluation [1], not least because of the great progress that has been made elucidating the genetic basis for a number of these muscular dystrophies. The definition of autosomal dominant and autosomal recessive types (also referred to as LGMD type 1 and type 2, respectively, see table 1) [2, 3] demonstrated the existence of genetically distinct entities within this group.The term LGMD thus is now frequently used for a genetically as well as clinically heterogeneous group of muscular dystrophies that are characterized by progressive weakness starting in the muscles of the pelvic or shoulder girdles, largely sparing the face and the extraocular muscles [4]. With the identification of a number of the responsible genes, clinical differences between as well as variability within the different entities can now be worked out more clearly [3, 5]. In the genetically based nomenclature [2], LGMD type 1 refers to autosomal dominant conditions and LGMD type 2 to autosomal recessive entities (see table 1). The four sarcoglycan (SG)-deficient muscular dystrophies (sarcoglycanopathies, SGpathies, LGMD 2D, 2E, 2C, 2F) form a distinct subgroup within the autosomal recessive LGMD and are the subject of this review.

Dystrophin-Associated Proteins The SG are part of the larger group of dystrophin-associated proteins (DAPs). The DAPs were defined essentially by virtue of their copurification

Table 1. Gene locus-based classification of LGMD Designation

Genetic locus

Gene symbol

Protein product

Number of exons

Autosomal dominant LGMD 1A LGMD 1B LGMD 1C LGMD 1D LGMD 1E

5q31 1q11-21 3p25 6q23 7q

MYOT LMNA CAV3

myotilin lamin A/C caveolin-3 ? ?

10 12 2

Autosomal recessive LGMD 2A LGMD 2B LGMD 2C LGMD 2D LGMD 2E LGMD 2F LGMD 2G LGMD 2H LGMD 2I

15q15 2p13 13q12 17q21 4q12 5q33-34 17q11-12 9q31-33 19q13.3

CAPN3 DYSF SGCG SGCA SGCB SGCD TCAP

calpain-3 dysferlin c-SG a-SG b-SG d-SG telethonin ? ?

24 D55 8 10 6 8 2

along with dystrophin in digitonin-solublized skeletal muscle membrane preparations using wheat germ agglutinin and anion exchange (DEAE) chromatography [6–8]. Dystrophin is a large monomeric rod-like subsarcolemmal molecule that binds cytoskeletal F-actin via its N-terminus and its rod domain [reviewed in 9, 10]. The subdomains of dystrophin’s C-terminus are directly or indirectly involved in binding a number of the DAPs [reviewed in 11]. It has become clear that the DAPs can be subdivided into different subcomplexes based on biochemical characteristics, binding and association to dystrophin and the other DAPs, as well as orientation with respect to the sarcolemma (fig. 1). A purely intracellular group of DAPs consists of the syntrophins and the dystrobrevins (the dystrobrevins are DAPs that at the same time are homologous to dystrophin’s C-terminus [12]). These molecules occur in diverse isoforms and form variable complexes attached to dystrophin’s C-terminus. This subcomplex may potentially be involved in organizing ion channels and receptors within the membrane by binding cytoplasmic domains [reviewed in 13]. Another transmembrane complex besides the SG complex is dystroglycan, composed of a- and b-dystroglycan, both of which are encoded by the same gene on chromosome 3p21 and are generated by posttranslational processing of a single precursor protein [reviewed in 14]. b-Dystroglycan binds dystrophin’s

Disorders of the Sarcoglycan Complex (Sarcoglycanopathies)

27

Fig. 1. Schematic representation of the DAPs. Only the cell-binding globular domain of the laminin trimer is shown. Disorders resulting from mutations in the various components are indicated by the arrows. AR>Autosomal recessive; AD>autosomal dominant; CMD>congenital muscular dystrophy.

C-terminus at the cysteine-rich domain, spans the membrane and extracellularly binds to a-dystroglycan. One of the extracellular binding partners of a-dystroglycan is the a2 chain of the heterotrimer laminin-2, also known as merosin [reviewed in 15]. A link from dystrophin via dystroglycan to the basement membrane component laminin-2 can thus be postulated. This link may serve purely mechanical functions of force transduction but may also be involved in transmembrane signalling and basement membrane organization [reviewed in 15]. A 25-kD component that was known to copurify with the DAPs has only recently been characterized [16]. It is predicted that the protein includes four putative transmembrane domains and was found to be identical to the human KRAG gene (Kirsten ras-associated gene) on chromosome 12p11.2 [16]. Because of its structural relation to the tetraspanin family of molecules it was renamed sarcospan. Sarcospan is transcribed in many tissues, including muscle and heart. It is particularly closely associated with the SG complex [17] so that the protein is lost from the membrane together with the SG complex in patients and animal models of SGpathies.

Bo¨nnemann

28

Sarcoglycan Complex SG was established as an independent transmembrane complex amongst the DAPs by demonstrating its separation from dystrophin and the other DAPs using the detergent n-octyl-glucoside [18]. Its currently known members in skeletal muscle include a-, b-, c- and d-SG. The genes for a-, b- and c-SG were identified on the basis of partial peptide sequences [19–22], whereas the d-SG cDNA was identified as an EST on the basis of homology to c-SG [23]. It was then formally shown to be a member of the SG complex in skeletal muscle by biochemical methods [24, 25]. Another SG protein identified electronically by homology this time to a-SG is e-SG, encoded on chromosome 7q21 [26, 27]. Whereas a- and c-SG are largely restricted in their expression to striated muscle [19, 22, 28], d-SG is found in addition in smooth muscle [23]. b-SG is transcribed at highest levels in skeletal muscle, but also in smooth muscle and in a number of other tissues [20, 21]. e-SG is found rather more ubiquitously [26, 27]. Although e-SG is not highly expressed in striated muscle and does not appear to be a regular member of the SG complex in skeletal or cardiac muscle, in smooth muscle there appears to be a version of the SG complex that contains b-, d- and e-SG [29]. The four SG proteins in striated muscle are single membrane spanning molecules that are glycosylated on the extracellular side [30, 31]. a-SG, encoded on chromosome 17q21, is a type I transmembrane molecule (the protein is preceded by a signal peptide so that its C-terminus is within the cell) with a molecular weight of 50 kD [19, 32]. The other three molecules are type II transmembrane molecules (there is no preceding signal peptide, so that the N-terminus is within the cell). b-SG, encoded on 4q12, has a molecular weight of 43 kD [20, 21], c-SG (13q12) of 35 kD [22], and d-SG (5q33) also of 35 kD [23]. The extracellular domain of all SG is rich in b-pleated sheets interrupted by a-helices [30, 31]. Although there is no direct sequence homology between b-SG on the one hand and c- and d-SG on the other, in all three molecules there is a similar arrangement of four spaced cysteine residues close to the (extracellular) C-terminus resembling a partial epidermal growth factor (EGF) module [33]. The biochemical structure of the complex is beginning to be elucidated and cross-linking as well as coimmunoprecipitation experiments suggest that b- and d-SG in particular are closely associated, along with c-SG. a-SG on the other hand may be situated more separately within the complex [18, 25, 34]. However, there is close interdependency of the SGs, since the entire complex is usually severely diminished in case of mutations of only one of its members (see below). In cell culture systems, assembly of the SG complex in the sarcolemma is also dependent on the simultaneous presence of all of its members [35]. Mutations introduced in one of the SGs do not disturb translation and


29

glycosylation of the remaining SG molecules in the ER, but the complex fails to assemble in its proper position in the membrane [35]. It is still unclear how exactly the SG complex is associated with dystrophin or dystroglycan and what its function may be. There is evidence that a-dystroglycan in particular is reduced in cases of SG deficiency in animal models [36–38] as well as in patients [39], suggesting perhaps a direct or indirect link (e.g. mediated by an additional protein) between SG and dystroglycan. Other evidence indicates reciprocal influence between the SG complex and the integrin system [40]. There appears to be close physical proximity of the SGdystrophin complex with integrin a5b1 and its associated proteins [40]. Different work suggested that a-SG may function as an ectoATPase, based on the fact that it binds ATP and has ATPase activity, thus potentially regulating the activity of purinergic receptors [41]. However, a coherent picture of SG function has yet to emerge. Obviously, any theory of SG function has to explain the severe muscular dystrophy resulting from its absence as well as the substantial loss of membrane integrity that is evident from analysis of animal models of sarcoglycanopathies [37, 38, 42, 43]. In these animals there is considerable in vivo uptake of the albumin-bound tracer Evans blue into affected muscle with an age-dependent onset [38]. Recent evidence from the c-SG-deficient mouse model suggests that muscle degeneration occurs independently from mechanical stress [44], thus directing attention away from a purely mechanical function of the complex towards a a role in signalling, possibly promoting cell survival.

Sarcoglycanopathies: Clinical Features Deficiency of SG proteins was demonstrated in dystrophin-positive autosomal recessive muscular dystrophy even before the identification of the first SG mutations [45, 46]. Primary a-SGpathy was first confirmed in a large family from France with evidence of genetic linkage to the a-SG gene locus on chromosome 17 [47]. b-SGpathy was independently identified in a genetic isolate of the Indiana Amish where it coexists with LGMD 2A (calpainopathy) [21] as well as by screening sporadic cases of muscular dystrophy for b-SG mutations [20]. c-SG was first found to be mutated in consanguineous families with autosomal recessive muscular dystrophy from northern Africa [22, 48] with evidence for genetic linkage to the c-SG gene locus on chromosome 13 [49], but the disease is now recognized worldwide. d-SGpathy was discovered by independently mapping the d-SG gene and a form of autosomal recessive muscular dystrophy to the same locus on chromosome 5q33-34 [23, 50, 51].

Bo¨nnemann

30

Population studies of the SGpathies are difficult to compare and will not be listed in detail, however, certain common trends seem to emerge. The proportion of SGpathies as a group within a population of LGMD patients clearly depends on the age group analyzed as well as the severity of the phenotype. In one large series, the proportion of SGpathies among autosomal recessive and sporadic LGMD patients of all ages was 25% [52]. In the wellstudied population in Brazil the proportion of SGpathies in the entire group of LGMD patients was 20%; however, calculated for just the severe cases it was as high as 68%, whereas for the milder cases the proportion was only 8.5% [53]. Comparable numbers were found in one region in Italy, where the proportion of SGpathy among severe childhood LGMD presentations was 54.5% as opposed to 17.5% for the milder group [54]. Similar relationships in principle were also found in the Turkish childhood LGMD population [55] and in an American series [56]. Comparing the relative proportion of the different SG gene mutations in various populations in which there is no strong founder effect for one single mutation, a-SG seems to be the most common SG gene mutated (40–50% of SG mutations), followed by both b- and c-SG [52, 54, 56]. Although d-SGpathy seems to be more common in Brazil [53], it is probably the least common form in other populations [57]. From the analysis of patients with SGpathies identified so far, no discriminating clinical differences between the four disorders have emerged as of yet. Therefore, their clinical features will be discussed jointly, emphasizing possible differences. Manifestation in childhood is the most common presentation with an age at onset of around 6–8 years on the average, but weakness may start earlier or considerably later, including adulthood [3]. Early motor milestones tend to be normal in the majority of cases but may be delayed with abnormal ambulation or toe walking from the beginning. The presenting complaints/ features in addition include waddling gait, difficulties with running or getting up from the floor, but also excercise intolerance, potentially muscle cramps or pain (‘pseudometabolic’ presentation [58]). The initial muscle weakness usually presents in the pelvic girdle followed by the muscles of the shoulder girdle. The clinical pattern of muscle involvement is reminiscent of Duchenne and Becker muscular dystrophies (DMD/BMD) with early involvement of glutei and hip adductors [58, 59], although involvement of the scapular fixators and the deltoid muscle may be more significant in the SGpathies compared to the dystrophinopathies, leading to early scapular winging and loss of arm elevation. Quadriceps and hamstrings may be equally involved. There may also be early Achilles’ tendon contractures and lordosis. Calf hypertrophy usually is present at some point, hypertrophy of other muscles as well as sometimes macroglossia can occur. The weakness is progressive and often rapidly so. In childhood onset cases there may be loss of ambulation at around


31

12–16 years of age, although wheelchair confinement before the age of 10 has been seen [51, 58, 60]. While later onset and milder progression are also possible in b- [21, 39, 52] and c-SGpathy [61, 62], the highest proportion of milder cases is found in of a-SGpathy [56, 58, 59, 63], including a case of an almost asymptomatic adult with high serum creatine kinase (CK) levels and some lordosis only [64]. There have been very few cases of genetically confirmed d-SGpathy so far; the ones reported have been severe [51, 65]. However, for a-SGpathy [58, 64], for b-SGpathy [21] as well as in particular for c-SGpathy [48, 61], there can be significant intra- and interfamilial variability that makes predictions of prognosis in an individual case difficult, even if there had been an affected sib in the same family. CK levels are significantly elevated, especially early in the course. A significant difference to the dystrophinopathy spectrum is the lack of intellectual involvement in the SGpathies. Although in the majority of cases there is no overt cardiomyopathy, a smaller number of patients may present with clinically significant dilative cardiomyopathy, sometimes even requiring cardiac transplantation [66]. Evidence for subclinical cardiac involvement by ECG and ECHO criteria may be seen more often [67–69] but less prominently so than in DMD/BMD. Significant cardiomyopathy can definitely occur in b-SGpathy [70]. Although reported patients with confirmed d-SGpathy have not had cardiomyopathy, there are patients with very probable d-SGpathy and significant cardiomyopathy [71; Voit, pers. commun.]. There have been no reported cases of a- or c-SGpathy with clinically overt cardiomyopathy so far, but the number may still be too small to reach definite conclusions (see discussion under Animal Models). In the later course of SGpathy there is also the possibility of progressive respiratory compromise concomitant with the development of progressive scoliosis.

Diagnosis, Protein Expression Patterns and Mutational Spectrum The diagnosis of a SGpathy rests on history and clinical examination, analysis of protein expression patterns in the muscle biopsy and ultimately on demonstrating a causative gene mutation in one of the SG genes. In informative families, genetic linkage analysis may help to include, and perhaps more importantly, to exclude known genetic loci from diagnostic considerations. More frequently encountered though is the sporadic patient with a proximal muscular dystrophy of unknown cause. Analysis of the muscle biopsy will frequently be the next step in the evaluation of such a patient, in particular when there is no genetic evidence for a dystrophinopathy based on the PCR deletion screening [72]. A characteristic feature of the disorders of the SG complex is the disintegration of the entire complex when mutations in one of the four

Bo¨nnemann

32

proteins occur [30, 31]. Thus, the primary mutated gene may not be obvious from analyzing the biopsy by immunohistochemistry or by Western blot. However, on utilizing all four SG antibodies certain helpful protein deficiency patterns may become evident. In a-SGpathy and c-SGpathy the protein most reduced or absent is the mutated one, while the other components may show significant preservation at the membrane, in particular d-SG in cases of c-SGpathy [53, 55, 73, 74]. On the other hand, in b- and also d-SGpathy in most cases there is complete absence of the entire complex or at least considerable reduction of all SG proteins [20, 21, 39, 53, 60, 73], although in d-SGpathy there may be some partial preservation of c-SG [75]. Only in a-SGpathy is there some correlation between the residual immunoreactivity for the mutated protein and the clinical severity [58, 59]; the same correlation does not hold true for the other SG. In c-SGpathy for instance, complete absence of c-SG immunoreactivity does not automatically imply a severe phenotype as this can be seen in severe as well as in milder cases [61]. In some patients there may be some decrease in dystrophin immunoreactivity as well, sometimes presenting with an immunohistochemical picture reminiscent of BMD [67, 73, 75]. Nonetheless, according to Western blot analysis the dystrophin molecule is of normal molecular weight, and the residual amount of dystrophin in a SGpathy is likely to be more than 20–30%, although it may be as low as 10% of normal [75]. Vice versa, there can also be a substantial reduction of the SG complex along with dystroglycan and the other DAPs and sarcospan in the dystrophinopathies [76]. It has been speculated that this loss of DAPs is in fact responsible for the development of muscular dystrophy [77, 78]. Mutations: a-SG Mutational analysis in the a-SG gene has revealed a predominance of missense mutations over truncating mutations [54, 79, 80]. Genotype/phenotype correlations are quite difficult to establish, in particular because of the high number of compound heterozygotes [80]. However, null mutations in a-SG consistently appear to lead to a severe phenotype, whereas there is much more clincical variability associated with the missense mutations [58, 80]. The misssense mutations described so far are located exclusively in the extracellular domain. One third of mutated chromosomes in the largest series from Europe carried the single most prevalent missense mutation, Arg77Cys [80]. This mutation in a CpG dinucleotide occurs on different genetic backgrounds and must, therefore, have arisen indepedently a number of times. The phenotype associated with this mutation again is quite variable, ranging from intermediate to mild in one study, the severity being correlated with the residual a-SG staining in the biopsy [58, 59]. Another relatively frequent mutation in the


33

large European series was Arg284Cys (12% of mutated chromosomes), which in the homozygous state was always associated with a mild clinical presentation [59, 80]. This mild phenotype probably is the only consistent genotype/ phenotype correlation for a-SG missense mutations. In this largest study so far, there was only one clinically severe homozygous a-SG missense mutation, Arg34Cys [80]. Mutations: b-SG In b-SG again there is a mixture of missense as well as truncating mutations. All the truncating mutations were associated with a severe phenotype [56, 59, 60]. In contrast to a-SG, in b-SG there is a relatively high proportion of missense mutations associated with a phenotype as severe as found for the truncating mutations [60]. Thus, overall the proportion of severe cases associated with missense mutations is considerably higher in b-SGpathy than in a-SGpathy. Most b-SG mutations are in the extracellular domain with the notable exception of Gln11Glu, located at the very N-terminus [56]. There is a certain cluster of b-SG mutations in a region immediately adjacent to the transmembrane domain, encoded on exon 3, most, but not all of which [39] are associated with a severe phenotype [60, 81]. The missense mutations that are associated with comparatively milder phenotypes and much more variability of presentation and progression tend to be located further away from that region [21, 52]. This data may imply, but does not prove, that the region immediately adjacent to the transmembrane domain in b-SG could be crucial for b-SG function within the complex, maybe mediating some of the interactions within the complex. Thr151Arg is the mutation that was identified in the original Indiana Amish pedgrees [21], although there appears to be a second mild b-SG mutation segregating within that population (Arg91Cys) [39]. Mutations: c-SG In c-SG most of the mutations identified so far have been truncating mutations, frequently caused by deletions [22, 33, 55, 56, 59, 61]. The deletions may also be larger, potentially encompassing most of the gene locus [97, Bo¨nnemann and Kunkel, unpubl. obs.]. Two recurrent mutations in the c-SG gene are important: A deletion of a single T residue within exon 6 (del521T), causing a frameshift and truncation of the protein, was originally described in the North African population [22] and subsequently in a number of patients originating from the Mediterranean region [61]. The mutation is associated with a rare 122-bp allele of the intragenic dinucleotide repeat marker (D13S232) suggesting a founder effect of the mutation [82]. Although most patients with this mutation are rather severly affected, some show much milder disease [61].

Bo¨nnemann

34

Since this is a truncating mutation, for c-SG the dictum that null mutations always are associated with a severe phenotype does not appear to be true. Thus, there probably are modifiers of severity that still remain to be identified. Another recurrent and relevant mutation is the missense mutation Cys283Tyr, found almost exclusively among Romano Gypsy patients, based on a founder effect [83, 84]. The missense mutation alters one of the terminal cysteine residues that form a partial EGF module. The consistently severe phenotype associated with this mutation [83] probably reflects the functional importance of the motif that could be involved in protein-protein interactions, although a binding partner has not been identified yet. Other missense mutations in the c-SG gene have been rare, and only one was in a homozygous state that would allow for genotype/phenotype correlations (Leu193Ser), in this case causing a milder phenotype [62]. Mutations: d-SG In d-SG only very few mutations have been identified so far, all of them causing a severe phenotype of early childhood onset and rapid progression to wheelchair confinement [51, 57]. This includes the only missense mutation known in the gene (Glu262Lys) [65]. In this tendency towards a more severe phenotype, d-SGpathy may be similar to b-SGpathy, even though the protein structure as well as the gene structure resemble more c-SG.

Animal Models The first known animal model of an SGpathy is the cardiomyopathic Syrian hamster which also has muscular dystrophy [85]. Originating from the original strain BIO 1.50, animals with predominantly hypertrophic (BIO 14.6 and UMX 7.1) as well as those with predominantly dilated cardiomyopathy (TO-2) have been bred [86]. The animals were found to lack expression of the entire SG complex [36, 87]. Subsequently a large 29.8-kb deletion in the 5 region upstream of exon 2 in the d-SG gene was found to cause the disease in all the different strains [88–90]. Since the deletion was identical in the different strains with dilated versus hypertrophic cardiomyopathy, the genetic background plays a role in determining the exact phenotype of the cardiomyopathy. In addition to this spontaneous animal model, mice strains with null mutations in each of four SGs have now been generated by gene targeting techniques [37, 42, 43, 91]. All mice develop significant muscular dystrophy that becomes evident histologically at an early age. Whereas the a-SG mice were clinically quite normal [37], the c-SG-deficient mice had clinically evident


35

muscular dystrophy with abnormal gait, motor difficulties and stunted growth [42]. From the reports published it is not clear how severe the muscular dystrophy was clinically in the b- and d-SG-deficient animals [43, 91]. All mice showed active necrosis and regeneration of skeletal muscle, as well as uptake of the dye Evans blue into muscle, indicating sarcolemmal disruption [37, 38, 42, 43, 91]. In addition, apoptotic cell death was observed in the c-SG-deficient animals [42]. In each of the animals there was evidence of disruption of the SG complex with severe reduction of the other three SGs. In the null mutants in which sarcospan expression was analyzed [a-, b-, d-SG-deficient mice; 37, 43, 91], there was concomitant severe reduction of this molecule as well, confirming its close association with the SG complex. a-Dystroglycan was analyzed in the a-SG- and the b-SG-deficient mice and appeared to be more loosely associated with the sarcolemma in membrane preparations from muscle lysates [37, 43]. These results indicate that the close membrane association of a-dystroglycan may at least in part be dependent on the SG complex. Similar to the d-SG-deficient hamster, the d-SG-deficient mice also develop significant cardiomyopathy [91]. Loss of d-SG-disrupts the smooth muscle SG complex composed of at least b-SG, d-SG and e-SG [29]. In the d-SG deficient mice, the lumen of the coronary arteries was irregular with a cardiac pathology that suggested ischemic lesions. The authors in that study thus propose that the cardiomyopathy develops on the basis of vascular dysfunction caused by the loss of the smooth muscle SG complex [91]. Part of the resulting cardiac pathology in mice after exercise could be reversed by pretreating the mice with the vasodilator nicorandil, opening important therapeutic considerations in these muscular dystrophies. In view of the vascular hypothesis of cardiomyopathy it is interesting to note that a reduced coronary reserve was reported in patients with SGpathy [69] and that the likelihood of significant cardiomyopathy is probably highest in patients with either b- or d-SGpathy. The b-SGdeficient mice should also have lost the smooth muscle SG complex although this was not directly shown. With increasing age these mice show larger fibrotic patches in the heart, even though there does not appear to be a clinical correlate such as increased mortality [43]. The a-SG-deficient mice, on the other hand, do not develop cardiomyopathy, even though the entire SG complex is significantly reduced in cardiac muscle [37]. Since a-SG is not part of the smooth muscle SG complex this finding would be consistent with a vascular pathogenesis of SG-deficient cardiomyopathy. However, one strain of c-SG-deficient mouse also develops significant hypertrophic cardiomyopathy of the left as well as the right ventricle contributing to significant early mortality in these animals [42], even though c-SG is also not expressed in smooth muscle [29]. Thus, the pathophysiology of cardiomyopathy in SGpathies may be multifactorial and heterogeneous.

Bo¨nnemann

36

Treatment There are a few observations in the literature suggesting that in the SGpathies there may be a response to corticosteroid treatment that is reminiscent of the response seen in DMD with a sustained benefit over a number of years [64, 92, 93]. However, there are no controlled studies yet and no data on long-term outcome. In any case, the similarity of the response is intriguing also from a physiological point of view, since dystrophinopathies and SGpathies may have certain aspects of pathophysiology in common [78]. Gene transfer has been accomplished in the SG-deficient cardiomyopathic hamster [94–96] as well as in the a-SG-deficient mouse [37], facilitated by the small size of SG cDNAs. In all attempts, the introduction of the cDNA for the mutated SG resulted in a restoration of the entire SG complex and led to a restoration of sarcolemmal integrity, as assayed by Evans blue [94] or Procion orange dye assays [95]. Even marked overexpression and intracellular accumulation of the excess-induced protein did not influence this result at the membrane [95, 96]. The transfer was performed by intramuscular recombinant adenovirus injection [37, 94], or using recombinant adenovirus-associated virus [96], in one study as an intra-arterial injection after pretreatment with papaverine and histamine to open the endothelial barrier of the vessels [95]. From these studies on SG gene transfer, there is good reason to surmise that successful gene transfer will result in the functional recovery of the treated muscle. However, while these results are encouraging with respect to human gene therapy in the SGpathies, the formidable challenge of safe delivery to all skeletal muscles (and potentially the heart including coronary arteries), long-term expression of the introduced gene, the degree of fibrosis, and immunotolerance remain to be addressed as prerequisites for such a treatment to become a success. Meanwhile, we currently have the means for a proper and precise genetic diagnosis of these disorders, certainly a sine qua non for any gene-based therapeutic approach to the patient with a SG-deficient muscular dystrophy.

Note Added in Proof A muscle-specific protein, filamin 2, has recently been identified as an intracellular interactor of the SG complex [98]. Another b-SG knock-out mouse model notably showed similar vascular anomalies as described for the d-SG knock-out mice before [99]. This and another paper also show that there appears to be a separate SG complex in striated muscle in which e-SG replaces a-SG [100].


37

Acknowledgements I would like to express my gratitude to Dr. Eijiro Ozawa, Tokyo, for his encouragement to write this review and to Dr. Volker Straub, Essen, for critically reviewing the text and for helpful discussions. The support of the Deutsche Gesellschaft fu¨r Muskelkranke e.V. (DGM) is gratefully acknowledged.

References 1 2 3

4 5 6 7 8 9 10 11 12

13 14 15 16 17

18

19

20

van der Kooi AJ, de Visser M, Barth PG: Limb girdle muscular dystrophy: Reappraisal of a rejected entity. Clin Neurol Neurosurg 1994;96:209–218. Bushby KM, Beckmann JS: The limb girdle muscular dystrophies – Proposal for a new nomenclature. Neuromuscul Disord 1995;4:337–343. Beckmann JS, Brown RH, Muntoni F, Urtizberea A, Bo¨nnemann CG, Bushby KMD: 66th/67th ENMC sponsored international workshop – The limb-girdle muscular dystrophies. Neuromuscul Disord 1999;9:436–445. Bushby KMD: Diagnostic criteria for the limb-girdle muscular dystrophies: Report of the ENMC Consortium on Limb-Girdle Dystrophies. Neuromuscul Disord 1995;5:71–74. Bushby KM: Making sense of the limb-girdle muscular dystrophies. Brain 1999;122:1403–1420. Campbell KP, Kahl SD: Association of dystrophin and an integral membrane glycoprotein. Nature 1989;338:259–262. Yoshida M, Ozawa E: Glycoprotein complex anchoring dystrophin to sarcolemma. J Biochem (Tokyo) 1990;108:748–752. Ervasti JM, Campbell KP: Membrane organization of the dystrophin-glycoprotein complex. Cell 1991;66:1121–1131. Ahn AH, Kunkel LM: The structural and functional diversity of dystrophin. Nat Genet 1993;3:283–291. Rybakova IN, Amann KJ, Ervasti JM: A new model for the interaction of dystrophin with F-actin. J Cell Biol 1996;135:661–672. Sadoulet-Puccio HM, Kunkel LM: Dystrophin and its isoforms. Brain Pathol 1996;6:25–35. Sadoulet-Puccio HM, Khurana TS, Cohen JB, Kunkel LM: Cloning and characterization of the human homologue of a dystrophin related phosphoprotein found at the Torpedo electric organ post-synaptic membrane. Hum Mol Genet 1996;5:489–496. Froehner SC, Adams ME, Peters MF, Gee SH: Syntrophins: Modular adapter proteins at the neuromuscular junction and the sarcolemma. Soc Gen Physiol Ser 1997;52:197–207. Tinsley JM, Blake DJ, Zuellig RA, Davies KE: Increasing complexity of the dystrophin-associated protein complex. Proc Natl Acad Sci USA 1994;91:8307–8313. Durbeej M, Henry MD, Campbell KP: Dystroglycan in development and disease. Curr Opin Cell Biol 1998;10:594–601. Crosbie RH, Heighway J, Venzke DP, Lee JC, Campbell KP: Sarcospan, the 25-kDa transmembrane component of the dystrophin-glycoprotein complex. J Biol Chem 1997;272:31221–31224. Crosbie RH, Lebakken CS, Holt KH, Venzke DP, Straub V, Lee JC, Grady RM, Chamberlain JS, Sanes JR, Campbell KP: Membrane targeting and stabilization of sarcospan is mediated by the sarcoglycan subcomplex. J Cell Biol 1999;145:153–165. Yoshida M, Suzuki A, Yamamoto H, Noguchi S, Mizuno Y, Ozawa E: Dissociation of the complex of dystrophin and its associated proteins into several unique groups by n-octyl beta-D-glucoside. Eur J Biochem 1994;222:1055–1061. Roberds SL, Anderson RD, Ibraghimov-Beskrovnaja O, Campbell KP: Primary structure and expression of the 50-kDa dystrophin-associated glycoprotein (adhalin). J Biol Chem 1993;268: 23739–23742. Bo¨nnemann CG, Modi R, Noguchi S, Mizuno Y, Yoshida M, Gussoni E, McNally EM, Duggan DJ, Angelini C, Hoffman EP, Ozawa E, Kunkel L M: b-Sarcoglycan (A3b) mutations cause autosomal recessive muscular dystrophy with loss of the sarcoglycan complex. Nat Genet 1995;11:266–273.

Bo¨nnemann

38

21

22

23

24

25 26 27 28

29

30 31 32 33

34 35 36

37

38

39

Lim LE, Duclos F, Broux O, Bourg N, Sunada Y, Allamand V, Meyer J, Richard I, Tome´ F, Fardeau M, Moomaw C, Slaughter C, Jackson CE, Beckmann JS, Campbell KP: b-Sarcoglycan (43 DAG): Characterization and involvement in a recessive form of limb-girdle muscular dystrophy linked to chromosome 4q12. Nat Genet 1995;11:257–265. Noguchi S, McNally EM, Ben Othmane K, Hagiwara Y, Mizuno Y, Yoshida M, Yamamoto H, Bo¨nnemann CG, Gussoni E, Denton PH, Kyriakides T, Middleton L, Hentati F, Ben Hamida M, Nonaka I, Vance JM, Kunkel LM, Ozawa E: Mutations in the dystrophin-associated protein c-sarcoglycan in chromosome 13 muscular dystrophy. Science 1995;270:819–822. Nigro V, Piluso G, Belsito A, Politano L, Puca AA, Papparella S, Rossi E, Viglietto G, Eposito MG, Abbondanza C, Medici N, Molinari AM, Nigro G, Puca GA: Identification of a novel sarcoglycan gene at 5q33 encoding a sarcolemmal 35 kDa glycoprotein. Hum Mol Genet 1996;5:1179–1186. Jung D, Duclos F, Apostol B, Straub V, Lee JC, Allamand V, Venzke DP, Sunada Y, Moomaw CR, Leveille CJ, Slaughter CA, Crawford TO, McPherson J, Campbell K: Characterization of d-sarcoglycan, a novel component of the oligomeric sarcoglycan complex involved in LGMD. J Biol Chem 1996;271:32321–32329. Yoshida M, Noguchi S, Wakabayashi E, Piluso G, Belsito A, Nigro V, Ozawa E: The fourth component of the sarcoglycan complex. FEBS Lett 1997;403:143–148. Ettinger AJ, Feng G, Sanes JR: Epsilon-sarcoglycan, a broadly expressed homologue of the gene mutated in limb-girdle muscular dystrophy 2D. J Biol Chem 1997;272:32534–32538. McNally EM, Ly CT, Kunkel LM: Human e-sarcoglycan is highly related to a-sarcoglycan (adhalin), the limb girdle muscular dystrophy 2D gene. FEBS Lett 1998;422:27–32. Mizuno Y, Yoshida M, Yamamoto H, Hirai S, Ozawa E: Distribution of dystrophin isoforms and dystrophin-associated 43 DAG (A3a) and 50 DAG (A2) in various monkey tissues. J Biochem (Tokyo) 1993;114:936–941. Straub V, Ettinger AJ, Durbeej M, Venzke DP, Cutshall S, Sanes JR, Campbell KP: Epsilonsarcoglycan replaces alpha-sarcoglycan in smooth muscle to form a unique dystrophin-glycoprotein complex. J Biol Chem 1999;274:27989–27996. Ozawa E, Noguchi S, Mizuno Y, Hagiwara Y, Yoshida M: From dystrophinopathy to sarcoglycanopathy: Evolution of a concept of muscular dystrophy. Muscle Nerve 1998;21:421–438. Lim LE, Campbell KP: The sarcoglycan complex in limb-girdle muscular dystrophy. Curr Opin Neurol 1998;11:443–452. McNally EM, Yoshida M, Mizuno Y, Ozawa E, Kunkel LM: Human adhalin is alternatively spliced and the gene is located on chromosome 17q21. Proc Natl Acad Sci USA 1994;91:9690–9694. McNally EM, Duggan DJ, Gorospe JR, Bo¨nnemann CG, Fanin M, Lidov HGW, Noguchi S, Ozawa E, Rulye SZ, Cruse RP, Angelini C, Kunkel LM, Hoffman EP: Mutations in the carboxyl-terminus of c-sarcoglycan cause muscular dystrophy. Hum Mol Genet 1996;5:1841–1847. Chan Y, Bo¨nnemann CG, Lidov HGW, Kunkel LM: Molecular organization of sarcoglycan complex in mouse myotubes in culture. J Cell Biol 1998;143:2033–2044. Holt KH, Campbell KP: Assembly of the sarcoglycan complex. Insights for muscular dystrophy. J Biol Chem 1998;273:34667–34670. Roberds SL, Ervasti JM, Anderson RD, Ohlendieck K, Kahl SD, Zoloto D, Campbell KP: Disruption of the dystrophin-glycoprotein complex in the cardiomyopathic hamster. J Biol Chem 1993; 268:11469–11499. Duclos F, Straub V, Moore SA, Venzke DP, Hrstka RF, Crosbie RH, Durbeej M, Lebakken CS, Ettinger AJ, van der Meulen J, Holt KH, Lim LE, Sanes JR, Davidson BL, Faulkner JA, Williamson R, Campbell KP: Progressive muscular dystrophy in alpha-sarcoglycan-deficient mice. J Cell Biol 1998;142:1461–1471. Straub V, Duclos F, Venzke DP, Lee JC, Cutshall S, Leveille CJ, Campbell KP: Molecular pathogenesis of muscle degeneration in the delta-sarcoglycan-deficient hamster. Am J Pathol 1998;153: 1623–1630. Duclos F, Broux O, Bourg N, Straub V, Feldman GL, Sunada Y, Lim LE, Piccolo F, Cutshall S, Gary F, Quetier F, Kaplan JC, Jackson CE, Beckmann JS, Campbell KP: Beta-sarcoglycan: Genomic analysis and identification of a novel missense mutation in the LGMD2E Amish isolate. Neuromuscul Disord 1998;8:30–38.


39

40 41 42

43

44 45

46

47

48 49

50

51

52

53

54

55

56 57

Yoshida T, Pan Y, Hanada H, Iwata Y, Shigekawa M: Bidirectional signaling between sarcoglycans and the integrin adhesion system in cultured L6 myocytes. J Biol Chem 1998;273:1583–1590. Betto R, Senter L, Ceoldo S, Tarricone E, Biral D, Salviati G: Ecto-ATPase activity of alphasarcoglycan (adhalin). J Biol Chem 1999;274:7907–7912. Hack AA, Ly CT, Jiang F, Clendenin CJ, Sigrist KS, Wollmann RL, McNally EM: c-Sarcoglycan deficiency leads to muscle membrane defects and apoptosis independent of dystrophin. J Cell Biol 1998;142:1279–1287. Araishi K, Sasaoka T, Imamura M, Noguchi S, Hama H, Wakabayashi E, Yoshida M, Hori T, Ozawa E: Loss of the sarcoglycan complex and sarcospan leads to muscular dystrophy in bsarcoglycan-deficient mice. Hum Mol Genet 1999;8:1589–1598. Hack AA, Cordier L, Shoturma DI, Lam MY, Sweeney HL, McNally EM: Muscle degeneration without mechanical injury in sarcoglycan deficiency. Proc Natl Acad Sci USA 1999;96:10723–10728. Matsumura K, Tome FMS, Huguette C, Azibi K, Chaouch M, Kaplan J-C, Fardeau M, Campbell KP: Deficiency of the 50K dystrophin-associated glycoprotein in severe childhood autosomal recessive muscular dystrophy. Nature 1992;359:320–322. Mizuno Y, Noguchi S, Yamamoto H, Yoshida M, Suzuki A, Hagiwara Y, Hayashi YK, Arahata K, Nonaka I, Hirai S, Ozawa E: Selective defect of sarcoglycan complex in severe childhood autosomal recessive muscular dystrophy muscle. Biochem Biophys Res Commun 1994;203: 979–983. Roberds SL, Leturcq F, Allamand V, Piccolo F, Jeanpierre M, Anderson RD, Lim LE, Lee JC, Tome´ FMS, Romero NB, Fardeau M, Beckmann JS, Kaplan J-C, Campbell KP: Missense mutations in the adhalin gene linked to autosomal recessive muscular dystrophy. Cell 1994;78:625–633. Ben Hamida M, Fardeau M, Attia N: Severe childhood muscular dystrophy affecting both sexes and frequent in Tunisia. Muscle Nerve 1983;6:469–480. Ben Othmane K, Ben Hamida M, Pericak-Vance MA, Ben Hamida C, Blel S, Carter SC, Bowcock AM, Petruhkin K, Gilliam T C, Roses AD, Hentati F, Vance JM: Linkage of Tunisian autosomal recessive Duchenne-like muscular dystrophy to the pericentromeric region of chromosome 13q. Nat Genet 1992;2:315–317. Passos-Bueno MR, Moreira ES, Vainzof M, Marie SK, Zatz M: Linkage analysis in autosomal recessive limb-girdle muscular dystrophy (AR LGMD) maps a sixth form to 5q33–34 (LGMD2F) and indicates that there is at least one more subtype of AR LGMD. Hum Mol Genet 1996;5: 815–820. Nigro V, de Sa´ Moriera E, Piluso G, Vainzof M, Belsito A, Politano L, Puca AA, Passos-Bueno MR, Zatz M: Autosomal recessive limb-girdle muscular dystrophy (LGMD 2F) is caused by a mutation in the d-sarcoglycan gene. Nat Genet 1996;14:195–198. Ginjaar HB, van der Kooi A, Ceelie H, Kneppers ALJ, Barth PG, Busch HFM, Wokke JHJ, Kerkhoff H, Broere D, Anderson LVB, Bo¨nnemann CG, Jeanpierre M, Bakker E, de Visser M, van Ommen GJB: Sarcoglycanopathies in Dutch patients with autosomal recessive limb girdle muscular dystrophies. Neuromuscul Disord 1997;7:440. Vainzof M, Passos-Bueno MR, Pavanello RC, Marie SK, Oliveira AS, Zatz M: Sarcoglycanopathies are responsible for 68% of severe autosomal recessive limb-girdle muscular dystrophy in the Brazilian population. J Neurol Sci 1999;164:44–49. Fanin M, Duggan DJ, Mostacciuolo ML, Martinello F, Freda MP, Soraru G, Trevisan CP, Hoffman EP, Angelini C: Genetic epidemiology of muscular dystrophies resulting from sarcoglycan gene mutations. J Med Genet 1997;34:973–977. Dincer P, Leturcq F, Richard I, Piccolo F, Yalnizoglu D, de Toma C, Akcoren Z, Broux O, Deburgrave N, Brenguier L, Roudaut C, Urtizberea JA, Jung D, Tan E, Jeanpierre M, Campbell KP, Kaplan JC, Beckmann JS, Topaloglu H: A biochemical, genetic, clinical survey of autosomal recessive limb girdle muscular dystrophies in Turkey. Ann Neurol 1997;42:222–229. Duggan DJ, Gorospe JR, Fanin M, Hoffman EP, Angelini C: Mutations in the sarcoglycan genes in patients with myopathy. N Engl J Med 1997;336:618–624. Duggan DJ, Manchester D, Stears KP, Mathews DJ, Hart C, Hoffman EP: Mutations in the dsarcoglycan gene are a rare cause of autosomal recessive limb-girdle muscular dystrophy (LGMD2). Neurogenetics 1997;1:49–58.

Bo¨nnemann

40

58

59 60

61

62

63

64

65

66

67

68

69

70

71

72 73

74

75

Eymard B, Romero NB, Leturcq F, Piccolo F, Carrie A, Jeanpierre M, Collin H, Deburgrave N, Azibi K, Chaouch M, Merlini L, Themar-Noel C, Penisson I, Mayer M, Tanguy O, Campbell KP, Kaplan JC, Tome FM, Fardeau M: Primary adhalinopathy (alpha-sarcoglycanopathy): Clinical, pathologic, genetic correlation in 20 patients with autosomal recessive muscular dystrophy. Neurology 1997;48:1227–1234. Angelini C, Fanin M, Freda MP, Duggan DJ, Siciliano G, Hoffman EP: The clinical spectrum of sarcoglycanopathies. Neurology 1999;52:176–179. Bo¨nnemann CG, Passos-Bueno R, McNally EM, Vainzoff M, de Sa´ Moreira E, Marie SK, Pavanello RCM, Noguchi S, Ozawa E, Zatz M, Kunkel LM: Genomic screening for b-sarcoglycan mutations: Missense mutations may cause severe limb-girdle muscular dystrophy type 2E (LGMD 2E). Hum Mol Genet 1996;5:1953–1961. McNally EM, Passos-Bueno R, Bo¨nnemann CG, Vainzoff M, De Sa´ Moreira E, Lidov HGW, Ben Othmane K, Denton PH, Vance JM, Zatz M, Kunkel LM: Mild and severe muscular dystrophy caused by a single c-sarcoglycan mutation. Am J Hum Genet 1996;59:1040–1047. van der Kooi AJ, de Visser M, van Meegen M, Ginjaar HB, van Essen AJ, Jennekens FGI, Jongen PJH, Leschot NJ, Bolhuis PA: A novel c-sarcoglycan mutation causing childhood onset, slowly progressive limb girdle muscular dystrophy. Neuromuscul Disord 1998;8:305–308. Passos-Bueno MR, Vainzof M, Moreira ES, Zatz M: Seven autosomal recessive limb-girdle muscular dystrophies in the Brazilian population: From LGMD2A to LGMD2G. Am J Med Genet 1999; 82:392–398. Angelini C, Fanin M, Menegazzo E, Freda MP, Duggan DJ, Hoffman EP: Homozygous a-sarcoglycan mutation in two siblings: One asymptomatic and one steroid responsive mild limb-girdle muscular dystrophy paitent. Muscle Nerve 1998;21:769–775. Moreira ES, Vainzof M, Marie SK, Nigro V, Zatz M, Passos-Bueno MR: A first missense mutation in the delta sarcoglycan gene associated with a severe phenotype and frequency of limb-girdle muscular dystrophy type 2F (LGMD2F) in Brazilian sarcoglycanopathies. J Med Genet 1998;35:951–953. Fadic R, Sunada Y, Waclawik AJ, Buck S, Lewandoski PJ, Campbell KP, Lotz BP: Brief report: Deficiency of a dystrophin-associated glycoprotein (adhalin) in a patient with muscular dystrophy and cardiomyopathy. N Engl J Med 1996;334:362–366. Prelle A, Comi GP, Tancredi L, Rigoletto C, Ciscato P, Fortunato F, Nesti S, Sciacco M, Robotti M, Bazzi P, Felisari G, Moggio M, Scarlato G: Sarcoglycan deficiency in a large Italian population of myopathic patients. Acta Neuropathol (Berl) 1998;96:509–514. Melacini P, Fanin M, Duggan DJ, Freda MP, Berardinelli A, Danieli GA, Barchitta A, Hoffman EP, Dalla Volta S, Angelini C: Heart involvement in muscular dystrophies due to sarcoglycan gene mutations. Muscle Nerve 1999;22:473–479. Gnecchi-Rusconi T, Taylor J, Mercuri E, Paternostro G, Pogue R, Bushby K, Sewry C, Muntoni F, Camici PG: Cardiomyopathy in Duchenne, Becker and sarcoglycanopathies: A role for coronary dysfunction? Muscle Nerve 1999;22:1549–1556. Barresi R, Di Blasi C, Negri T, Brugnoni R, Vitali A, Felisari G, Salandi A, Daniel S, Cornelio F, Morandi L, Mora M: Disruption of heart sarcoglycan complex and severe cardiomyopathy caused by beta-sarcoglycan mutations. J Med Genet 2000;37:102–107. Jungbluth H, Dunckley M, Wilson L, Sewry C, Manzur AY, Mercuri E, Nigro V, Dubowitz V, Muntoni F: Becker or limb girdle muscular dystrophy? An intriguing case. Dev Med Child Neurol 1998;40(suppl 79):20–21. Bushby KMD: The limb-girdle muscular dystrophies: Diagnostic guidelines. Eur J Paediatr Neurol 1999;3:53–58. Vainzof M, Passos-Bueno MR, Moreira ES, Pavanello RCM, Marie SK, Anderson LVB, Bo¨nnemann CG, McNally EM, Nigro V, Kunkel LM, Zatz M: The sarcoglycan complex in the six autosomal recessive limb-girdle muscular dystrophies. Hum Mol Genet 1996;5:1963–1969. Higuchi I, Kawai H, Umaki Y, Kawajiri M, Adachi K, Fukunaga H, Nakagawa M, Arimura K, Osame M: Different manners of sarcoglycan expression in genetically proven a-sarcoglycan and c-sarcoglycan deficiency. Acta Neuropathol (Berl) 1998;96:202–206. Vainzof M, Moreira ES, Ferraz G, Passos-Bueno MR, Marie SK, Zatz M: Further evidence for the organisation of the four sarcoglycan proteins within the dystrophin-glycoprotein complex. Eur J Hum Genet 1999;7:251–254.


41

76

77

78 79

80

81

82

83

84

85 86 87 88

89

90 91

92

93

Ohlendieck K, Matsumuru K, Ionasescu VV, Towbin JA, Bosch EP, Weinstein SL, Sernett BS, Campbell KP: Duchenne muscular dystrophy: Deficiency of dystrophin-associated proteins in the sarcolemma. Neurology 1993;43:795–800. Matsumura K, Campball KP: Deficiency of dystrophin-associated proteins: A common mechanism leading to muscle cell necrosis in severe childhood muscular dystrophies. Neuromuscul Disord 1993; 3:109–118. Campbell KP: Three muscular dystrophies: Loss of cytoskeleton-extracellular matrix linkage. Cell 1995;80:675–679. Piccolo F, Roberds SL, Jeanpierre M, Leturcq F, Azibi K, Beldjord C, Carrie A, Recan D, Chaouch M, Reghis A, El Kerch F, Sefiani A, Voit T, Merlini L, Collin H, Eymard B, Beckmann JS, Romero NB, Tome´ FMS, Fardeau M, Campbell KP, Kaplan J-C: Primary adhalinopathy: A common cause of autosomal recessive muscular dystrophy of variable severity. Nat Genet 1995; 10:243–245. Carrie A, Piccolo F, Leturcq F, de Toma C, Azibi K, Beldjord C, Vallat JM, Merlini L, Voit T, Sewry C, Urtizberea JA, Romero N, Tome FM, Fardeau M, Sunada Y, Campbell KP, Kaplan JC, Jeanpierre M: Mutational diversity and hot spots in the alpha-sarcoglycan gene in autosomal recessive muscular dystrophy (LGMD2D). J Med Genet 1997;34:470–475. Bo¨nnemann CG, Wong J, Ben Hamida C, Ben Hamida M, Hentati F, Kunkel LM: LGMD 2 E in Tunisia is caused by a missense mutation Arg91Leu in b-sarcoglycan. Neuromuscul Disord 1998; 8:193–197. Ben Othmane K, Speer MC, Stauffer J, Blel S, Middleton L, Ben Hamida C, Etribi A, Loeb D, Hentati F, Roses AD, Ben Hamida M, Pericak-Vance MA, Vance JA: Evidence for linkage disequilibrium in chromosome 13-linked Duchenne-like muscular dystrophy (LGMD2C). Am J Hum Genet 1995;57:732–734. Piccolo F, Jeanpierre M, Leturcq F, Dode C, Azibi K, Toutain A, Merlini L, Jarre L, Navarro C, Krishnamoorthy R, Tome FM, Urtizberea JA, Beckmann JS, Campbell KP, Kaplan JC: A founder mutation in the gamma-sarcoglycan gene of gypsies possibly predating their migration out of India. Hum Mol Genet 1996;5:2019–2022. Lasa A, Piccolo F, de Diego C, Jeanpierre M, Colomer J, Rodriguez MJ, Urtizberea JA, Baiget M, Kaplan J, Gallano P: Severe limb girdle muscular dystrophy in Spanish gypsies: Further evidence for a founder mutation in the gamma-sarcoglycan gene. Eur J Hum Genet 1998;6:396–399. Homburger F, Nixon CW, Eppenberger M, Baker JR: Hereditary myopathy in the Syrian hamster: Studies on pathogenesis. Ann NY Acad Sci 1966;138:14–27. Jasmin G, Eu HY: Cardiomyopathy of hamster dystrophy. Ann NY Acad Sci 1979;317:46–58. Mizuno Y, Noguchi S, Yoshida M, Nonaka I, Hirai S, Ozawa E: Sarcoglycan complex is selectively lost in dystrophic hamster muscle. Am J Pathol 1995;146:530–536. Nigro V, Okazaki Y, Belsito A, Piluso G, Matsuda Y, Politano L, Nigro G, Ventura C, Abbondanza C, Molinari AM, Acampora D, Nishimura M, Hayashizaki Y, Puca GA: Identification of the Syrian hamster cardiomyopathy gene. Hum Mol Genet 1997;6:601–607. Sakamoto A, Ono K, Abe M, Jasmin G, Eki T, Murakami Y, Masaki T, Toyo-oka T, Hanaoka F: Both hypertrophic and dilated cardiomyopathies are caused by mutation of the same gene, deltasarcoglycan, in hamster: An animal model of disrupted dystrophin-associated glycoprotein complex. Proc Natl Acad Sci USA 1997;94:13873–13878. Sakamoto A, Abe M, Masaki T: Delineation of genomic deletion in cardiomyopathic hamster. FEBS Lett 1999;447:124–128. Coral-Vazquez R, Cohn RD, Moore SA, Hill JA, Weiss RM, Davisson RL, Straub V, Barresi R, Bansal D, Hrstka RF, Williamson R, Campbell KP: Disruption of the sarcoglycan-sarcospan complex in vascular smooth muscle: A novel mechanism for cardiomyopathy and muscular dystrophy. Cell 1999;98:465–474. Bo¨nnemann CG, Darras BT, Lidov HGW, Feener CA, Shapiro FD, Kunkel LM: Strength improvement on prednisone in a patient with limb-girdle muscular dystrophy 2C (primary c-sarcoglycanopathy). Ann Neurol 1997;42:531–532. Connolly AM, Pestronk A, Mehta S, Al-Lozi M: Primary alpha-sarcoglycan deficiency responsive to immunosuppression over three years. Muscle Nerve 1998;21:1549–1553.

Bo¨nnemann

42

94

95

96 97

98

99

100

Holt KH, Lim LE, Straub V, Venzke DP, Duclos F, Anderson RD, Davidson BL, Campbell KP: Functional rescue of the sarcoglycan complex in the BIO 14.6 hamster using delta-sarcoglycan gene transfer. Mol Cell 1998;1:841–848. Greelish JP, Su LT, Lankford EB, Burkman JM, Chen H, Konig SK, Mercier IM, Desjardins PR, Mitchell MA, Zheng XG, Leferovich J, Gao GP, Balice-Gordon RJ, Wilson JM, Stedman HH: Stable restoration of the sarcoglycan complex in dystrophic muscle perfused with histamine and a recombinant adeno-associated viral vector. Nat Med 1999;5:439–443. Li J, Dressman D, Tsao YP, Sakamoto A, Hoffman EP, Xiao X: rAav vector-mediated sarcogylcan gene transfer in a hamster model for limb girdle muscular dystrophy. Gene Ther 1999;6:74–82. Nowak KJ, Walsh P, Jacob RL, Johnsen RD, Peverall J, McNally EM, Wilton SD, Icakulas BA, Laing NG: Severe c-sarcoglycanopathy caused by a novel missense mutation and a large deletion. Neuromuscul Disord 2000;10:100–107. Thompson TG, Chan YM, Hack AA, Brosius M, Rajala M, Lidov HG, McNally EM, Watkins S, Kunkel LM: Filamin 2 (FLN2): A muscle-specific sarcoglycan interacting protein. J Cell Biol 2000;148:115–126. Durbeej M, Cohn RD, Hrstka RF, Moore SA, Allamand V, Davidson BL, Williamson RA, Campbell KP: Disruption of the beta-sarcoglycan gene reveals pathogenetic complexity of limb-girdle muscular dystrophy type 2E. Mol Cell 2000;5:141–151. Liu LA, Engvall E: Sarcoglycan isoforms in skeletal muscle. J Biol Chem 1999;274:38171–38176.

Carsten G. Bo¨nnemann, Department of Neuropediatrics, University Childrens’ Hospital, Robert-Koch-Strasse 40, D–37075 Go¨ttingen (Germany) Fax +49 551 39 6252, E-Mail [email protected]


43


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

Facioscapulohumeral Muscular Dystrophy: Diagnostic and Molecular Aspects Peter W. Lunt Clinical Genetics Unit, Bristol Royal Hospital for Sick Children, Bristol, UK

Facioscapulohumeral muscular dystrophy (FSHD) has a prevalence estimated at between 2.5–5.0/100,000, and a birth incidence of heterozygote gene carriers around 1/20,000 [1, 2]. The genetic defect follows autosomal dominant inheritance, but with new mutation accounting for around 10% of recognised cases [3]. A typical symptomatic presentation is in teenage (the 2nd decade of life), with shoulder girdle weakness and associated muscle wasting, with examination at that time revealing characteristic peri-oral and/or peri-orbital facial weakness, which in retrospect has usually preceded the shoulder girdle symptoms. The condition is progressive, with proximal lower limb involvement sufficient to require a wheelchair developing in 10–20% of cases [2]. However, the range of age and severity of presentation between families, and also to a considerable degree within families, is extremely wide, varying from minimal signs of asymptomatic scapular girdle or facial weakness in the 7th decade, to cases with severe progressive proximal and distal upper and lower limb involvement together with ‘Mo¨bius-like’ expressionless facies by early childhood [4]. The molecular basis underlying this variability is now beginning to be understood, and facilitating some degree of prediction within a family. FSHD is caused by a so far unique mutation mechanism, and is perhaps also unique amongst genetic conditions in being one where the DNA mutation is known, but where there is yet no knowledge of the gene (or genes) affected by this. Thus, the genetic defect causing FSHD is deletion of an integral number of 3.3-kb repeat units located at subtelomeric chromosome region 4q35, reducing the normal 10–96 tandem repeat copies (40- to 320-kb fragment) down to a residual 1–10 copies (10- to 40-kb fragment) [5, 6]. There is

a closely homologous 3.3-kb repeat array located at the subtelomeric chromosome region 10q26, and some evidence for chromosomal translocation (or sequence conversion) between these two regions, possibly predisposing to the DNA rearrangement causing FSHD [7]. Diagnostic molecular testing can be offered through this ‘uniform mutation mechanism’, both within families for prediction, and for single representative cases where confirmation of diagnosis is sought. This is made possible with a high specificity where there is a ‘positive’ molecular result [3]. The sensitivity of testing has still to be improved, and exclusion of diagnosis in a single representative case with a neuromuscular presentation is still not possible from a molecular genetic laboratory result alone. It is found that the age at onset and severity of clinical presentation correlates broadly and inversely with the size of the residual DNA fragment at 4q35, and, by inference, therefore correlates directly with the number of repeat units deleted [8–10]. Thus, the smallest residual fragment lengths at 10–17 kb (1–3 repeat copies) are usually associated with a severe infantile or childhood presentation [4, 11], medium lengths (18–30 kb, or 4–7 repeat copies) are often found in the largest recognised dominant families, while the largest lengths (31–38 kb, or 8–10 repeat copies) have been associated with a milder predominantly scapulohumeral presentation, and may well have reduced penetrance, particularly in females [8, 12, 13]. New mutation cases are seen predominantly with the smallest residual fragment lengths, giving matching clinical severity, and may originate predominantly on the maternal copy of chromosome 4 [6, 13]. Study of parental DNA suggests that around 20–30% of new mutations occur as somatic and germline events in one of the parents, this usually also being the mother [3, 13, 14, 43]. The favoured hypothesis for the molecular pathogenesis is for the deletion of 3.3-kb repeats to alter the chromosomal configuration or folding at terminal 4q region, and thereby exert a positional effect on a gene or genes located proximally to this, possibly through proximal spread of telomeric heterochromatin [15]. The 3.3-kb repeat units appear not to be transcribed themselves, although the presence of a paired homeodomain sequence within each repeat, with sequence similarity to other transcription regulators, induces other plausible hypotheses for a more direct role in regulation of expression of other genes [16, 17]. To date, no gene has been identified whose expression is influenced directly by the 3.3-kb repeat lengths. Clinically, it is now recognised that many patients with FSHD have evidence for a subclinical hearing loss, most marked at 6,000 Hz [18]. Others may have evidence of a retinal vasculopathy, manifesting at its most severe (as Coats’ disease-like changes) in the youngest onset FSHD cases [19, 20]. Similarly, it is now being reported that epilepsy and mental retardation may be associated with some of the most

Facioscapulohumeral Muscular Dystrophy: Diagnostic and Molecular Aspects

45

severe FSHD presentations, suggesting the possibility of a CNS effect with the largest deletions [11]. The concept of the clinical presentation in an FSHD case being determined by the probability of significant disturbance of site-specific gene expression dependent on the total length of the 3.3-kb repeats is emerging. With such a model, there would be no strict threshold at the mildest end of the spectrum, but rather an overlap between minimally affected cases and unaffected subjects from the normal population, with overlap between repeat copy numbers at the lowest end of the ‘normal range’ with those which could manifest as a mildest presentation of FSHD or scapulohumeral muscular dystrophy. Delineation of factors controlling this probability, particularly demonstrated by wide clinical variability within families, could potentially lead to methods of therapeutic intervention, which currently are restricted to ameliorative surgery, such as scapular fixation, tendon transfers, and blepharorrhaphy [20, 21]. Meantime empirical treatments with corticosteroids or b2-agonists are being evaluated on a trial basis [22], while increasing interest in FSHD patient groups is providing a focus for raising and providing support.

Diagnosis of Index Case The diagnosis of FSHD in an apparently isolated case, or in the index case in a family context, now depends on a combination of the clinical presentation, family history, and molecular genetic analysis. Published clinical diagnostic criteria comprise ‘inclusion’ and ‘exclusion’ criteria with ‘additional comments’ [23]. The inclusion criteria broadly require shoulder girdle involvement prior to any pelvic girdle involvement, facial weakness (peri-oral, peri-orbital or both) in an index case, or in at least 50% of the affected cases in a family, and proximal prior to distal involvement in the upper limbs, but with early distal (peroneal) weakness often a characteristic feature in the lower limbs. Initial asymmetry of shoulder girdle involvement is usual, with the right side affected first in over 75% of cases, and possibly with a relationship to handedness [2]. Suggested exclusion criteria (or at least cause for considerable caution) are a sustained improvement of symptoms, presence of ptosis or extra-ocular muscle involvement, and onset of weakness in the pelvic girdle prior to the shoulder girdle, without typical facial weakness [23]. Muscle hypertrophy, particularly in forearm and calf muscles, can be seen, and muscular pain can be a prominent symptom [20]. Muscle biopsy may show few or non-specific changes, but can show typical dystrophic features [1]. Inflammatory change and small angular fibres are often seen. Specific biopsy features of other neuromuscular conditions should not be present. However, in a clinically

Lunt

46

suspected case, DNA testing rather than muscle biopsy could now be the first line investigation. Clinical presentation in a typical case is usually as a teenager who first becomes aware of developing symptoms of shoulder girdle weakness, or of signs of muscle wasting in this region. Data from large families with many affected individuals suggests a median onset age of 11 years for clinical signs (usually as facial weakness) to be first recognisable by an examiner, preceding by a few years the median age for the patient to first complain of symptoms [2, 24]. Typically the teenager notices a ‘dropped’ shoulder contour or thinning of the upper arms, possibly with increased winging of the scapulae, and admits to difficulty raising one arm (usually the right one). Facial examination will usually show weakness of eye closure – minimal involvement being an inability to bury the eyelashes on attempted tight eye closure. Weakness of peri-oral muscles is often evident to the attuned eye as a slight asymmetry of the lips, but is best demonstrated by air escape when the examiner tries to force air out of the patient’s puffed cheeks. As well as confirmatory findings of shoulder girdle weakness, an exaggeration of the normal lumbar lordosis with a positive Beevor’s sign [25], and a degree of peroneal weakness may already be present at this stage. In more severe infantile-onset cases, facial weakness is the earliest and most prominent sign. Thus, the infant may show little or no facial expression, appearing unable to smile, and may be initially misdiagnosed as having Mo¨bius syndrome [4]. Pelvic girdle weakness in the most severe cases can be prominent by age 10 years, leading to consideration of Xp21 or limb girdle types of muscular dystrophy, but unlike these conditions, FSHD is still characterised by an even greater degree of shoulder girdle weakness rather than pelvic weakness. FSHD is inevitably progressive, and an overall 20% of patients require a wheelchair by the 5th decade, although this can be required before age 20 years in many of the most severe new mutation cases [2]. Harder to recognise is the milder, later onset presentation, which tends to be associated with larger (32–38 kb) DNA fragment sizes at 4q35 with fewer copies deleted of the 3.3-kb repeats. Many of these gene carriers may remain unaware of symptoms, or not attribute symptoms to the family condition, but can be recognised in a family context by signs of peri-oral or peri-orbital weakness, by a minor degree of scapular winging or scapular weakness, or most often by an asymmetrically ‘dropped’ shoulder contour. However, in this group, signs of facial weakness may be minimal or absent, and some families classified with a dominant scapulohumeral form of muscular dystrophy represent the mild end of the spectrum of FSHD-associated 3.3-kb repeat deletion with fragment size of 35–38 kb [12]. Penetrance of clinical signs, previously estimated at 95% by age 20 years [24], now appears to be significantly lower in females compared to males,


47

particularly with the larger DNA fragment sizes [13]. The family history is therefore often particularly helpful in establishing diagnosis. Stories of relatives who have had difficulty raising their arms, have had a ‘rounded’ shoulder contour, or have apparently ‘thrown their leg’ when walking are particularly helpful. Since it is only in the more severely affected cases that reproductive fitness may be reduced, one can expect a high proportion of FSHD cases in the milder half of the spectrum (i.e. with fragment size q25 kb) to have other affected relatives. Examination of parents and sibs of all cases can therefore be very helpful diagnostically, particularly in relation to facial weakness, in providing more than one clinical presentation for evaluating against the standard clinical criteria, but this has to be balanced against the problems inherent in uncalled-for genetic diagnosis of relatives who might have preferred to remain unaware of the presence of clinical signs and diagnosis in themselves. Information on relatives identified in family histories should also be verified where possible. Anticipation It is uncertain whether there may be evidence of clinical anticipation in FSHD, whereby the severity tends to increase with successive generations [8–10]. At present it is not clear how this could come about with a fixed mutation in each family, but the same has been proposed in at least one other neurological condition with a fixed mutation – familial amyloid polyneuropathy [26]. One could hypothesise that the fixed deletional mutation, by affecting chromosome folding or telomeric pairing at meiosis, might be inducing a dynamic expansion (say) in a different type of tandem repeat located more proximally on chromosome 4q, leading to further expansion at subsequent meioses, but this must remain as pure conjecture in the absence of any known dynamic repeat sequence in the 4q35 region. Furthermore, the finding of DNA evidence for somatic and germline mosaicism for a ‘severe’ mutation [13, 14], as an explanation for minimal symptoms in one parent in some ‘new mutation’ cases, and the knowledge that females may generally have a milder presentation than males [2, 14], might provide more plausible explanations for at least some cases of apparent anticipation [43].

Molecular Testing: Confirmation of Diagnosis In 90–95% of cases of FSHD, as defined by meeting the diagnostic criteria, the diagnosis can effectively be confirmed by showing the presence of a shortened (=35 kb) DNA fragment at 4q35 (recognised by probe p13E-11), which arises from deletion of an integral number of copies of the 3.3-kb repeats

Lunt

48

from that region [3, 5]. The DNA probe used (p13E-11) also detects the closely homologous 3.3-kb repeat array from 10q26 [7]. However, each chromosome 10-type repeat has an additional BlnI restriction site [27]. For the specific diagnostic test, a double digest with EcoRI/BlnI is employed on genomic DNA (obtained from peripheral blood), which removes chromosome 10-type repeats, but leaves chromosome 4-type repeats intact (albeit reduced by 3 kb in size compared to EcoRI single digest) [27]. Test specificity for BlnI-resistant fragments smaller than 35 kb (approximately 8¶3.3-kb repeats) in someone with a neuromuscular presentation is very high (98%), since BlnI-resistant fragments of p35 kb are found in =2% of the normal population [3]. However, the sensitivity of the test is lower. In around 6% of clinically affected cases which satisfy the diagnostic criteria but appear as ‘false-negatives’, and in the 2% or so of controls who would be ‘false-positives’, a more complicated situation exists with respect to the arrangement of 3.3-kb repeat sequencies at 4q35 and 10q26, requiring characterisation of all four repeat arrays for interpretation [7, 28]. This is achieved by using pulse field gel electrophoresis (PFGE) or field inversion type (FIGE), with EcoRI and EcoRI/BlnI digests, to identify DNA fragments up to 200 kb in size, hence the length and Bln sensitivity ‘type’ of the four separate repeat arrays [6]. Only 80% of the normal control population have two repeat arrays of each type; 20% show polymorphism for either a translocation or gene sequence conversion between the repeats at the 4q35 and the 10q26 regions, recognised on PFGE as having three fragments of one type and one of the other, or rarely by four fragments of the same type [7]. FSHD arises when the repeat array on chromosome 4 is reduced in size (copy number), irrespective of whether the repeat units are of 4-type or 10-type. Hence there are 5% of patients with FSHD who have a =38-kb fragment, but which is Bln-sensitive, owing to this being a shortened 10-type repeat array on one chromosome 4, and usually associated with a 1:3 ratio of 4-type:10type repeats on PFGE. Variation in length of the repeat arrays attached to chromosome 10 seems to be of no consequence, and nearly 60% of controls have a fragment which appears in the same size range as FSHD-associated fragments (=38 kb, if families with scapulohumeral presentation are included) [6], although in =4% is this Bln-resistant and hence might confuse diagnostic testing [3]. Table 1 shows the distribution of fragment size and type in FSHD patients and controls. The presence of a shortened Bln-resistant fragment on conventional gel electrophoresis as a diagnostic test for FSHD would for fragments =32 kb give test sensitivity of 85% and specificity of 98.5%, and for fragments =38 kb give test sensitivity of 94% and specificity of 96.5%. For milder presentations with fragment sizes 32–38 kb, or for Bln-sensitive fragments, or for


49

Table 1. Distribution of fragment size and type (Bln-resistant or Bln-sensitive) detected on standard gel electrophoresis in FSHD patients and in controls Conventional gel fragment seen

FSHD, % Overall With 2EB:2E With 1EB:2–3E With 0EB:3–4E With 2–3EB:1E With 3–4EB:0E Controls, % Overall With 2EB:2E With 1EB:2–3E With 0EB:3–4E With 2–3EB:1E With 3–4EB:0E

Bln-resistant

no Bln-resistant, but Bln-sensitive

=32 kb 32–38 kb

=32 kb 32–38 kb

85 73 4 – 8 0.2 1.5 0.6 0.01 – 0.85 0.04

9 8 0.4 – 0.9 0.02 2 0.2 0.005 – 1.7 0.1

Likelihood ratios of FSHD:control With 2EB:2E 120:1 40:1 With 1EB:2–3E 400:1 80:1 With 0EB:3–4E – – With 2–3EB:1E 10:1 1:2 With 3–4EB:0E 5:1 1:5

4.7 0.5 4 0.2 0.015 – 19 16 2 0.05 1 – 1:30 2:1 4:1 1:60 –

0.9 0.05 0.8 0.05 0.02 – 39 33 3.7 0.1 1.8 – 1:600 1:5 1:2 1:100 –

total

no frag.=38 kb

0.4 Not FSHD 0.34 0.02 0.04 0.001

100 81.5 9.5 0.3 9 0.2

39 31 3.8 0.1 3.8 0.01

100 81 9.5 0.25 9 0.15

Not FSHD 1:10 1:5 1:100 1:10

This is given as the overall percentage distribution seen in FSHD or in controls, and also as the distribution according to the possible ratios of 4-type (EB):10-type (E) repeat arrays seen on PFGE. From this a likelihood ratio of FSHD: control has been calculated for each fragment size/type and possible 4-type:10-type fragment ratio.

testing to exclude a diagnosis of FSHD, it is essential to add in PFGE. Therefore, for each category of shortened fragment, table 1 also gives a calculated likelihood ratio of FSHD:control according to the overall ratio of 4-type:10-type repeat arrays seen on PFGE. Two other rare situations have been observed which can add further complication: (1) deletion extending into the p13E-11 hybridisation site, and (2) hybrid fragments, comprising repeats of both chromosome 10 and chromosome 4 type [28]. Recognition of either of these situations requires use of a

Lunt

50

second probe 9B6A which is complimentary to the repeat unit (D4Z4) itself. Subjects deleted for the p13E-11 site will appear to have no small fragment on 4q35 when hybridised with p13E-11, which may lead to an initial interpretation as exclusion of FSHD. However, they show only 3 bands on PFGE/FIGE using this pobe, but a ‘hidden’ fourth band (which is =38 kb in FSHD) if 9B6A is used instead [28]. This has been observed with a normal length repeat array in an unaffected father and son, ascertained through further extension of the deletion resulting in a shorter repeat array and de novo FSHD in an affected son [28]. It is not known whether p13E-11 site deletion may be prevalent in the general population, or whether it would have a high propensity to further extension and inevitably be ascertained only as a rare association with de novo FSHD. If we are correct that a more proximal gene locus is being influenced by the repeat deletion, since 4q35 haploinsufficient states (with cytogenetically detectable terminal deletions) are not known to show any features of FSHD [29], the proximal limit of molecular deletions causing FSHD defines a distal limit for the location of an influenced gene locus [28]. Hybrid repeat arrays, consisting of a run of 10-type repeats attached to the distal end of a 4-type repeat array, could lead to a false-positive diagnosis of FSHD since EcoRI/BlnI double digest may show a small Bln-resistant fragment segregating with 4q35, but corresponding to the length of the 4-type repeats only. Using PFGE /FIGE the apparent disappearence of a larger EcoRI fragment (from the hybrid array) may be possible to recognise [28], but also requires use of probe 9B6A to avoid confusion where a 10-type repeat array at 10q26 happens to be of similar size to the Bln-resistant residual fragment. Development of a new technique, using a BlnI/BglII double digest, is anticipated to provide a ‘dosage’ test which on standard gel electrophoresis using p13E-11 will be able to assess the ratio of 4-type to 10-type repeats in the most proximal repeat unit for all four chromosomes concerned, and hence help identify translocated (or sequence-converted) repeats and p13E-11 site deletions [30]. The clinical importance of these laboratory findings is that in sending a sample to the laboratory for diagnostic testing, given that although test specificity may be very high, because the sensitivity is lower the clinician must inform the lab whether he is expecting a diagnosis of FSHD to be confirmed, or whether he is trying to exclude the diagnosis. An estimated prior probability of the diagnosis being truly FSHD can be combined by Bayes’ calculation with a likelihood ratio for a fragment of given size and Bln-type to be associated with FSHD or not (FSHD:control ratio in table 1). Note that owing principally to 0.5% of FSHD subjects who have a double exchange with a 4-type array at 10q26 and a shortened 10-type array at 4q35, and 6% of controls who have three 10-type fragments with a shortened one at 10q26, the sensitivity and specificity of testing


51

cannot yet reach higher overall than 99.5 and 94%, respectively. Also, DNA samples suitable for PFGE must be extracted in a specific way and from a fresh blood sample, imposing some practical limitations on the testing. Our own experience of running a diagnostic molecular service in Bristol, UK based on conventional gel electrophoresis only has been that 21% of 270 referrals have no fragments =40 kb and are probably not FSHD; 14% of those with fragments =35 kb have this as a Bln-sensitive fragment. This data suggests that if 25% of controls have a Bln-sensitive fragment =35 kb, then it will be 5% of true FSHD who have a shortened 10-type fragment on 4, as predicted. In the above calculations for sensitivity and specificity of testing, independence of deletional mutation and background chromosome translocation pattern has been assumed. However, if parents of ‘new mutation’ cases might have an excess of shorter or translocated repeat types, perhaps predisposing to de novo deletion, the sensitivity and specificity of testing may be less good [31, 43]. This would also imply that some people (or perhaps the fertilised eggs of some couples) could have an increased susceptibilty to deletional reduction of the 4q35 repeat array, and therefore to having a child with ‘de novo’ FSHD. Scapulohumeral Presentation and Fragment Lengths of 34–40 kb An as yet undefined proportion of families with FSHD have a milder scapulohumeral presentation with a Bln-resistant fragment on 4q35 in the size range 34–40 kb (8–10 repeats) [8]. In one family from Somerset, UK, in which affected subjects demonstrated a scapulohumeral syndrome without facial weakness, a 4q35-cosegregating 38-kb fragment is found in all definitely affected members, and in 2 at-risk young adults who are presumed to be presymptomatic [12]. In this and similar families, penetrance is likely to be reduced, particularly in females [13]. Bln-resistant fragments of this size can also be seen in some normal controls – thereby reducing test specificity in this size range to a level which may be impracticable for diagnostic use. Hence there is probably no definite threshold between the normal unaffected situation and the most mild degree of shoulder girdle weakness. Rather, one can envisage with increasing repeat lengths (fragment size) a gradually decreasing probability that a shorter than usual repeat array will in any particular cell in the body affect the expression of some other gene (or genes) adversely, and hence an overall decreasing probability that symptoms will develop in any one individual. Addition of 4q35 and 10q26 Linkage Analysis In families where there remains doubt about the diagnosis from molecular testing, the addition of linked polymorphism analysis may be very helpful in determining whether particular fragments cosegregate with 4q35 or 10q26 markers. The author has experience of 2 teenage brothers who presented with

Lunt

52

scapular winging (in both) and spondylolisthesis (in one), but normal muscle biopsy. The finding of Bln-sensitive fragments of size 28 kb in one boy and 33 kb and 37 kb in the other was resolved by 4q35 and 10q26 marker analysis in both boys, their sister and their parents. FSHD was excluded by showing that none of these fragments appeared to segregate with 4q35 markers, but all 3 were compatable with 10q26 linkage of the fragments.

Questions of Patients and Parents Once Diagnosis Has Been Confirmed Prognosis FSHD inevitably involves a progressive weakness, although can plateau for long periods. Loss of ambulation, requiring wheelchair use, is found in 20% of patients aged between 40 and 83 years, but in this group, all patients with disabling proximal lower limb weakness had in retrospect already been aware of at least some lower limb weakness by age 20 years [2]. Several studies of the retrospectively reported age at onset of first clinical signs have now each shown a broad correlation between average age at onset and FSHD-associated DNA fragment size [8–10], this being 10–18 kb in most infantile-onset cases; 18–34 kb in most typical teenage-onset cases, and 30 kb or above in the eldest-onset cases. There may be an additional generational (anticipation) effect, with the mildest clinical cases within a family (after exclusion of the direct family line) still tending to be those in the top generation [8–10], but further study of additional pedigrees is required. Males may have a slightly younger average onset age than females, and adult females may be more likely than males of the same age to show nonpenetrance [13]. This may be a hormonal effect rather than a genetic one, with severity more equal above age 50–60 years [32]. Currently, parents can be advised that the presentation of FSHD in any affected offspring is likely to be at least as severe as in a same-sex affected parent and quite possibly greater where a mother is passing the condition to a son. Prediction of the likely severity in a daughter receiving the condition from her father is less certain, although anecdotally it is hard to think of any definite examples where it has been significantly later onset or milder. Therapeutic Strategies and Other Support There are no specific treatments identified for FSHD. The observation of a tendency to earlier and greater periscapular and upper arm weakness on the side of the dominant hand (usually right side) [2], together with the often irreversible muscle wasting of a limb following enforced immobility (e.g. secondary to a fracture) [1] suggest that the primarily impaired process might be muscle re-


53

generation following damage. If so, strenuous body-building exercise would certainly be contraindicated, while investigation of factors controlling the process of muscle regeneration could lead to potential treatments. A trial of steroids has not shown any sustained benefit [33], but a trial of a b2-agonist (albuterol) is currently being evaluated [22]. For certain patients, surgery to fixate the scapula to the chest wall has proved successful, and enabled greater use of the arms, albeit at the restriction of becoming unable to raise them above shoulder level [20, 21]. A proper understanding of factors controlling variation of severity within a family, and even within a sibship, might potentially lead to therapeutic intervention to influence a genetic modifier mechanism. General support can be very helpful, including physiotherapy and occupational therapy, but equally, contact with specific or more general neuromuscular patient support groups will become increasingly helpful, particularly if contact between patients in different families can be facilitated through the Internet. Genetic and Clinical Testing Wider in a Family De novo cases of FSHD can only be proven as such by recognition of de novo appearence of a typical Bln-resistant short fragment, where DNA from both parents is available and paternity is proven. In all other cases there remains a possibility of genetic risk wider in the family, through unrecognised or asymptomatic gene carriers, and a wider family study is indicated [31]. Proven de novo cases have almost all involved a DNA fragment smaller than the average size for familial affected cases (i.e.=25 kb) [8]. This may reflect an ascertainment bias for paediatric onset cases, or a higher genetic fitness and smaller proportion of new mutations amongst cases with larger fragments, but could also possibly indicate a higher overall mutation rate for smaller fragments. New mutations, whether occurring in meiosis or mitosis (the latter giving rise to somatic and germline mosaicism), have predominantly been maternal (80% of recognised parental mosaics being in the mother) [6, 3, 13, 14], but whether this might indicate a parental sex difference for the size of de novo deletion, analagous to the parental sex difference between de novo deletion and point mutation in Xp21 dystrophy, has not yet been tested. Alternatively, more male mosaics may show symptoms [43]. A family study involves both clinical and molecular evaluation, which is only possible if the index case has a proven and readily recognisable repeat array deletion at 4q35, giving a characteristic DNA fragment. Quite often, the attuned clinical examiner will detect subtle but typical clinical signs in a relative, which had previously gone unnoticed, particularly with respect to minor peri-oral or peri-orbital weakness. The issue of informed consent becomes paramount, and if a relative believes themselves to be clear, perhaps clinical examination as well as DNA testing of that person should follow a

Lunt

54

similar protocol to predictive DNA testing in other late-onset conditions. Supportive linkage analysis at 4q35 and 10q26 should also be performed to minimise any chance of a spurious positive prediction from a 10q26 fragment. This would usually require DNA samples from unaffected spouses as well as blood relatives. In apparently clinically isolated cases, the family study should commence with both parents of the affected subject, to recognise in particular the 20% of cases where one parent is mosaic for the mutation, which shows as a faint band on the gel corresponding to the affected band in their offspring. In a further 12% of cases, one parent (equal sex ratio) has demonstrated the fragment in full dose, but remains asymptomatic [3], including at least 2 cases where a very small (13 kb) fragment has been associated with a very severe presentation in their offspring [31], although in one of these, further PFGE study identifying 5 bands in the father suggests that he may be a mosaic. The presence in two other families of a 25-kb 4q35-cosegregating fragment in both a parent and grandparent who each remain asymptomatic would seem inadequately explained by the possibility of a reduced penetrance in females, and has led to the suggestion of a 2-step mutation process [31]. Penetrance and Risk to Offspring Original data suggesting that a family member, born at 50% risk, has only a 4.8% chance after age 20 years of still carrying the FSHD mutation if they show no detectable clinical sign of the condition (95% penetrance) was based on data from families demonstrating dominant inheritance and of sufficient size to be able to exclude the proband, his sibs, parents and grandparents from analysis [24]. Most such families have fragments from the middle size range (18–30 kb), since ascertainment of families with larger fragments is likely to be less complete, while with the shortest fragment lengths the greater severity limits family size. With smaller fragments (10–18 kb) the age-dependent penetrance will approach 100% by age 20 years, whereas with larger fragments (30–38 kb) it may well be much lower, particularly in women, but the presentation is also likely to remain relatively mild. Hence, a relative of someone with a severe FSHD presentation (apart from the parents, one of whom may be a mosaic) can generally be reassured that they are very unlikely to be carrying the same gene defect if they remain clear above age 20 years. However, details of the fragment size in the index case should be checked first, and with a potential possibility of a 2-step mutation process, reassurance can only really be given to a relative following a negative DNA test. If, as seems likely, there is a gradually reducing penetrance with increasing fragment size, there will be a significant region of overlap between normality and a mild FSHD presentation, such that any proactive widening of a family study in families with the larger fragment sizes (32–40 kb) would seem unwise. However, this is an area requiring further research.


55

Testing of Children It is usually inappropriate to test a child for a potentially later-onset genetic condition for which there is no treatment, if they are not showing any sign of the condition. For FSHD, only if an affected sibling has had a severe early childhood-onset presentation with matching small fragment size, such that a younger sib inheriting the same 3.3-kb repeat deletion would be expected inevitably to develop significant symptoms also in their early childhood, might testing be considered. Linked DNA marker analysis should be performed simultaneously. However, a single report of deletional expansion occurring between (an albeit asymptomatic) parent and one offspring [28] behoves caution even in the situation above, to avoid predicting for a milder later-onset presentation. Prenatal Testing This is possible from chorion villus biopsy (CVB) from 11 weeks’ gestation onwards, but similarly requires certainty about the FSHD-associated fragment in the family, must be run together with DNA from both parents, and must be backed up by linked markers. Analysis is by Southern blot, which requires a larger CVB sample and a longer time for analysis than in other conditions which can employ PCR techniques. The possibility of developing a PCR assay for the fragments is being investigated, but so far has only been able to detect fragments up to 25 kb in length [3].

Basic Science FSHD is a condition where the mutation is known, but not the gene or genes on which it acts. If there were unequivocal genetic heterogeneity, the localisation and possibility of cloning of a gene involved in a second and non4q35 locus could well add additional clues to the molecular and cellular pathogenesis. To date, there are only two large FSHD families published which appear to be unlinked to 4q35 (both from one US genetic centre) [34], but in neither has any alternative chromosomal localisation, including 10q26, been identified [35]. Hypotheses for possible mechanisms of action of the deletion of 3.3 kb repeats include the following. Position Effects: (1) If the repeats are normally important for subtelomeric chromosome folding, the disturbance of this may well alter the expression of a gene or genes sited more proximally along chromosome 4. (2) The telomeric region of all chromosomes is folded/methylated as heterochromatin. Deletion of part of the area involved in this might cause a region more proximally to be folded/methylated as heterochromatin, and therefore genes from that area

Lunt

56

could be inactivated by ‘spreading heterochromatisation’. (3) The 3.3-kb repeats might help align the 4q and 10q telomeres on the spindle during cell division, perhaps facilitating some expressional interaction between genes on these chromosomes, which would be disturbed by 3.3-kb repeat deletion. More Direct Regulator Role of the 3.3-kb Repeats: (4) Each repeat contains a paired homeodomain suggesting some role in temporal, regional or tissuespecific regulation of gene expression [16]. ‘STOP’ codons in the DNA sequence of the repeat preclude an open reading frame, and the repeats appear not to be transcribed. However, sequence homology with genes for a DUX family of DNA-binding regulatory proteins seems more than a coincidence [17]. Perhaps an RNA transcript itself might have a regulatory role. In all these possible models, it is suggested that the effect of the mutation occurs at a distance from the mutation itself. Attempts to find a gene more proximally on 4q35, whose protein product is expressed in muscle and is regulated by the number of 3.3-kb repeats, have to date been unsuccessful. At least three possibilities for candidate genes have been identified (FRG1, ALP, TUB4q), but none of these show altered expression in FSHD muscle tissue [36–38]. Some evidence suggests that the expression of multiple genes may be affected by the repeat deletion, and variously up- or downregulated, but it is not yet clear whether this might be a secondary phenomenon consequent on muscle damage. Cases of karyotypic deletion of 4q35 have not exhibited any muscular dystrophy [29]. This suggests that FSHD cannot be consequent on haploinsufficiency, but is probably a ‘gain of function’ effect. The correlation of deletion size with severity, the absence of facial weakness with the smallest deletions, and the singular distribution of muscle involvement in FSHD suggest that a small deletion may leave intact the regulatory mechanism for expression of some gene in facial muscles but not in the shoulder girdle. Similarly the tendency for the most severe cases to be the ones with the most marked retinal changes and hearing loss, and possibly also to have associated seizures and mental retardation [11], suggests that the largest deletions may tend to affect gene expression at sites additional to muscle. Several parallels with myotonic dystrophy could be more than a coincidence; in both conditions facial and peroneal muscle weakness can be a prominent feature, and in both there is a paired homeodomain sequence involved in, or in the vicinity of, the mutation [39]. That the only limb girdle muscular dystrophy in which the upper limb girdle is the one predominantly affected (type 2A) is the only one where the gene codes for a proteolytic enzyme (calpain 3) rather than a structural protein [40] is also noteworthy. Comparative studies with the genomes of other organisms have not yet provided useful leads. The 3.3-kb repeats seem to be limited to higher primate species [41]. The 4q35 region shares homologies in the mouse with sequencies


57

and genes on murine chromosome 8, but the only neuromuscular phenotype (myodystrophy) determined by a gene (myd) in this murine chromosome region is inherited as a recessive condition, and has closest flanking markers which show homology much more proximally on human 4q [42].

Future (Speculation) Further research is expected eventually to identify the nature of the gene (or genes) which are influenced by the 3.3-kb repeat deletions, the normal role of the repeats themselves, and the mechanism of mutation. Clues may come potentially from genetic linkage studies for polymorphisms which may influence variation of severity between siblings. Identification of any such modifier gene could potentially lead to initial forms of therapy, at least to help minimise the severity of presentation. Knowledge of the mutation mechanism may have wider implications for understanding the control of chromosome folding, and the possible role of this in regulation of gene expression.

References 1 2 3 4

5

6

7

8

9

Padberg GW: Facioscapulohumeral Disease; thesis, Leiden University, 1982. Lunt PW, Harper PS: Genetic counselling in facioscapulohumeral muscular dystrophy. J Med Genet 1991;28:655–664. Lunt PW: 44th ENMC International Workshop: Facioscapulohumeral Muscular Dystrophy: Molecular Studies. Neuromuscul Disord 1998;8:126–130. Jardine PE, Koch MC, Lunt PW, Maynard J, Bathke KD, Harper PS, Upadhyaya M: De novo facioscapulohumeral muscular dystrophy defined by DNA probe p13E-11 (D4F104S1). Arch Dis Child l994;71:221–227. van Deutekom JC, Wijmenga C, van Tienhoven EAE, Gruter A-M, Hewitt JE, Padberg GW, van Ommen G-J, Hofker MH, Frants RR: FSHD associated DNA rearrangements are due to deletions of integral copies of a 3.2 kb tandemly repeated unit. Hum Mol Genet 1993;2:2037–2042. Wijmenga C, van Deutekom JCT, Hewitt JE, Padberg GW, van Ommen G-JB, Hofker MH, Frants RR: Pulsed-field gel electrophoresis of the D4F104S1 locus reveals the size and parental origin of the facioscapulohumeral muscular dystrophy (FSHD)-associated deletions. Genomics 1994;19: 21–26. van Deutekom JC, Bakker E, Lemmers RJ, van der Wielen MJR, Bik E, Hofker MH, Padberg GW, Frants RR: Evidence for subtelomeric exchange of 3.3 kb tandemly repeated units between chromosomes 4q35 and 10q26: Implications for genetic counselling and etiology of FSHD1. Hum Mol Genet 1996;5:1997–2003. Lunt PW, Jardine PE, Koch MC, Maynard J, Osborn M, Williams M, Harper PS, Upadhyaya M: Correlation between fragment size at D4F104S1 and age at onset or at wheelchair use, with a possible generational effect, accounts for much phenotypic variation in 4q35-facioscapulohumeral muscular dystrophy (FSHD). Hum Mol Genet 1995;4:951–958 & erratum 1243–1244. Zatz M, Marie SK, Passos-Bueno MR, Vainzof M, Campiotto S, Cerqueira A, Wijmenga C, Padberg G, Frants R: High proportion of new mutations and possible anticipation in Brazilian facioscapulohumeral muscular dystrophy families. Am J Hum Genet 1995;56:99–105.

Lunt

58

10

11 12

13

14

15

16

17

18 19

20 21 22

23

24

25 26

27 28

Tawil R, Forrester J, Griggs RC, Mendell J, Kissel J, NcDermott M, King W, Weiffenbach B, Figelwicz D, FSH-DY group: Evidence for anticipation and association of deletion size with severity in facioscapulohumeral muscular dystrophy. Ann Neurol 1996;39:744–748. Funakoshi M, Goto K, Arahata K: Epilepsy and mental retardation in a subset of early onset 4q35-facioscapulohumeral muscular dystrophy. Neurology 1998;50:1791–1794. Jardine PE, Upadhyaya M, Maynard J, Harper P, Lunt PW: A scapular onset muscular dystrophy without facial involvement: Possible allelism with facioscapulohumeral muscular dystrophy. Neuromuscul Disord l994;4:477–482. Zatz M, Marie SK, Cerqueira A, Vainzof M, Pavanello RCM, Passos-Bueno MR: The facioscapulohumeral muscular dystrophy gene affects males more severely and more frequently than females. Am J Med Genet 1998;77:155–161. Kohler J, Rupilius B, Otto M, Bathke K, Koch MC: Germline mosaicism in 4q35 facioscapulohumeral muscular dystrophy (FSHD1A) occurring predominantly in oogenesis. Hum Genet 1996;98: 485–490. Winokur ST, Bengtsson U, Feddersen J, Mathews KD, Weiffenbach B, Bailey H, Markovich RP, Murray JC, Wasmuth JJ, Altherr MR, Schutte BC: The DNA rearrangement associated with facioscapulohumeral muscular dystrophy involves a heterochromatin-associated repetitive element: Implications for a role of chromatin structure in the pathogenesis of the disease. Chromosome Res 1994;2:225–234. Hewitt JE, Lyle R, Clark LN, Valleley EM, Wright TJ, Wijmenga C, van Deutekom JCT, Francis F, Sharpe PT, Hofker M, Frants RR, Williamson R: Analysis of the tandem repeat locus D4Z4 associated with facioscapulohumeral muscular dystrophy. Hum Mol Genet 1994;3:1287–1295. Ding H, Beckers M-C, Plaisance S, Marynen P, Collen D, Belayew A: Characterization of a double homeodomain protein (DUX1) encoded by a cDNA homologous to 3.3 kb dispersed repeated elements. Hum Mol Genet 1998;7:1681–1694. Brouwer OF, Padberg GW, Ruys CJM, Brand R, de Laat JAPM, Grote JJ: Hearing loss in facioscapulohumeral muscular dystrophy. Neurology 1991;41:1878–1881. Fitzsimons RB, Gurwin EB, Bird AC: Retinal vascular abnormalities in facioscapulohumeral muscular dystrophy. A general association with genetic and therapeutic associations. Brain 1987;110: 631–648. Padberg GW: Facioscapulohumeral muscular dystrophy; in Emery AEH (ed): Neuromuscular Disorders: Clinical and Molecular Genetics. Chichester, Wiley, 1998, pp 105–121. Twyman RS, Harper GD, Edgar MA: Thoracoscapular fusion in facioscapulohumeral dystrophy: Clinical review of a new surgical method. J Shoulder Elbow Surg 1996;5:201–205. Kissel JT, McDermott MP, Natarajan R, Mendell JR, Pandya S, King WM, Griggs RC, Tawil R, FSH- DY group: Pilot trial of albuterol in facioscapulohumeral muscular dystrophy. Neurology 1998;50:1402–1406. Padberg GW, Lunt PW, Koch M, Fardeau M: Facioscapulohumeral muscular dystrophy; in Emery AEH (ed): Diagnostic Criteria for Neuromuscular Disorders, ed 2. London, Royal Society of Medicine Press, 1997, chap 3, pp 9–15. Lunt PW, Compston DAS, Harper PS: Estimation of age-dependent penetrance in facioscapulohumeral muscular dystrophy by minimising ascertainment bias. J Med Genet l989;26:755– 760. Awerbuch GI, Nigro MA, Wishnow R: Beevor’s sign and facioscapulohumeral dystrophy. Arch Neurol 1990;47:1208–1209. Yamamoto K, Ikeda S, Hanyu N, Takeda S, Yanagisawa N: A pedigree analysis with minimised ascertainment bias shows anticipation in Met30-transthyretin related familial amyloid polyneuropathy J Med Genet 1998;35:23–30. Deidda G, Cacurri S, Piazzo N, Felicetti L: Direct detection of 4q35 rearrangements implicated in facioscapulohumeral muscular dystrophy (FSHD). J Med Genet 1996;33:361–365. Lemmers RJLF, van der Maarel S, van Deutekom JCT, van der Wielen MJR, Deidda G, Dauwerse HG, Hewitt J, Hofker M, Bakker E, Padberg G, Frants RR: Inter- and intrachromosomal subtelomeric rearrangements on 4q35: Implications for facioscapulohumeral muscular dystrophy (FSHD) aetiology and diagnosis. Hum Mol Genet 1998;7:1207–1214.


59

29

30

31

32 33 34

35

36

37

38

39 40

41

42

43

Tupler R, Berardinelli A, Barbierato L, Frants R, Hewitt JE, Lanzi G, Maraschio P, Tiepolo L: Monosomy of distal 4q does not cause facioscapulohumeral muscular dystrophy. J Med Genet 1996;33:366–370. van der Maarel SM, Deidda G, Lemmers RJLF, Bakker E, van der Wielen MJR, Sandkuijl L, Hewitt JE, Padberg GW, Frants RR: A new dosage test for subtelomeric 4;10 translocations improves conventional facioscapulohumeral muscular dystrophy (FSHD) diagnosis. J Med Genet 1999;36: 823–828. Lunt PW, Jardine PE, Stevenson A, Tyfield L: Genetic counselling in facioscapulohumeral muscular dystrophy (FSHD): Lessons from ‘clinically unaffected’ mutation carriers. Muscle Nerve 1998 (suppl 7):S25. Padberg GW, Vogels OJM, van der Kooi EL: The clinical picture of FSHD. Muscle Nerve 1998 (suppl 7):S25. Tawil R, McDermott MP, Pandya S, King W, Kissel J, Mendell JR, Griggs RC, FSH-DY group: A pilot trial of prednisone in facioscapulohumeral muscular dystrophy. Neurology 1997;48:46–49. Gilbert JR, Stajich JM, Wall S, Carter SC, Qiu H, Vance JM, Stewart CS, Speer MC, Pufky J, Yamaoka LH, Rozear M, Samson F, Fardeau M, Roses AD, Pericak-Vance MA: Evidence for heterogeneity in facioscapulohumeral muscular dystrophy (FSHD). Am J Hum Genet 1993;53: 401–408. Gilbert JR, Speer MC, Stajich J, Clancy R, Lewis K, Qiu H, Yamaoka L, Kumar A, Vance J, Stewart C, Rozear M, Roses AD, Pericak-Vance MA: Exclusion mapping of chromosomal regions which cross hybridise to FSHD1A associated markers in FSHD1B. J Med Genet 1995;32:770–773. van Deutekom JCT, Lemmers RJLF, Grewal PK, van Geel M, Romberg S, Dauwerse HG, Wright T, Padberg GW, Hofker MH, Hewitt JE, Frants RR: Identification of the first gene (FRG1) from the FSHD region on human chromosome 4q35. Hum Mol Genet 1996;5:581–590. Bouju S, Pietu G, Le Cunff M, Cros N, Malzac P, Pellissier J-F, Pons F, Leger J-J, Auffray C, Dechesne CA: Exclusion of muscle specific actinin-associated LIM protein (ALP) gene from 4q35 facioscapulohumeral muscular dystrophy (FSHD) candidate genes. Neuromuscul Disord 1999;9: 3–10. van Geel M, Beck AF, Eichler EE, Frants RR, de Jong PJ: Possible activation of a b-tubulin pseudogene (TUB4q) could explain the FSHD molecular defect. Am J Hum Genet 1999;65(suppl 4): A93. Harris S, Moncrieff C, Johnson K: Myotonic dystrophy: Will the real gene please step forward! Hum Mol Genet 1996;5:1417–1423. Richard I, Broux O, Allamand V, Fougerousse F, Chiannilkulchai N, Bourg N, Brenguier L, Devaud C, Pasturaud P, Roudaut C, Hillaire D, Passos-Bueno M-R, Zatz M, Tischfield JA, Fardeau M, Jackson CE, Cohen D, Beckmann JS: Mutations in the proteolytic enzyme calpain 3 cause limbgirdle muscular dystrophy type 2A. Cell 1995;81:27–40. Winokur ST, Bengtsson U, Vargas JC, Wasmuth JJ, Altherr MR: The evolutionary distribution and structural organization of the homeobox-containing repeat D4Z4 indicates a functional role for the ancestral copy in the FSHD region. Hum Mol Genet 1996;5:1567–1575 & corrigendum: Hum Mol Genet 1997;6:502. Grewal PK, van Deutekom JC, Mills KA, Lemmers RJ, Mathews KD, Frants RR, Hewitt JE: The mouse homolog of FRG1, a candidate gene for FSHD, maps proximal to the myodystrophy mutation on chromosome 8. Mamm Genome 1997;8:394–398. van der Maarel SM, Deidda G, Lemmers RJLF, van Overveld PGM, van der Wielen M, Hewitt JE, Sandkuijl L, Bakker B, van Ommen GJB, Padberg GW, Frants RR: De novo facioscapulohumeral muscular dystrophy: Frequent somatic mosaicism, sex-dependent phenotype, and the role of mitotic transchromosomal repeat interaction between chromosomes 4 and 10. Am J Hum Genet 2000;66: 26–35.

Dr. Peter W. Lunt, Clinical Genetics Unit, Bristol Royal Hospital for Sick Children, St. Michael’ Hill, Bristol BS2 8BJ (UK) Fax +44 117 9285167

Lunt

60


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

Myotonic Dystrophy Richard T. Moxley a, b, Giovanni Meola c, d a

University of Rochester School of Medicine and University of Rochester Medical Center, Rochester, N.Y., USA; c University of Milan and d San Donato Hospital, Milan, Italy b

The Trinucleotide Expansion [CTG]n in Myotonic Dystrophy: Impact of This Discovery on Diagnosis and Genetic Counseling Myotonic dystrophy (DM) is an autosomal dominant disorder with highly variable clinical manifestations [1, 2]. It affects specific tissues, such as distal limb and facial muscles, smooth muscles (gastrointestinal tract, uterus), the heart (primarily, the conducting system), the eye (primarily, the lens), the brain (especially, anterior temporal and frontal lobes), and endocrine function (testosterone deficiency, abnormal growth hormone regulation, insulin resistance) [1, 2]. DM results from an unstable trinucleotide repeat expansion containing cytosine, thymine and guanine [CTG]n, located in the 3 untranslated region of chromosome 19q13.3 [3–5]. The DM gene codes for a serine-threonine kinase (DMPK) [3–5]. This type of enzyme often regulates various intracellular processes by phosphorylation-dephosphorylation reactions [3–6]. The function of DMPK in normals and in DM is unknown. A deficiency of DMPK probably plays a pathophysiologic role in DM [6, 7], but other factors, such as an altered function of flanking genes [8–13] and toxic effects of the abnormally enlarged DMPK mRNA [12–19], are likely to contribute in a critical way to the variable multisystem manifestations of DM. Diagnosis The gold standard for the diagnosis of DM is the identification of an abnormally enlarged [CTG]n repeat expansion in the DM gene. Normal alleles for the DM gene range in size from [CTG]5 to [CTG]37 [3–5, 20]. DM patients

Table 1. Overview of genetics of DM Frequency

Inheritance Sex bias for transmission of the severe form Protein/expression Disease-causing mechanism Repeat/location Size of repeat in normal alleles Predisposing normal alleles (frequency) Linkage disequilibrium

1/8,000 (likely to be a significant underestimate; new calculations in the future will use data obtained by using specific gene probes) Autosomal dominant with anticipation Female (almost all cases of congenital DM result from female transmission of the DM allele) Serine/threonine kinase Unknown speculation (see text) CTG/3 untranslated region (CTG)5–37 Predisposed normal alleles have repeat size of (CTG)19–30 (approximately 0.10) Yes (absolute)

Table 2. Correlations of clinical manifestations with the age of onset Mildest [CTG]n =150 repeats Onset: middle to old age Major findings: cataracts, minimal or no muscle weakness Classical [CTG]n 300–1,000 repeats Onset: adolescence and early adult life Major findings: myotonia, muscle weakness (face, distal muscles in arms and legs) Congenital [CTG]n ?1,500 repeats with some cases occurring with sizes as small as 800 repeats Onset: at birth, frequent history of hydramnios and reduced fetal movements Major findings: neonatal respiratory distress, hypotonia, bilateral facial weakness, feeding difficulty, talipes, mental retardation

have repeat expansions ranging from 50 to over 2,000 [20–47]. Table 1 provides an overview of the genetics in DM. The specific alteration that leads to the unstable expansion of [CTG]n repeat expansion in DM remains a mystery. There is a general correlation between the degree of repeat expansion and the severity of the manifestations of DM [17–47]. However, a recent report, evaluating the relationship between leukocyte [CTG]n repeat length and clinical onset, has found that a significant correlation for age of onset only occurs in patients with relatively small repeat expansions [29]. These observations point out that the proposed classification of DM using age of onset, clinical severity and the length of the [CTG]n repeat expansion given in table 2 below is an approximation and not a rigorously proven classification.

Moxley/Meola

62

Diagnostic tests of primary value in DM include (1) DNA testing for the abnormal enlargement of the [CTG]n repeat, (2) a focused clinical examination to look for the skeletal muscle and nonmuscular manifestations, (3) electromyography to identify subclinical myotonia, and (4) slit lamp examination to detect characteristic cataracts [1, 2]. Tests of secondary value include (1) serum creatine kinase, which is often mildly elevated, and (2) muscle biopsy, which frequently shows an increase in central nuclei, type I fiber atrophy, and ringed fibers [1, 2]. The major challenge in diagnosis is to consider the possibility of DM. DM is so variable. It disguises itself in many ways. It may present as a floppy infant to a neonatalogist, a patient with failed labor or placenta previa to the obstetrician; a case of postoperative apnea to the anesthesiologist or surgeon, a patient with sudden heart block or tachyarrhythmias to the cardiologist, a patient with pseudoobstruction of the intestine to the gastroenterologist, a patient with recurrent atelectatic pneumonitis or sleep apnea to the pulmonologist, a baby with talipes to the orthopedist, a child with behavior disorder to the psychologist, and a young or old adult with cataracts to the ophthalmologist [1, 2]. Other associated clinical manifestations of DM may elude detection by each of these specialists because they have focused on ‘their own area’. Genetic Counseling Analysis of DNA isolated from amniocytes or chorionic villus samples is able to predict the delivery of infants with DM [17, 18, 21–25]. However, there are certain limitations. The [CTG]n repeat enlargement varies in the DM alleles in different tissues [12, 13, 23, 26, 30, 33–36, 38–47]. On occasion the [CTG]n repeat expansion in the DM gene isolated from amniocytes is smaller than the repeat isolated from maternal or fetal leukocyte DNA and yet the child ultimately turns out to have the severe form of disease, congenital DM [17]. Prenatal diagnosis of congenital DM, as opposed to the less severe types of DM, needs to rely on a combination of factors, including maternal pregnancy history, clinical findings, and cautious interpretation of maternal and fetal DNA analysis [17]. Clearly DNA analysis permits identification of whether the fetus has the abnormally expanded DM allele and proves that the disease allele is present. It is not, however, a reliable means to predict the severity of DM after birth. At the time of prenatal evaluation of at risk mothers and infants it is useful to discuss the different obstetric complications so that optimal use of preventive therapy is available [1, 2]. Counseling of women with DM is important so that they have an understanding of the increased risk of complications during pregnancy and delivery and are aware of the fact that the degree of [CTG]n repeat expansion may not reliably predict problems [19].

Myotonic Dystrophy

63

Fig. 1. Serial photographs of a father and son affected with DM are shown. To the right are two photographs in the upper panel of the father (age 77 years) and two in the lower panel of the son (age 41 years) obtained in 1991. The two photographs furthest on the left were obtained when the father was 45 years of age and the son was 18 years old. Those photographs immediately adjacent to the ones obtained in 1991 show the father at age 62 years and the son at age 26 years. Both patients have had cataract surgery, right eye lens implant for the father and left eye for the son. Ptosis, bifacial weakness, wasting of the sternocleidomastoid, temporalis, and masseter muscles as well as frontal balding are apparent in the son and less obvious in the father.

The DM Gene and the Phenomenon of Anticipation The phenomenon of anticipation, the earlier onset of more severe symptoms in individuals in successive generations within a family, occurs in DM [1]. The discovery of the unstable CTG repeat expansion in the DM gene provides a genetic explanation for this clinical phenomenon. Figure 1 illustrates anticipation between a father and son. The father has mild DM and his son has earlier onset classical type of DM (table 2).

Moxley/Meola

64

Fig. 2. Mother with mild to moderate DM with her two children, both of whom have the congenital form of DM.

Figure 2 presents a mother with mild DM and her two daughters who have congenital DM. Figure 2 is also a reminder that almost all cases of congenital DM occur in offspring of mothers with DM [1, 17, 21–25, 27, 31]. There is a genetic explanation for the relatively exclusive predilection of females with myotonic dystrophy to have offspring with the congenital form of DM. Oocytes remain viable in DM mothers despite having CTG repeat expansions up to several thousand repeats in length. Males have some mechanism that provides an upper limit in repeat size for sperm, such that the male gametes with hugely enlarged CTG repeats do not survive or are not capable of producing a viable pregnancy [31]. In contrast to the female preponderance that accounts for the congenital form of DM, there is an excess of male grandparental transmissions in DM [27, 31]. In particular, there is an excess of males among those first documented in a pedigree to have transmitted DM to individuals who have clear clinical signs of disease [27, 31]. A similar male predilection to transmit unstable repeat expansions occurs in Huntington’s disease and suggests a possible similarity between DM and Huntington’s related to the influence of male gender on repeat instability [31]. Anticipation occurs because of the predilection of the CTG repeat in the DM gene to enlarge. However, there are examples of the opposite phenomenon, that is the contraction in CTG repeat length in successive generations [20, 28, 33]. The mechanism responsible for the maintenance and regulation of CTG repeat size in the DM gene is unknown. As our knowledge of this process increases, a new, perhaps curative, strategy for treatment may emerge.

Myotonic Dystrophy

65

[CTG]n Repeat Mosaicism in DM: Its Impact on Clinical Features and Treatment Marked variability in the clinical presentation and in the severity of the different manifestations are hallmarks of DM [1, 2]. Certain organs have an increased incidence of symptoms in DM [1, 2]. DNA analysis has demonstrated somatic mosaicism affecting these organs. There is an increased length of the CTG repeat expansion (greater than that observed in leukocyte DNA) in skeletal muscle [30, 33–42], brain [34, 40–42], and heart [34, 40–42]. These findings and other related observations have led to the hypothesis that the severity of alterations in the genotype (the degree of abnormal CTG repeat expansion in the DM gene) predicts the severity of clinical manifestations in the specific tissue under study. There are several reports that support this hypothesis and describe an increased severity of skeletal muscle manifestations (phenotype) that directly correlates with an increase in CTG repeat length measured in leukocyte DNA [3–5, 21, 44–47]. These observations offer encouragement to clinicians and researchers that there may be, at least in some patients, a straightforward, direct relationship between genotype and phenotype. However, the results of recent investigations raise important questions about the reliability of predicting phenotype from genotype. These reports indicate a need for caution in individual patients before concluding that they show a direct relationship between the CTG repeat expansion and severity. For example, there is often a much greater enlargement of CTG repeat expansion in skeletal muscle DNA compared to that in circulating leukocytes [30, 33–42]. This agrees with the clinical observation that skeletal muscle dysfunction is a dominant feature in DM and that abnormal function of leukocytes is uncommon. However, specific muscle groups waste to a greater degree in DM patients. The muscles of the face and of the distal limbs waste much more than the proximal limb muscles [1, 2]. Recently, investigators have examined muscle biopsy specimens obtained from different muscles in the same patient to determine if the extent of CTG repeat expansion in specific muscles predicts their degree of weakness [36]. The investigators have found that the degree of abnormal CTG repeat expansion in the vastus lateralis (proximal) and tibialis anterior (distal) muscles is similar, but the degree of muscle weakness is much greater in the tibialis anterior compared to the quadriceps femoris muscles [36]. More study is necessary to identify those factors, other than CTG repeat expansion, that account for the selective pattern of muscle wasting and weakness. Cognitive and behavioral abnormalities, as well as alterations on imaging of the brain (ventricular enlargement, white matter changes in temporal lobes)

Moxley/Meola

66

occur in DM [1, 2]. Recent investigations indicate that there is a lack of direct correlation between the degree of CTG repeat enlargement in leukocyte DNA and (1) the severity of cognitive/behavioral impairment in adults with DM [48], (2) the alterations that occur in brain structure [49–52], and (3) the mental retardation that is characteristic in congenital DM [50]. Brain manifestations attributable to DM can sometimes occur years before the development of skeletal muscle weakness [50]. These examples of alterations in brain function and structure provide further evidence of the restricted manifestation of symptoms that can occur in DM and point out the tenuous nature of the hypothesis that there is a straight forward, direct correlation between CTG repeat expansion in leukocyte DNA and the various manifestations of DM in different tissues. The pathomechanism responsible for the brain manifestations of DM may differ from that in other target tissues. There is a decrease in cerebral blood flow in the anterior temporal and frontal lobes of patients with DM [48]. Metabolic factors and alterations in the control of cerebral blood flow may play a role in the development of brain manifestations in adult DM [48]. Cardiac conduction defects are common in DM and are a major cause for sudden death in patients [1, 2, 53–58]. However, there is no direct relationship between the degree of CTG repeat enlargement in leukocyte DNA and atrial or ventricular arrhythmias [53–55], between cardiac conduction disturbance and skeletal muscle involvement [53], or between cardiac conduction disturbance and the extent of fibrolipomatous infiltration and cardiac dysfunction [58]. The lack of a straightforward correlation between CTG repeat length in leukocyte DNA and cardiac symptoms may relate to the somatic mosaicism of repeat expansion and indicate greater expansion of the CTG repeat in fibers of the Purkinje system than in circulating leukocytes. Further investigations of postmortem cardiac tissue and of cardiac function are necessary to clarify the relationship between the severity of cardiac manifestations of DM and the gene defect. Studies of human fetal development in DM demonstrate that the CTG repeat instability develops only after 13 weeks’ gestational age and before 16 weeks [40]. This observation suggests that CTG repeat size in different tissues is significantly influenced by developmental genes at specific times in fetal growth. It further suggests that repeat expansion and somatic mosaicism are largely complete by a gestational age of 16 weeks. However, CTG repeat expansion occurs in leucocyte DNA [38, 52] and sperm [38] during adult life. These observations raise the possibility that the DM gene may expand further during adult life in tissues, such as skeletal muscle. This ongoing additional expansion of the DM gene may influence flanking genes and/or increase the accumulation of DMPK mRNA in a way that triggers cellular dysfunction

Myotonic Dystrophy

67

or cell death. These ideas and other theories about the genetic pathomechanism in DM are under investigation. Clinical Features and Management Leukocyte DNA analysis provides an earlier opportunity to diagnose [20] and to anticipate clinical problems in patients with DM. It is now possible to pursue a more aggressive monitoring program to detect early cardiac conduction disturbance, cataract formation and respiratory difficulties [1, 2]. Table 3 presents a summary of the major problems and their treatment in patients with congenital and childhood onset DM. Table 4 outlines the problems and management of complaints related to skeletal muscle dysfunction in DM. Females with DM can encounter significant complications during pregnancy and have an increased frequency of problems at the time of delivery. These are summarized in table 5. Patients with DM are at risk for developing intraoperative and postoperative complications of anesthesia, especially the development of delayed onset apnea [1, 2]. Table 6 outlines strategies to identify and/ or to avoid complications from anesthesia.

Pathophysiology and Therapeutic Trials Natural History: Opportunity to Establish New Measures DNA diagnostic testing can establish the diagnosis of DM in individuals with only minimal as well as those with advanced symptoms of DM. This capability provides us with a new opportunity to establish the natural history of all of the various clinical manifestations of DM. The development of standardized, reliable, reproducible testing to assess the various clinical manifestations of DM is in progress [38]. Table 7 provides examples of possible clinical tests that may be useful in documenting the natural history of these various manifestations. Trials of Testosterone, Growth Hormone, Insulin-Like Growth Factor-1, Troglitazone and Dehydroepiandrosterone Sulfate The pathomechanism responsible for the muscle wasting in DM remains unclear. Possible causes include toxic DMPK mRNA-mediated muscle fiber death [12, 13–16, 38], DMPK deficiency [6, 10, 12, 13], abnormal function of genes that flank the DM gene [8–13], failed muscle fiber growth [59], insulin resistance [2, 60, 61], and deficient protein anabolism [2, 60, 62, 63]. Gonadal failure and a deficiency of testosterone are common in men with DM [1, 2]. To determine if maintaining a high physiologic level of testosterone increases muscle mass and function in DM, investigators have given either

Moxley/Meola

68

Table 3. Management of DM in infancy and childhood [adapted from 1] Problem

Management

Respiratory insufficiency or failure

Neonatal intensive care, ventilator support, chest x-ray to check on elevation of diaphragm, poor prognosis for survival if on ventilator longer than 4 weeks; standard care as necessary for pulmonary immaturity

Feeding problems and aspiration

Frequent monitoring of respiration and check chest x-ray; poor swallowing monitor for recurrent episodes of aspiration; neonatal intensive care or highly skilled nursing care needed; evaluate esophageal function if problems persist

Risk of anoxia and cerebral hemorrhage in infants

Neonatal intensive care; serial cerebral ultrasound measurements are helpful

Talipes

Early diagnosis and active correction; splinting; surgery, if indicated

General delay in development

Accurate evaluation of contributing factors: especially, muscular and cerebral components

Speech problems

Anticipate early: distinguish palatal weakness and other muscular defects from mental retardation

Abdominal pain

A frequent and difficult problem; treat as for ‘spastic colon’; role of smooth muscle disease and myotonia in this complaint is unknown; response to medications is variable and not established

Surgery and anesthesia

Be aware of risks; avoid general anesthesia if possible

Recurrent otitis, hearing difficulty

Active management of otitis, awareness of later neural hearing loss

School decisions

May need special classroom setting and vocational education in later school years; requires full assessment of functional capacities, awareness of facial immobility and hearing and speech defects which exaggerate mental retardation; physical disability rarely severe and usually is static during childhood

Should physical activity be restricted

Not feasible or justified; obtain yearly ECG to search for signs of cardiac conduction disturbance

Myotonic Dystrophy

69

Table 4. Treatment of skeletal muscle problems in DM Problem

Management

Foot drop

Light-weight molded ankle foot orthoses

Knee extensor weakness

Light-weight long leg knee-ankle-foot orthoses may allow prolonged weight bearing and prevent the pain and chronic effusion in the knee joint

Low back strain/pain

Often occurs when patients have to arise from sitting with their legs abducted their thorax flexed forward, and their back in exaggerated lordosis; PT to advise about elevated seats, self-help devices to assist in arising and reducing low back strain; use NSAIDs and low-dose tricyclic antidepressants for several weeks or as maintenance therapy; use cyclobenzaprine for several days during acute severe pain; mexiletine treatment to reduce myotonia in paraspinous muscles may be necessary on a chronic basis to decrease the susceptibility of these muscles to recurrent strain/pain

Anterior shoulder pain

Develops, as scapular fixator weakness becomes apparent; rotator cuff strain, tendonitis, and overuse spasm in the anterior deltoid and upper trapezius muscles contribute individually or in combination; need to avoid lifting with arms outstretched and abducted; PT instruction is helpful; for acute pain use rest, NSAIDs, low dose tricyclic antidepressants, and cyclobenzaprine; long-term mexiletine therapy may also be necessary for chronic or recurrent shoulder muscle pain; obtain orthopedic consultation if rotator cuff injury is suspected

Myotonia

Mexiletine is often more effective than phenytoin

Generalized weakness

Wheelchair; use intermittently with braces in earlier stages of generalized weakness (e.g. for traveling long distances within the home or trips outside)

Respiratory muscle weakness with ventilatory failure

Consider nasal ventilation, initially at night and subsequently during daytime; in selected cases, consider positive pressure ventilation via tracheostomy

Neck weakness

Fitted collar; head supports in car seats and chairs

Ptosis

Lid elevating crutches or props on eyeglasses; lid surgery is necessary only infrequently

Recurrent dislocation of the mandible due to weakness of temporalis and masseter muscles

Mandible surgery is infrequently required; guidelines for preoperative and postoperative care need to be followed carefully; often there is muscle spasm in the masseter muscles during attacks of pain with dislocation; local heat packs and mexiletine therapy on a long-term basis can lessen the severity of the episodes; occasionally an attack of mouth closure with dislocation of the mandible creates a medical emergency for ‘mouth breathers’; patients on critically timed oral medication

Moxley/Meola

70

Table 5. Maternal-obstetric complications in DM Maternal complications Increased muscle weakness, especially respiratory Increased rate of spontaneous abortion Reduced fetal movements and hydramnios Prolonged and often ineffective labor Sensitivity to general anesthesia (arrhythmias, apnea following cesarean section) Obstetric complications Retained placenta Placenta previa Neonatal deaths

Table 6. Preoperative and postoperative care and complications of anesthesia in DM Preoperative evaluation ECG, pulmonary function testing (including supine and upright forced vital capacity) and arterial blood gas measurements Intraoperative monitoring Monitor ECG; measure arterial blood pressure; use a peripheral nerve stimulator to monitor blockade of peripheral muscle; monitor temperature; warm mattress; warm intravenous fluids; maintain humidification of anesthetic gases Postoperative care Retain endotracheal tube in place and ventilate if necessary on an intensive care unit; monitor respiratory efficiency with checks of oxygen saturation and pCO2 for at least 24 h postoperatively to avoid overlooking delayed onset apnea; use controlled flow oxygen therapy with close monitoring of ventilation in patients relying upon hypoxic drive due to chronic respiratory insufficiency; provide early physiotherapy; monitor ECG; keep patient warm; monitor swallowing closely to check for signs of aspiration; treat all infections vigorously Anesthetic agents When possible use local or regional anesthesia, such as, an epidural block; avoid suxamethonium and other depolarizing muscle relaxants; avoid or use only minimal doses of thiopental; for muscle relaxation use short-acting agents, such as atracurium or vecuronium; avoid or use only minimal doses of opiates to avoid respiratory depression; when possible, avoid general anesthesia; if necessary, use combination of nitrous oxide/oxygen mixture with an agent, such as 0.8% enflurane or 1.0% isoflurane; use anticholinesterases, such as neostigmine, with care; may be preferable to ventilate the patient until residual curarization wears off

Myotonic Dystrophy

71

Table 7. Different measures to define the natural history of DM Skeletal muscle weakness (manual muscle testing, quantitative isometric muscle contraction force testing) and wasting (dual energy x-ray absorptiometry, total body potassium, magnetic resonance imaging, urinary creatinine) Myotonia (electromechanical measures, in vitro techniques) CNS (neuropsychological tests, imaging, position emission tomography) Respiratory (forced vital capacity, sleep apnea testing, electromyography, ventilatory drive) Cardiac (electrocardiogram, 24 h monitor, echocardiography, magnetic resonance imaging, position emission tomography) G1 (cinevideoswallowing, gastric emptying time) Endocrine (gonadal, pituitary axis, thyroid, pancreas, hormone levels, hormone challenges, oral glucose tolerance testing, intravenous glucose tolerance testing)

testosterone or placebo in a randomized controlled fashion to 40 men with DM [64]. Active treatment involved testosterone enanthate (3 mg/kg per week) for 12 months. After 1 year those patients who received testosterone had a significant increase in muscle mass. However, they had no significant improvement in strength or function [64]. There is a definite role for testosterone in providing replacement therapy in DM for testosterone deficiency. The use of testosterone as a treatment for muscle wasting requires further study. In addition to the occurrence of testosterone deficiency, many DM patients have an abnormal release and regulation of growth hormone [1, 2]. A therapeutic trial of growth hormone isolated from pooled human brains [65] raised hope that it might be a useful treatment. Daily intramuscular injections for 2 weeks led to an improvement in nitrogen balance and a shortening of the abnormally prolonged PR interval that was present at baseline [65]. However, the patients had no increase in muscle strength or function. A more recent trial using recombinant human growth hormone given daily for 16 weeks to 7 men with classical DM produced a 10% increase in lean body mass and a 28% increase in muscle protein synthesis [66]. Despite this improvement in protein anabolism there was no significant increase in strength or function. These studies of growth hormone and testosterone treatment suggest that future longer-term therapeutic trials may prove beneficial since both growth hormone and testosterone produced an increase in muscle mass. However, it is unclear whether the increase in muscle tissue actually represented an increase in the contractile proteins. Further studies are needed.

Moxley/Meola

72

In recent years human recombinant DNA-derived insulin-like growth factor-1 (IGF-1) has become available. To determine if IGF-1 is more effective than its ‘parent’, growth hormone, investigators have recently completed a therapeutic trial of IGF-1 in DM [67]. Seven men and 2 women received a 4-month trial of 5 mg of human recombinant IGF-1 subcutaneously at 12hour intervals. Four patients, whose dosage of IGF-1 was greater than 0.070 mg/ kg per dose, showed an improvement both in strength and in function. Patients also showed improvements in the rate of protein synthesis and in insulin sensitivity. The results of this trial of IGF-1 are promising. Future trials merit consideration. However, there are restraining circumstances. The cost of IGF-1 (as well as growth hormone) is very high. There is also a risk of accelerating the growth of a coexisting neoplasm or of worsening prostatic hypertrophy with IGF-1 therapy. Insulin is an essential anabolic hormone and skeletal muscle resistance to insulin occurs in DM [2, 60]. A loss of the usual anabolic actions of insulin may contribute to the muscle wasting and weakness in DM [60]. Therapeutic trials to reverse skeletal muscle insulin resistance are in progress. One trial is using the newly marketed thiazolidinedione derivative, troglitazone [38, 68]. A recent case report indicates that troglitazone has reduced insulin resistance and decreased myotonia in a patient with DM [69]. This forecasts a beneficial result of the trial of troglitazone in DM. The results of the troglitazone study that is in progress will help to establish if insulin-enhancing agents, like thiazolidinediones, are useful in DM. A previous study has identified a decrease in circulating levels of dehydroepiandrosterone (DHEAS) in patients with DM [70]. This finding has raised the possibility that a deficiency of this adrenal hormone may contribute to the muscle wasting and weakness. To investigate this possibility researchers have initiated therapeutic trials using daily intravenous infusions of 200 mg of DHEAS. They have found very encouraging results in their initial open trials [71, 72]. Patients have shown increased muscle function and decreased myotonia. Further controlled studies are in progress. Trials of Antimyotonia Therapy The pathomechanism that underlies the myotonia in DM is unclear. Some evidence suggests that it involves apamin-sensitive potassium channels [38, 73, 74]. However, previous reports have also described alterations in sodium conductance [38, 75]. No explanation is available to account for the absence of myotonia during the first several years of life in patients with congenital DM, and no explanation is available to account for the gradual appearance of clinical and electrical myotonia that occurs later in childhood [1, 2].

Myotonic Dystrophy

73

Only a few small scale controlled studies that evaluate antimyotonia therapy are available in DM [75–79]. In one investigation 7 men and 3 women participated in a randomized, blinded, crossover study [78]. Each received a 2-week treatment of procainamide (250 mg every 6 h for 7 days and then 500 mg every 6 h for the 2nd week) or disopyramide (100 mg every 8 h for the 1st week then 200 mg every 8 h for the 2nd week). Both medications decreased myotonia but did not produce a change in grip strength. Five patients developed side effects with procainamide and 6 patients had side effects with disopyramide. Another trial of antimyotonia treatment involved 6 patients in a complicated double-blind study design [77]. The study involved 8 separate 15-day treatment periods: 2 periods of phenytoin therapy (1 of 200 mg per day, the other of 300 mg per day), and 2 periods of treatment with carbamazepine (1 using 600 mg per day, the other using 800 mg per day). Each of the above 15-day trial periods had a preceding 15-day trial of placebo treatment. Both phenytoin and carbamazepine reduced myotonic stiffness. Patients tolerated both medications well. Another trial involved 6 men and 4 women treated in an open fashion with tocainide [76]. Each patient received 400 mg a day followed by increases in increments of 400 mg a day to a total dose of 1,200 mg daily. After 2 weeks of treatment the patients had a 36–80% reduction in myotonic stiffness. No significant change in their electrocardiograms occurred. Three of the 10 patients experienced transient dizziness and nausea which disappeared following a reduction in the dosage of tocainide to 800 mg a day. The most encouraging trial of antimyotonia therapy has demonstrated a superior efficacy of mexiletine and tocainide compared to phenytoin and disopyramide in 9 patients with DM [75]. Daily doses of 400–600 mg of mexiletine produced a maximum decrease in myotonic stiffness equal to that of 1,200 mg of tocainide and produced a significantly better reduction of myotonia than that achieved with 300 mg of phenytoin daily. The investigators have recommended mexiletine treatment over tocainide in view of the increased risk of bone marrow suppression with tocainide. Future large scale trials of mexiletine in DM deserve consideration. Such trials will help to determine if mexiletine is safe and effective in reducing not only myotonic stiffness of skeletal muscle, but whether it can ameliorate the myotonia that interferes with swallowing and gastrointestinal function [1, 2].

Opportunities for Future Treatment: Studies Using Animal Models The development of useful animal models for DM remains a challenge. Mouse knockout models of the DM gene and overexpression models have not produced animals with disease manifestations that closely resemble human

Moxley/Meola

74

DM [12, 13, 38, 80]. However, more recent studies have raised hopes that abnormal CTG repeat expansions in the small and moderate range can be successfully transmitted in successive litters of mice and that the clinical and histopathologic alterations observed are similar to human DM [13, 38, 81, 82]. There are also hopes that knockout models in mice of genes that flank the DM locus will provide clues about the molecular pathomechanism of DM and about the role that flanking genes may have in some of the manifestations of the disease [8, 12, 13, 38]. There is optimism that in the near future new ideas will be forthcoming from animal studies, and that ultimately the investigations using animal models will lead to effective treatments for the various manifestations of DM in humans.

References 1 2 3

4 5 6

7

8

9

10 11 12

13 14 15

Harper PS: Myotonic Dystrophy. London, Saunders, 1989. Moxley RT: Myotonic muscular dystrophy; in Rowland LP, DiMauro S (eds): Handbook of Clinical Neurology. New York, Elsevier Science, 1992, pp 209–259. Brook JD, McCurrach ME, Harley HG, et al: Molecular basis of myotonic dystrophy: Expansion of a trinucleotide (CTG) repeat at the 3 end of a transcript encoding a protein kinase family member. Cell 1992;68:799–808. Fu YH, Pizzuti A, Fenwick RG Jr, et al: An unstable triplet repeat in a gene related to myotonic muscular dystrophy. Science 1992;255:1256–1258. Mahadevan M, Tsilfidis C, Sabourin L, et al: Myotonic dystrophy mutation: An unstable CTG repeat in the 3 untranslated region of the gene. Science 1992;255:1253–1255. Epstein HF, Jin S: Biochemical studies of DM protein kinase (DMPK); in Wells RD, Warren ST (eds): Genetic Instabilities and Hereditary Neurological Diseases. San Diego, Academic Press, 1998, pp 147–167. Hamshere MG, Newman EE, Alwazzan M, Athwal BS, Brook JD: Transcriptional abnormality in myotonic dystrophy affects DMPK but not neighbouring genes. Proc Natl Acad Sci USA 1997;94: 7394–7399. Winchester CL, Johnson K: Is myotonic dystrophy (DM) the result of a contiguous gene Defect?; in Wells RD, Warren ST (eds): Genetic Instabilities and Hereditary Neurological Diseases. San Diego, Academic Press, 1998, pp 169–179. Boucher CA, King SK, Carey N, et al: A novel homeodomain-encoding gene is associated with a large CpG island interrupted by the myotonic dystrophy unstable (CTG)n repeat. Hum Mol Genet 1995;4:1919–1925. Thornton CA, Wymer JP, Simmons Z, McClain C, Moxley RT: Expansion of the myotonic dystrophy CTG repeat reduces expression of the flanking DMAHP gene. Nat Genet 1997;16:407–409. Klesert TR, Otten AD, Bird TD, Tapscott SJ: Trinucleotide repeat expansion at the myotonic dystrophy locus reduces expression of DMAHP. Nat Genet 1997;16:402–406. Waring J, Korneluk R: Genetic studies of the myotonic dystrophy CTG repeat; in Wells RD, Warren ST (eds): Genetic Instabilities and Hereditary Neurological Diseases. San Diego, Academic Press, 1998, pp 131–146. Groenen P, Wieringa B: Expanding complexity in myotonic dystrophy. Bioessays 1998;20:901–912. Taneja KL, McCurrach M, Schalling M, Housman D, Singer RH: Foci of trinucleotide repeat transcripts in nuclei of myotonic dystrophy cells and tissues. J Cell Biol 1995;128:995–1002. Davis BM, McCurrach ME, Taneja KL, Singer RH, Housman DE: Expansion of a CUG trinucleotide repeat in the 3 untranslated region of myotonic dystrophy protein kinase transcripts results in nuclear retention of transcripts. Proc Natl Acad Sci USA 1997;94:7388–7393.

Myotonic Dystrophy

75

16

17 18 19

20 21 22 23 24 25 26

27

28 29

30 31

32 33

34 35 36

37 38

Timchenko LT, Timchenko NA, Caskey CT, Roberts R: Novel proteins with binding specificity for DNA CTG repeats and RNA CUG repeats: Implications for myotonic dystrophy. Hum Mol Genet 1996;5:115–121. Geifman-Holtzman O, Fay K: Prenatal diagnosis of congenital myotonic dystrophy and counseling of the pregnant mother: Case report and literature review. Am J Med Genet 1998;78:250–253. Ohya K, Tachi N, Sato T, Kon S, Kikuchi K, Chiba S: Detection of the CTG repeat expansion in congenital myotonic dystrophy. Jpn J Hum Genet 1997;42:169–180. Rudnik-Schoneborn S, Nicholson GA, Morgan G, Rohrig D, Zerres K: Different patterns of obstetric complications in myotonic dystrophy in relation to the disease status of the fetus. Am J Med Genet 1998;80:314–321. Shelbourne P, Davies J, Buxton J, et al: Direct diagnosis of myotonic dystrophy with a diseasespecific DNA marker. N Engl J Med 1993;328:471–475. Redman JB, Fenwick RJ, Fu YH, Pizzuti A, Caskey CT: Relationship between parental trinucleotide GCT repeat length and severity of myotonic dystrophy in offspring. JAMA 1993;269:1960–1965. Harley HG, Rundle SA, MacMillan JC, et al: Size of the unstable CTG repeat sequence in relation to phenotype and parental transmission in myotonic dystrophy. Am J Hum Genet 1993;52:1164–1174. Lavedan C, Hofmann-Radvanyi H, Shelbourne P, et al: Myotonic dystrophy: Size- and sex-dependent dynamics of CTG meiotic instability, and somatic mosaicism. Am J Hum Genet 1993;52:875–883. Tsilfidis C, MacKenzie AE, Mettler G, Barcelo J, Korneluk RG: Correlation between CTG trinucleotide repeat length and frequency of severe congenital myotonic dystrophy. Nat Genet 1992;1:192–195. Hunter A, Tsilfidis C, Mettler G, et al: The correlation of age of onset with CTG trinucleotide repeat amplification in myotonic dystrophy. J Med Genet 1992;29:774–779. Jansen G, Willems P, Coerwinkel M, et al: Gonosomal mosaicism in myotonic dystrophy patients: Involvement of mitotic events in (CTG)n repeat variation and selection against extreme expansion in sperm. Am J Hum Genet 1994;54:575–585. Brunner HG, Bruggenwirth HT, Nillesen W, et al: Influence of sex of the transmitting parent as well as of parental allele size on the CTG expansion in myotonic dystrophy (DM). Am J Hum Genet 1993;53:1016–1023. Ashizawa T, Anvret M, Baiget M, et al: Characteristics of intergenerational contractions of the CTG repeat in myotonic dystrophy. Am J Hum Genet 1994;54:414–423. Hamshere MG, Harley H, Harper P, Brook JD, Brookfield JF: Myotonic dystrophy: The correlation of (CTG) repeat length in leucocytes with age at onset is significant only for patients with small expansions. J Med Genet 1999;36:59–61. Ashizawa T, Dubel JR, Harati Y: Somatic instability of CTG repeat in myotonic dystrophy. Neurology 1993;43:2674–2678. Harper PS: Myotonic dystrophy as a trinucleotide repeat disorder – A clinical perspective; in Wells RD, Warren ST (eds): Genetic Instabilities and Hereditary Neurological Diseases. San Diego, Academic Press, 1998, pp 115–130. Matsumura R, Namikawa T, Miki T, et al: An intergenerational contraction of the CTG repeat in Japanese myotonic dystrophy. J Neurol Sci 1996;139:48–51. Anvret M, Ahlberg G, Grandell U, Hedberg B, Johnson K, Edstrom L: Larger expansions of the CTG repeat in muscle compared to lymphocytes from patients with myotonic dystrophy. Hum Mol Genet 1993;2:1397–1400. Thornton CA, Johnson K, Moxley RT: Myotonic dystrophy patients have larger CTG expansions in skeletal muscle than in leukocytes. Ann Neurol 1994;35:104–107. Ansved T, Edstrom L, Grandell U, Hedberg B, Anvret M: Variation of CTG-repeat number of the DMPK gene in muscle tissue. Neuromuscul Disord 1997;7:152–155. Hedberg B, Anvret M, Ansved T: CTG-repeat length in distal and proximal leg muscles of symptomatic and non-symptomatic patients with myotonic dystrophy: Relation to muscle strength and degree of histopathological abnormalities. Eur J Neurol 1999;6:341–346. Laurent A, Costa JM, Assouline B, Voyer M, Vidaud M: Myotonic dystrophy protein kinase gene expression in skeletal muscle from congenitally affected infants. Ann Genet 1997;40:169–174. Anonymous: AFM/MDA 1st International Myotonic Dystrophy Consortium Conference 30 June– 1 July 1997, Paris, France. Neuromuscul Disord 1998;8:432–437.

Moxley/Meola

76

39 40 41 42 43 44 45 46 47

48 49 50

51 52

53

54 55

56 57 58

59 60 61

62

Tachi N, Kozuka N, Ohya K, Chiba S, Kikuchi K: CTG repeat size and histologic findings of skeletal muscle from patients with congenital myotonic dystrophy. J Child Neurol 1996;11:430–432. Martorell L, Johnson K, Boucher CA, Baiget M: Somatic instability of the myotonic dystrophy (CTG)n repeat during human fetal development. Hum Mol Genet 1997;6:877–880. Joseph JT, Richards CS, Anthony DC, Upton M, Perez-Atayde AR, Greenstein P: Congenital myotonic dystrophy pathology and somatic mosaicism. Neurology 1997;49:1457–1460. Wong LJ, Ashizawa T: Instability of the (CTG)n repeat in congenital myotonic dystrophy (letter). Am J Hum Genet 1997;61:1445–1448. Ishii S, Nishio T, Sunohara N, et al: Small increase in triplet repeat length of cerebellum from patients with myotonic dystrophy. Hum Genet 1996;98:138–140. Gennarelli M, Novelli G, Andreasi BF, et al: Prediction of myotonic dystrophy clinical severity based on the number of intragenic [CTG]n trinucleotide repeats. Am J Med Genet 1996;65:342–347. Ashizawa T, Dubel JR, Dunne PW, et al: Anticipation in myotonic dystrophy. II. Complex relationships between clinical findings and structure of the GCT repeat. Neurology 1992;42:1877–1883. Jaspert A, Fahsold R, Grehl H, Claus D: Myotonic dystrophy: Correlation of clinical symptoms with the size of the CTG trinucleotide repeat. J Neurol 1995;242:99–104. Zatz M, Passos-Bueno MR, Cerqueira A, Marie SK, Vainzof M, Pavanello RC: Analysis of the CTG repeat in skeletal muscle of young and adult myotonic dystrophy patients: When does the expansion occur? Hum Mol Genet 1995;4:401–406. Meola G, Sansone V, Perani D, et al: Reduced cerebral blood flow and impaired visual-spatial function in proximal myotonic myopathy. Neurology 1999;53:1042–1050. Martinello F, Piazza A, Pastorello E, Angelini C, Trevisan CP: Clinical and neuroimaging study of central nervous system in congenital myotonic dystrophy. J Neurol 1999;246:186–192. Spranger M, Spranger S, Tischendorf M, Meinck HM, Cremer M: Myotonic dystrophy. The role of large triplet repeat length in the development of mental retardation. Arch Neurol 1997;54: 251–254. Akiguchi I, Nakano S, Shiino A, et al: Brain proton magnetic resonance spectroscopy and brain atrophy in myotonic dystrophy. Arch Neurol 1999;56:325–330. Martorell L, Martinez JM, Carey N, Johnson K, Baiget M: Comparison of CTG repeat length expansion and clinical progression of myotonic dystrophy over a five year period. J Med Genet 1995;32:593–596. Lazarus A, Varin J, Ounnoughene Z, et al: Relationships among electrophysiological findings and clinical status, heart function, and extent of DNA mutation in myotonic dystrophy. Circulation 1999;99:1041–1046. Phillips MF, Harper PS: Cardiac disease in myotonic dystrophy. Cardiovasc Res 1997;33:13–22. Tokgozoglu LS, Ashizawa T, Pacifico A, Armstrong RM, Epstein HF, Zoghbi WA: Cardiac involvement in a large kindred with myotonic dystrophy. Quantitative assessment and relation to size of CTG repeat expansion. JAMA 1995;274:813–819. Annane D, Merlet P, Radvanyi H, et al: Blunted coronary reserve in myotonic dystrophy. An early and gene-related phenomenon. Circulation 1996;94:973–977. Hayashi Y, Ikeda U, Kojo T, et al: Cardiac abnormalities and cytosine-thymine-guanine trinucleotide repeats in myotonic dystrophy. Am Heart J 1997;134:292–297. De Ambroggi L, Raisaro A, Marchiano V, Radice S, Meola G: Cardiac involvement in patients with myotonic dystrophy: Characteristic features of magnetic resonance imaging. Eur Heart J 1995; 16:1007–1010. Carpenter S, Karpati G: Pathology of Skeletal Muscle. New York, Churchill Livingstone, 1984, pp 616–631. Livingston JN, Moxley RT 3d: Myotonic dystrophy: Phenotype-genotype and insulin resistance. Diabetes 1994;2:29–42. Morrone A, Pegoraro E, Angelini C, Zammarchi E, Marconi G, Hoffman EP: RNA metabolism in myotonic dystrophy: Patient muscle shows decreased insulin receptor RNA and protein consistent with abnormal insulin resistance. J Clin Invest 1997;99:1691–1698. Griggs RC, Kingston W, Herr BE, Forbes G, Moxley RT: Lack of relationship of hypogonadism to muscle wasting in myotonic dystrophy. Arch Neurol 1985;42:881–885.

Myotonic Dystrophy

77

63 64 65

66

67

68 69 70 71 72

73

74 75 76 77 78 79 80

81

82

Griggs RC, Halliday D, Kingston W, Moxley RT: Effect of testosterone on muscle protein synthesis in myotonic dystrophy. Ann Neurol 1986;20:590–596. Griggs RC, Pandya S, Florence JM, et al: Randomized controlled trial of testosterone in myotonic dystrophy. Neurology 1989;39:219–222. Chyatte SB, Rudman D, Patterson JH, Ahmann P, Jordan A: Human growth hormone in myopathy: Myotonic dystrophy, Duchenne muscular dystrophy, and limb-girdle muscular dystrophy. South Med J 1974;67:170–172. Thornton CA, Griggs RC, Welle S, Forbes G, Moxley RT: Recombinant human growth hormone (rHGH) treatment increases lean body mass in patients with myotonic dystrophy (abstract). Neurology 1993;43:A280. Vlachopapadopoulou E, Zachwieja JJ, Gertner JM, et al: Metabolic and clinical response to recombinant human insulin-like growth factor I in myotonic dystrophy – A clinical research center study. J Clin Endocrinol Metab 1995;80:3715–3723. Saltiel AR, Olefsky JM: Thiazolidinediones in the treatment of insulin resistance and type II diabetes. Diabetes 1996;45:1661–1669. Kashiwagi K, Nagafuchi S, Sekiguchi N, et al: Troglitazone not only reduced insulin resistance but also improved myotonia in a patient with myotonic dystrophy. Eur J Neurol 1999;41:171–172. Carter JN, Steinbeck KS: Reduced adrenal androgens in patients with myotonic dystrophy. J Clin Endocrinol Metab 1985;60:611–614. Sugino M, Ohsawa N, Ito T, et al: A pilot study of dehydroepiandrosterone sulfate in myotonic dystrophy. Neurology 1998;51:586–589. Sugino M, Ohsawa N, Ito T, Ishida A, Kimura F, Keiichi S: Clinical effects of dehydroepiandrosterone sulfate on myotonic dystrophy: Effects on ADL, myotonia and arrhythmia (abstract). Muscle Nerve 1998(suppl 7):S186. Renaud JF, Desnuelle C, Schmid-Antomarchi H, Hugues M, Serratrice G, Lazdunski M: Expression of apamin receptor in muscles of patients with myotonic muscular dystrophy. Nature 1986;319: 678–680. Behrens MI, Jalil P, Serani A, Vergara F, Alvarez O: Possible role of apamin-sensitive K+ channels in myotonic dystrophy. Muscle Nerve 1994;17:1264–1270. Rudel R, Ruppersberg JP, Spittelmeister W: Abnormalities of the fast sodium current in myotonic dystrophy, recessive generalized myotonia, and adynamia episodica. Muscle Nerve 1989;12:281–287. Kwiecinski H, Ryniewicz B, Ostrzycki A: Treatment of myotonia with antiarrhythmic drugs. Acta Neurol Scand 1992;86:371–375. Mielke U, Haass A, Sen S, Schmidt W: Antimyotonic therapy with tocainide under ECG control in the myotonic dystrophy of Curschmann-Steinert. J Neurol 1985;232:271–274. Sechi GP, Traccis S, Durelli L, Monaco F, Mutani R: Carbamazepine versus diphenylhydantoin in the treatment of myotonia. Eur Neurol 1983;22:113–118. Finlay M: A comparative study of disopyramide and procainamide in the treatment of myotonia in myotonic dystrophy. J Neurol Neurosurg Psychiatry 1982;45:461–463. Monckton DG, Ashizawa T, Siciliano MJ: Murine models for myotonic dystrophy; in Wells RD, Warren ST (eds): Genetic Instabilities and Hereditary Neurological Diseases. San Diego, Academic Press, 1998, pp 181–193. Groenen P, van der Broek W, Coerwinkel M, Wieringa B: Mouse models for myotonic dystrophy: Not simple gain-or-loss of function mutations after all! (abstract). Muscle Nerve 1998(suppl 7): S44. Gourdon G, Lia A, Seznec H, et al: Transgenic mice carrying the human DM region: A model for CTG repeat intergenerational and somatic instability (abstract). Muscle Nerve 1998(suppl 7):S45.

Dr. Richard T. Moxley, Department of Neurology, University of Rochester, School of Medicine and Dentistry, New York, NY 14642 (USA) E-Mail [email protected]

Moxley/Meola

78


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

Muscle Ion Channel Diseases Reinhardt Ru¨del, Karin Jurkat-Rott, Frank Lehmann-Horn Department of Physiology, University of Ulm, Germany

The combination of electrophysiological and molecular genetic investigations has led to the identification of a growing family of ‘ion channel diseases’ that are caused by mutations in various genes encoding voltage-gated or ligandgated ion channels. Since ion channels provide the basis for the regulation of excitability in nerve and muscle cells, it is not surprising that such mutations may lead to dysfunction of these highly specific, membrane-spanning proteins that results in hyper- or hypoexcitability of the respective cells. Channelopathies are known both with skeletal and cardiac muscle. These are the myotonias, periodic paralyses, and the inherited cardiac arrhythmias (the long QT syndromes), respectively. Malignant hyperthermia susceptibility (MHS) and central core disease are two skeletal muscle disorders that are not associated with excitation proper, but with excitation-contraction coupling. This chapter deals with hereditary skeletal muscle diseases caused by mutations in genes encoding voltage-gated ion channels. Defects in the sodium, calcium and chloride channel, but not in the potassium channels of skeletal muscle are known (see table 1).

Sodium Channelopathies Hyperkalemic Periodic Paralysis, Paramyotonia congenita and Potassium-Aggravated Myotonia Clinical and Electrophysiological Characterizations. These three dominantly inherited diseases are all linked to SCN4A, the gene encoding the a subunit of the skeletal muscle sodium channel. Hyperkalemic periodic paralysis (HyperPP) is characterized by episodes of muscle weakness associated with hyperkalemia, often with signs of myotonia in the interval between attacks.

Table 1. List of hereditary muscle diseases caused by pathologic function of voltage-dependent ion channels (voltage dependence of RYR1 is given via DHPR) Channel Disease

Locus

Protein

HyperPP PC PAM

17q23

hSkm-1

4 10 7

Ca+

HypoPP MHS-5 MHS-1 CCD

H q31-32 H 19q12-13.2

DHPR DHPR RYR1 RYR1

3 2 ?20 ?5

Large membrane depolarization Altered muscle metabolism caused by faulty excitation-contraction coupling

ClÖ

DMC RMC

7q35

ClC1 ClC1

?5 ?30

Reduced ClÖ conductance causing after-depolarization

Na+

Mutation number

Pathology

Incomplete channel inactivation causes sustained small or large membrane depolarization

H

MHS-1 and MHS-5>Malignant hyperthermia susceptibility type 1 and type 5; CCD>central core disease; DMC>dominant myotonia congenita; RMC>recessive myotonia congenita.

Paramyotonia congenita (PC) is characterized by a stiffening of the muscles during exercise or when exposed to the cold. In PC the stiffness can give way to weakness, which then may persist for several hours even when the muscles are rapidly rewarmed. Potassium-aggravated myotonia (PAM) is characterized by severe permanent myotonia or fluctuating muscle stiffness that is most prominent about 20 min after exercise [delayed onset myotonia; 1, 2]; the mild form is symptomatically very similar to dominant myotonia congenita Thomsen (which is a chloride channel disease [3]). To date, 21 missense mutations have been discovered leading to the different symptoms described above (fig. 1). The three allelic diseases do not always appear in their pure forms, e.g. PC patients often suffer from spontaneous episodes of weakness which may go along with an elevated serum potassium level. As a clear distinction, HyperPP patients never show substantial stiffness when cold, and muscle weakness never occurs in PAM. Although intermediate forms are frequent, it seems reasonable to retain the classification of three separate nosological entities because, in the pure forms, not only the symptoms but also the recommended treatments differ. The clinical symptoms of the three diseases, muscle stiffness and – in HyperPP and PC – muscle weakness, are not present all the time. Rather, they are elicited by various stimuli. A typical trigger for an episode of weakness in HyperPP is rest after a heavy workload; stiffness and weakness in PC are

Ru¨del/Jurkat-Rott/Lehmann-Horn

80

Fig. 1. Subunits of the voltage-gated sodium channel of skeletal muscle. The a subunit consists of four highly homologous domains (repeats I–IV) containing six transmembrane segments each (S1–S6). The S5–S6 loops form the ion selective pore, and the S4 segments contain positively charged residues conferring voltage dependence to the protein. The repeats are connected by intracellular loops; one of them, the III–IV linker, contains the supposed inactivation particle of the channel. b is an auxiliary subunit. When inserted in the membrane, the four repeats of the protein fold to generate a central pore as schematically indicated at the bottom on the right. Conventional 1-letter abbreviations are used for the replaced amino acids. The different symbols used for the point mutations indicate the resulting diseases as explained at the bottom of the left-hand side.

triggered by muscle exercise and/or exposure of the muscles to the cold; ingestion of potassium-rich food may induce muscle stiffness in PAM patients. In each case, the symptoms taper off spontaneously within a few hours. The episodes reduce the patient’s quality of life considerably, although they may be prevented to a certain extent by appropriate behavior and symptomatic treatment with drugs [for more details, see 1]. Electrophysiological Basis of the Symptoms of the Three Sodium Channelopathies. Stiffness and weakness are caused by the same pathogenetic mechanism, namely a long-lasting depolarization of the muscle fiber membranes. For the explanation of the pathogenesis of the diseases it is important that patients possess two populations of sodium channels, i.e. mutant and wild type.

Muscle Ion Channel Diseases

81

When after an action potential the membrane depolarization caused by the mutant channels is mild (5–10 mV), the wild-type sodium channels that physiologically recover from inactivation can be reactivated by this low ‘resting potential’ [2]. This activation may lead to one or more successive action potentials, which is the basis for the involuntary muscle activity that the patients experience as muscle stiffness. The repetitive firing ends either with an increase of the depolarization, which would leave the wild-type channels in the state of inactivation, or a decrease of the depolarization, which would prevent the wild-type channels from getting reactivated. Such a hyperexcitable state can be computer-simulated and mimicked with anemone toxin [4]. When the depolarization is strong (20–30 mV), the majority of the wildtype channels remain in the state of inactivation, which gradually renders more and more muscle fibers inexcitable. This is the basis of the process of muscle weakness turning into complete flaccid paralysis. This state is also temporary, as the excitability of the muscle fibers returns when, by action of the sodium/potassium pump, the membrane resting potential slowly assumes the physiological value of about Ö80 mV [2]. Knowledge Obtained from Electrophysiological Experiments on Mutant Channels Expressed in Cultured Cells. A detailed specification of the altered channel properties produced by the various disease-causing mutations was possible by heterologous expression of the respective mutant cDNA in cultured cells and subsequent studies of the sodium currents conducted by the membranes of such cells upon depolarizing steps. Whole-cell and patch-clamp recordings showed that all mutations affect the channel inactivation in one way or another (fig. 2). The alterations that were observed with such mutant channels were a reduced speed of current decay following a depolarization step, a more or less incomplete decay of the current, an increased speed of the recovery from inactivation, a shifted position of the steady-state inactivation curve (Hodgkin and Huxley’s h= curve) and an altered degree of uncoupling of inactivation from activation [2]. The alterations found with the different disease-causing mutants are summarized in table 2. They can be generalized as follows. (1) Sodium currents conducted by HyperPP mutants show a large fraction of persistent current and an incomplete slow inactivation. These changes may cause strong and long-lasting depolarization, which is the basis of weakness in HyperPP [4]. (2) Sodium currents conducted by PC mutants are characterized by a slowing of fast inactivation. This change explains paradoxical myotonia. Another typical change is acceleration of recovery from inactivation and a left shift of the steady-state inactivation curve. In combination with an increased persistent current it could explain the cold-induced weakness. (3) Sodium currents conducted by PAM mutants are characterized by an increased persistent fraction and/or slowing of


82

Fig. 2. Hinged-lid model of fast inactivation of sodium channels and the effects of mutations at various locations on the current decay. a Bird’s eye view on the channel consisting of four similar repeats (I–IV). The channel is cut and spread open between repeats II and III to allow the view on the intracellular loop between repeats III and IV. The loop acts as the inactivation gate whose ‘hinge’ GG (>a pair of glycines) allows it to ‘swing’ between two positions, i.e. the noninactivated channel state (pore open, left panel) and the inactivated state (pore blocked by the ‘plug’ IFM representing amino acid sequence isoleucine, phenylalanine, methionine, right panel). b The substitution of E (Glu) for Gly-1306 slows channel inactivation (left two panels, compare fast current decay in wild-type channel on far left) and leads to a life-threatening form of PAM. The designed substitution of QQQ (Gln-GlnGln) for IFM (Ile-Phe-Met) completely abolishes channel inactivation (two right hand panels) proving that the loop between repeats III and IV is indeed the inactivation gate [adapted from 15, 16].

fast inactivation. These alterations explain the slight depolarization which causes the myotonia. An also existing right shift of the steady-state inactivation curve might be the reason why PAM patients do not experience weakness. Discussion of the Various Disease-Causing Mutations. Most of the amino acid substitutions caused by disease-causing mutations are in the ‘inactivating’ linker between repeats III and IV or adjacent in the ‘voltage-sensing’ segment S4 of repeat IV. Almost all remaining mutations are situated at the inner side of the membrane where they could impair the docking site for the inactivation particle (fig. 2).


83

Table 2. Summary of the electrophysiological properties of PAM, PC and HyperPP sodium channel mutations Disease

Slowing of fast Persistent Steady-state fast Recovery from Impaired slow inactivation current inactivation fast inactivation inactivation

PC PAM HyperPP

++ + Ö

+ +/++ +++

k K Ö

++ Ö/+ Ö

Ö Ö +

+ to +++ indicate the severity of the alteration; Ömeans no change. Arrows show direction of shift of the steady-state fast inactivation curve (to the left>to more negative potentials).

Three of the mutations in the III/IV linker (Gly-1306-Ala,Val,Glu) produce different amino acid substitutions for one of a pair of glycines proposed to act as the ‘hinge’ for the inactivation gate. They all cause PAM and, interestingly, the more the structure of the substituting amino acid differs from the physiological Gly-1306 (longer side chains and/or greater charge) the more intensive is the hyperexcitability of the muscles and the more severe the myotonia. Alanine, with a short side chain, results in a benign, often ‘subclinical’ form of myotonia. Valine, having a side chain of intermediate size, causes moderate exercise-induced myotonia. Glutamic acid, an amino acid with a long side chain, causes permanent myotonia, the most severe form of the disease. Thus the natural mutations affecting Gly-1306 provided evidence for an increased rigidity of the amino acid chain at the position of the highly conserved pair of glycines increasingly hampering channel inactivation [5]. Mutations at this hinge site also altered channel activation and deactivation. A similar correlation between the structural differences between wild-type and mutant amino acids on one hand, and the severity of clinical symptoms on the other was found for four PC-causing mutants at another identical site (Arg-1448-Cys,His,Pro,Ser) near the extracellular face of IVS4. This finding led to a systematic application of site-directed mutagenesis in this supposed channel activation domain. All tested mutants primarily affected channel inactivation. Therefore, it was hypothesized that depolarization-induced movements in IVS4 concern both the inactivation gate and the docking site for the inactivation particle [6]. Pathogenesis of Symptoms. Although the symptoms in PC are very much aggravated in the cold, the sodium currents conducted by PC-causing mutants expressed in heterologous cells did not show a corresponding dependence on temperature. The mechanism by which the cold enhances muscle stiffness in


84

PC patients is not completely clear. Both the time constant of fast inactivation and the persistent current increase with cooling, and mutant and normal channels show the same temperature dependence; however, the absolute figures are larger for mutant channels at any temperature. Therefore, it was proposed that a certain threshold has to be exceeded in the cold environment to induce myotonic and/or paralytic symptoms. In contrast to the cold-induced symptoms, the pathogenesis of the potassium-induced stiffness and paralysis is well understood. The physiological depolarization, which follows an elevation of serum potassium according to Nernst increases the open probability of the sodium channels and unmasks their inactivation defect. Thus, potassium exerts its effect via depolarization. Open Questions. It is not entirely clear why patients with PC are temperature-sensitive whereas those with PAM and HyperPP are not, since a specific temperature dependence could not be found with any of the PC-causing mutants in vitro. On the other hand, the cold-induced weakness is clearly linked to membrane depolarization due to the increased sodium inward current, so that a mechanism other than via the sodium channel seems unlikely. As to the aggravation of the clinical symptoms upon potassium intake with PAM and HyperPP patients, studies on PAM- or PC-causing mutants also showed sensitivity to extracellular potassium. Therefore, the effect of potassium is most likely explained by a membrane depolarization. Further not yet explained problems with sodium channelopathies are the occurrence of a myopathy with the HyperPP-causing mutation Thr-704-Met and the pathology of normokalemic periodic paralysis.

Calcium Channelopathies Two Important Calcium Channels Are Expressed in Skeletal Muscle These are the dihydropyridine receptor, DHPR, an L-type voltage-dependent calcium channel, and the ryanodine receptor, RYR1, coupled to it (fig. 3, 4). They are situated in the triadic junctions of the transverse tubular system and the sarcoplasmic reticulum (SR), respectively. In skeletal muscle, the DHPR appears to be physiologically unimportant as ion-conducting channel; it rather functions as a voltage sensor for the ryanodine receptor (RYR1) which releases calcium from the SR, thus initiating contraction [7]. The a1S subunit of the a1S-a2/d-b1-c pentameric DHPR complex interacts with the RYR1 via the II–III interlinker. Disease-causing mutations are known in the genes for either channel. Certain point mutations in CACNAS, the gene encoding a1S, cause familial hypokalemic periodic paralysis (HypoPP), a disease characterized by episodes of muscle weakness. Other


85

point mutations in this gene, as well as mutations in the RYR1 gene, cause malignant hyperthermia, a potentially lethal condition triggered by certain anesthetics. Hypokalemic Periodic Paralysis Characterization of the Disease. The major symptom of this dominantly inherited disease, episodes of generalized paralysis, may occur less frequently and be on average of longer duration than in HyperPP. Decisive for the classification is the level of serum potassium, [K]e, during a paralytic attack, which may fall below 2 mM in HypoPP, whereas in HyperPP it may rise beyond 4.5 mM. The hypokalemia is assumed to be caused by stimulation of the Na/K pump by insulin which is the physiological mechanism by which potassium ions are transported from the extracellular to the intracellular space. Very low [K]e causes instability of the membrane potential because then the potassium conductance approaches zero. Even in normal muscle, a [K]e 01.0 mM causes membrane depolarization, not hyperpolarization. An increase in [K]e is then the easiest therapy for normalization and stabilization of the resting potential [for a review, see 1]. The Genetic Defect in HypoPP. The disease is linked to chromosome 1q3132 and cosegregates with the gene encoding the DHPR a1S subunit which is located in this region. Sequencing of cDNA derived from muscle biopsies of patients revealed three mutations [6, 8]. Two of them are analogous predicting arginine to histidine substitutions within the highly conserved S4 regions of repeats II and IV (Arg-528-His and Arg-1239-His, respectively), the rare third mutation predicts an arginine to glycine substitution in IVS4 (Arg-1239-Gly) (fig. 3). Pathogenesis. Electrophysiological investigation of biopsied muscle specimens from HypoPP patients in vitro showed that the paralysis induced by hypokalemia is due to a sustained depolarization of the sarcolemma to about Ö50 mV. The crucial role of the depolarization for the pathogenesis of paralysis was demonstrated by experiments in which openers of ATP-sensitive potassium channels were given in vitro. Their hyperpolarizing action was able to prevent muscle weakness or even restore normal force. The pathogenesis of the membrane depolarization is still unclear. L-type calcium currents conducted by Arg-528-His and Arg-1239-His mutant channels were studied in various native and heterologous expression systems. In summary, they revealed neither a significant alteration of gating nor a change of the resulting calcium transients that could explain the membrane depolarization following a decrease in serum potassium. It seems likely that additional structures are involved that interact somehow with the DHPR and/or may respond to hypokalemia. Recently, a reduced potassium current conducted


86

Fig. 3. Subunits of the voltage-gated calcium channel of skeletal muscle. The a1 subunit resembles that of the sodium channel, however the function of the various parts, e.g. the III–IV linker, may not be the same. a2/d, b1–b4, and c are auxiliary subunits. Mutations in the a1 subunit have been described for HypoPP and MHS. Conventional 1-letter abbreviations are used for the replaced amino acids. The symbols used for the point mutations indicate the resulting diseases as explained at the bottom of the left-hand side.

by ATP-dependent channels was reported in biopsied muscle specimens from three HypoPP patients carrying the Arg-528-His mutation [9]. The authors proposed a contribution of this finding to the pathogenesis of HypoPP. Malignant Hyperthermia Susceptibility Not a Disease, But a Genetically Determined Disposition. Susceptibility to malignant hyperthermia (MH), a potentially lethal event in carriers of voltageor ligand-gated calcium channel mutations, may cause clinically inconspicuous individuals to respond abnormally when exposed to volatile anesthetics or depolarizing muscle relaxants. MH is generated by a pathologically high increase of the myoplasmic calcium concentration during exposure to the triggering agents. This leads to increased muscle metabolism and heat production resulting in symptoms of muscle rigidity, hyperthermia associated with metabolic acidosis, hyperkalemia and hypoxia [10, 11] The metabolic alterations usually progress rapidly. Early administration of dantrolene, an inhibitor of


87

Fig. 4. Cartoon of the homotetrameric ryanodine receptor (RYR1), the calcium release channel situated in the membrane of the SR. The cytosolic part of the protein complex, the so-called foot, bridges the gap between the transverse tubular system and the SR. The mutations shown cause susceptibility to MH and central core disease. Conventional 1-letter abbreviations are used for the replaced amino acids whose positions are given by the ascribed amino acid numbers of RYR1 [modified after 7].

calcium release from the SR has successfully aborted numerous fulminant crises and has reduced the mortality rate to less than 10%. MHS is genetically heterogeneous. In many families, mutations in the gene encoding RYR1 (fig. 4) have been found. Also two mutations in the DHPR a1 subunit have been described (see below), which underlines the functional link between the two protein complexes. Another possible locus contains the gene encoding the a2/d subunit of the DHPR [6].


88

MHS Due to RYR1 Mutations. More than 20 disease-causing point mutations in RYR1 have been identified, all situated in the long N terminus of the protein, the so-called foot of the channel complex (see fig. 4) which contains the binding sites for various activating ligands like calcium (lM), ATP, calmodulin (which binds in the absence of calcium), caffeine and ryanodine (nM), and inactivating ligands like calcium (110 lM) and magnesium in mM concentrations. The diagnostically important increased sensitivity of MH-susceptible muscle to caffeine is considered to be caused by an altered RYR1 function. Functional tests, so far only performed with mutant porcine muscle showing an MHS equivalent in isolated SR vesicles, have shown that calcium regulation is disturbed. Lower calcium concentrations activate the channel to a higher than normal level, and higher than normal calcium concentrations are required to inhibit the channel. Investigations of reconstituted RYR1 in lipid bilayers, designed to find the reason for the increased caffeine sensitivity, led to controversial results. Single-channel measurements on RYR1 did not show increased caffeine sensitivity whereas pharmacological studies showed increased sensitivity [11]. Functional characterization of the various mutations in the N-terminus and the central part of the ‘foot’ gave similar results, i.e. increased sensitivity of the mutant RYR1 to activating concentrations of calcium and exogenous and diagnostically used ligands such as caffeine, halothane, and 4-chloro-mcresol. Overexpression of mutant ryanodine receptors in normal human primary muscle cells also led to an increased calcium response during exposure to a triggering agent [11]. A reduced inhibition of calcium release by magnesium has been reported for MHS muscle and proposed as the major pathomechanism of MHS. MHS Caused by L-Type Channel a1 Subunit Mutations. Cosegregation of MHS with markers on chromosome 1q32 was shown for two families. Screening for causative mutations in the candidate gene, CACNA1S, revealed arginine1086 to histidine and cysteine substitutions in the intracellular interlinker connecting domains III and IV of the protein (fig. 3), a region that hitherto was not known to be important for excitation-contraction coupling [6, 11]. Central Core Disease This congenital proximal myopathy with structural alterations mainly of type 1 fibers is allelic to MH [10]. It is transmitted as an autosomal dominant trait. Name giving are central areas along the whole fiber length that contain structured or unstructured myofibrils and lack mitochondria. Affected individuals show hypotonia at birth (floppy infant syndrome). Later in life muscle strength usually improves except for rare cases showing progressive muscle weakness. Exercise-induced muscle cramps are often reported. Anesthesia-


89

induced events of MH have been reported, and central core disease patients usually give positive results in the diagnostic in vitro contracture test. These observations induced genetic linkage studies on chromosome 19q12-13.2, the first MH locus. Linkage was indeed found and mutations in RYR1 were detected (see fig. 4). Even though events similar to MH may occur in numerous muscle disorders during general anesthesia, a genetic relation to MH exists with certainty only in central core disease and possibly in the King-Denborough syndrome, a disease characterized by dwarfism, scoliosis, ptosis and further skeletal or muscular symptoms [11]. Data on the molecular genetics of the latter disease is still missing. Lethal events during anesthesia have been reported for both of these rare diseases.

Chloride Channelopathies Myotonia may not only be due to sodium channel mutations as in PAM, but also to changes in the chloride channel CLC-1 as in myotonia congenita (Thomsen). The clinical symptoms of these two diseases are almost indistinguishable although their pathogenetic mechanisms are rather different [12]. Myotonia congenita Characterization of the Two Types. This disease is transmitted with either a dominant or recessive mode of inheritance; both types are caused by mutations in CLCN1, the gene encoding the major skeletal muscle chloride channel [13]. Muscle stiffness is temporary and can affect every skeletal muscle of the body. Myotonic stiffness is most pronounced when a forceful movement is abruptly initiated after the muscles were rested for 5 min or more. For instance, after making a hard fist, the patient may not be able to extend the fingers fully for several seconds. Myotonia decreases or completely subsides when the same movement is repeated several times (warm-up phenomenon), but it always recurs after a few minutes of rest. On rare occasions, a sudden, frightening noise may cause instantaneous generalized stiffness. The patient may then fall to the ground and remain rigid and helpless for some seconds or even minutes. Even more disabling may be a transient weakness. Typically myotonic muscles generate a characteristic pattern in the acoustic electromyogram, i.e. bursts of action potentials with amplitude and frequency modulation, so-called dive bombers [3]. The dominant form is very rare, as less than 10 different families were identified at the molecular level. The recessive form is much more frequent, between 1:23,000 and 1:50,000. Males seem to be affected more often than


90

Fig. 5. Membrane topology model of the skeletal muscle chloride channel monomer, CLC-1, originally based on hydropathy analysis. The functional channel is a homodimer. The different symbols used for the known mutations leading to dominant Thomsen-type myotonia and recessive Becker-type myotonia are explained at the bottom on the left. Conventional 1-letter abbreviations are used for replaced amino acids [modified after 13].

females with a ratio of 3:1 when only the typical clinical features are taken into account. However, family studies disclosed that women are affected at the same frequency though to a much lesser degree. Molecular Pathology. The muscle stiffness is caused by the fact that, following voluntary excitation, the membranes of individual muscle fibers may continue for some seconds to generate runs of action potentials. This activity prevents immediate muscle relaxation from occurring. The overexcitability is caused by a permanent reduction of the resting chloride conductance of the muscle fiber membranes. The high chloride conductance is necessary for a fast repolarization of the transverse tubular membranes, when these tend to stay depolarized by potassium accumulated in the tubules during tetanic muscle excitation [3]. Both dominant and recessive myotonia congenita are linked to chromosome 7q35 and CLCN1, the gene encoding the chloride channel. It spans at least 40 kb and contains 23 exons whose boundaries have been located [13]. Knowledge Obtained from Electrophysiological Experiments on Mutant Channels Expressed in Cultured Cells. Functional expression of CLCN1 has


91

6


92

been accomplished in Xenopus oocytes, human embryonic kidney (HEK-293) cells and the insect cell line Sf-9 [13]. The resulting currents were similar to those found in native muscle fibers. Electrophysiological studies of wild-type and mutant channel proteins have provided first insight into the pharmacology and structure-function relationships of CLC-1, and led to the identification of regions involved in gating and permeation [13]. Inferences from experiments with the chloride channel CLC-0 and studies of CLC-1 constructs strongly suggest that functional channels are formed as homodimers [6]. More than 30 point mutations and three deletions have been found in the channel gene, and they cause either dominant or recessive myotonia congenita by producing change or loss of function of the gene product (fig. 5). Experiments with myotonia-generating drugs showed that blockade of 50% of the physiological chloride current is not sufficient to produce myotonic activity. This could be the reason why heterozygous carriers of recessive mutations that completely destroy the channel function are without clinical myotonia. Dominant inheritance is explained by a mutant channel monomer that can form dimers and, in doing so, produces a dominant negative effect. The most common feature of the thereby resulting chloride currents is a shift of the activation curve towards more positive membrane potentials reducing the total chloride conductance (fig. 6). Surprisingly, the degree of the shift and the clinical severity sometimes disagree, e.g. Gln-552-Arg causes an unusually large potential shift, however a very mild clinical phenotype, myotonia levior [6]. Open Questions. No modern experiments have been reported designed to test the explanation for the warm-up phenomenon that stems unchallenged from pre-molecular biology days. Several mutations were found that lead to myotonia congenita which under certain circumstances is dominantly and under other circumstances recessively transmitted. What the decisive circumstance is has so far not been elucidated. Perhaps it is connected to one or the other polymorphisms in CLCN1 that have no functional consequences within the wild type.

Fig. 6. Recordings from human skeletal muscle CLC-1 channels expressed in a mammalian cell line. Currents from normal (WT) and dominant myotonia-causing mutant (Gly200-Arg) channels are compared. a, b Macroscopic currents, recorded in the whole-cell mode, were activated from a holding potential of 0 mV by voltage steps to potentials of Ö145 to +95 mV, and deactivated after 400 ms by polarization to Ö105 mV. c Voltage dependence of the relative open probability that is much reduced for the mutant channel in the physiological potential range. All mutations that cause such a voltage shift have dominant effects [modified after 14].


93

All dominant mutations shift the activation curve of CLC-1 towards more positive membrane potentials. There is, however, no simple relation between the amount of shift and the severity of symptoms seen in patients carrying the various mutations. Apparently other, not yet detected factors act in an ancillary fashion. Such factors may also play a role in cases of recessive myotonia where the symptoms fluctuate [14]. Even more enigmatic is the finding that some ‘recessive’ mutations do not lead to the loss of a gene product but to channels that in the usual expression systems conduct chloride currents with normal amplitude and gating behavior. Relatively little is known about the structure/function relation of CLC-1. Several groups of investigators are involved in finding the pore region of the channel and the mechanism of gating. Molecular biological methods, such as site-directed mutagenesis or use of channel chimeras, will hopefully help to overcome this unsatisfactory situation.

Acknowledgments The support by the Deutsche Forschungsgemeinschaft (Ru 138-20), the Interdisziplina¨res Zentrum fu¨r Klinische Forschung (IZKF) of the University of Ulm, the Muscular Dystrophy Association (MDA), and the European Community, TMR Programme on Excitation-Contraction Coupling, is gratefully acknowledged.

References 1

2 3 4 5

6 7 8 9

Lehmann-Horn F, Engel AG, Ricker K, Ru¨del R: The periodic paralyses and paramyotonia congenita; in Engel AG, Franzini-Armstrong C (eds): Myology, ed 2. New York, McGraw-Hill, 1994, vol 2, pp 1303–1334. Lehmann-Horn F, Ru¨del R: Molecular pathophysiology of voltage-gated ion channels. Rev Physiol Biochem Pharmacol 1996;128:195–268. Ru¨del R, Lehmann-Horn F, Ricker K: The nondystrophic myotonias; in Engel AG, FranziniArmstrong C (eds): Myology, ed 2. New York, McGraw-Hill, 1994, vol 2, pp 1291–1302. Cannon SC: From mutation to myotonia in sodium channel disorders. Neuromuscul Disord 1997; 7:241–249. Mitrovic N, George AL Jr, Lerche H, Wagner S, Fahlke C, Lehmann-Horn F: Different effects on gating of three myotonia-causing mutations in the inactivation gate of the human muscle sodium channel. J Physiol (Lond) 1995;487:107–114. Lehmann-Horn F, Jurkat-Rott K: Voltage-gated ion channels and hereditary disease. Physiol Rev 1999;79:1317–1371. Melzer W, Herrmann-Frank A, Lu¨ttgau HC: The role of Ca2+ ions in excitation-contraction coupling of skeletal muscle fibres. Biochim Biophys Acta 1995;1241:59–116. Ptaćˇek LJ: The familial periodic paralyses and nondystrophic myotonias. Am J Med 1998;105: 58–70. Tricarico D, Servidei S, Tonali P, Jurkat-Rott K, Camerino DC: Impairment of skeletal muscle adenosine triphosphate-sensitive K+ channels in patients with hypokalemic periodic paralysis. J Clin Invest 1999;103:675–682.


94

10 11 12 13 14

15

16

Loke J, MacLennan DH: Malignant hyperthermia and central core disease: Disorders of Ca2+ release channels. Am J Med 1998;104:470–486. Jurkat-Rott K, McCarthy TV, Lehmann-Horn F: Genetics and pathogenesis of malignant hyperthermia. Muscle Nerve 2000;23:4–17. Hudson AJ, Ebers GC, Bulman DE: The skeletal muscle sodium and chloride channel diseases. Brain 1995;118:547–563. Pusch M, Jentsch TJ: Molecular physiology of voltage-gated chloride channels. Physiol Rev 1994; 74:813–827. ¨ zdemir Wagner S, Deymeer F, Ku¨rz LL, Benz S, Schleithoff L, Lehmann-Horn F, Serdaroglu P, O C, Ru¨del R: The dominant chloride channel mutant G200R causing fluctuating myotonia: Clinical findings, electrophysiology, and channel pathology. Muscle Nerve 1998;21:1122–1128. West JW, Patton DE, Scheuer T, Wang Y, Goldin AL, Catterall WA: A cluster of hydrophobic amino acid residues required for fast Na+-channel inactivation. Proc Natl Acad Sci USA 1992;89: 10910–10914. Mitrovic N, Lerche H, Heine R, Fleischhauer R, Pika-Hartlaub U, Hartlaub U, George AL Jr, Lehmann-Horn F: Role in fast inactivation of conserved amino acids in the IV/S4-S5 loop of the human muscle Na+ channel. Neurosci Lett 1996;214:9–12.

Dr. Reinhardt Ru¨del, Department of Physiology, University of Ulm, D–89069 Ulm (Germany) E-Mail [email protected]


95


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

Congenital Myasthenic Syndromes Andrew G. Engel a, Kinji Ohno a, Anthony A. Stans b a b

Department of Neurology and Orthopedic Surgery, Mayo Clinic, Rochester, Minn., USA

Congenital myasthenic syndromes (CMS) are inherited diseases in which the safety margin of neuromuscular transmission is compromised by one or more distinct mechanisms [1, 2]. Although congenital myasthenia was recognized as early as 1937 [3], the CMS were not investigated until after the discovery of the autoimmune origin of myasthenia gravis (MG) in the 1970s. In the 1970s and 1980s, ultrastructural, cytochemical, and in vitro microelectrode studies of CMS patients revealed a heterogeneous group of disorders: endplate (EP) acetylcholinesterase (AChE) deficiency [4], the slow-channel syndrome [5], a defect in the synthesis or vesicular packaging of acetylcholine (ACh) [6], decreased quantal release due to paucity of synaptic vesicles [7], and EP acetylcholine receptor (AChR) deficiency [8,9]. In 1990s, following the discovery of the cDNA sequences of all human AChR subunits and of the catalytic and tail subunits of AChE, molecular genetic studies took center stage. In addition, patch clamping of human intercostal muscle EPs allowed resolution and analysis of single channel currents through EP AChRs [10], and the advent of mammalian expression systems facilitated detailed analysis of the properties of engineered mutant AChRs and AChEs.

Classification of CMS During the past 12 years, my coworkers and I investigated 120 CMS kinships at the Mayo Clinic. On the basis of our studies, we classify the CMS as presynaptic (7%), synaptic (12.5%), postsynaptic (78%), and of unknown origin (2.5%) (see table 1).

Table 1. Classification of CMS based on 120 index patients Defect site Presynaptic defects Paucity of synaptic vesicles Defect in ACh synthesis/packaging Congenital Lambert-Eaton-like syndrome Synaptic defect EP AChE deficiency Postsynaptic defects Primary kinetic abnormality with or without AChR deficiency Primary AChR deficiency with or without minor kinetic abnormality No identified defecta Total a

Index cases

1 6 1 15 30 64 3 120

Includes 2 cases of limb-girdle myasthenia.

Diagnosis of a CMS A typical clinical history for CMS is one of ocular, bulbar or respiratory muscle symptoms worsened by crying or activity in the neonatal period, fluctuating ocular palsies and abnormal fatigability on exertion during infancy and childhood, normal or delayed motor milestones, sometimes progression of symptoms during adolescence or adult life, and negative tests for antiAChR antibodies. Some syndromes (e.g. the slow-channel syndrome and some CMS caused by mutations in the AChR e-subunit) may not present until the 2nd or 3rd decade of life, and in the CMS with episodic apnea the patients have only mild symptoms or are asymptomatic between attacks of respiratory and bulbar paralysis precipitated by fever, infection, excitement or overexertion [6]. The intravenous edrophonium test is positive except in EP AChE deficiency and in the CMS with episodic apnea between attacks, and is inconsistently positive in the slow-channel syndrome. On examination, the most important clue to a defect of neuromuscular transmission is increasing weakness on sustained exertion, as shown by increasing ptosis during upward gaze, an arm elevation time less than 30 s, or increasing difficulty in performing even a few deep-knee bends. Patients with severe involvement of the truncal muscles rapidly develop postural scoliosis and shift their weight from one foot to another on standing. Selectively severe weakness of cervical and of wrist and finger extensor muscles is found in older patients with EP AChE deficiency [11] and in the slow-channel syndrome [5]. The pupillary light reflexes are delayed in patients with EP AChE deficiency [11].

Congenital Myasthenic Syndromes

97

Ocular muscle involvement can be absent or mild in some cases of EP AChE deficiency [11], the slow-channel syndrome [5], and in the CMS with episodic apnea [12]. The tendon reflexes are preserved but can be hypoactive. Despite the above diagnostic clues, in most instances the clinical examination cannot reliably distinguish between different types of CMS. The generic diagnosis of a CMS must be supported by a decremental electromyographic (EMG) response at low-frequency (2 Hz) stimulation in at least one muscle, or by abnormal jitter and blocking during single-fiber EMG. The decremental response may be absent during the interictal periods in the CMS with episodic apnea. Here a decremental response can be elicited by 10 Hz stimulation for 5–10 min, or by exercise for several minutes before 2 Hz stimulation [6]. In patients taking high doses of AChE inhibitors, in patients with EP AChE deficiency [4, 11], and in the slow-channel CMS (SCCMS) [5], single-nerve stimuli evoke a primary compound muscle action potential (CMAP) followed by one or more repetitive CMAPs, each separated by an interval of 5–10 ms. The repetitive potentials are smaller and decrement faster than the primary response at all frequencies of stimulation. Therefore, the test must be done in patients not exposed to AChE inhibitors, after a period of rest, and initially with single nerve stimuli. Observations in the EMG laboratory can provide an objective estimate of responsiveness to AChE inhibitors or other cholinergic agents. For example, one can compare the decrement observed in a given muscle before and a few minutes after an intravenous dose of edrophonium, or 60–90 min after an oral dose of 3,4-diaminopyridine (3,4-DAP). The differential diagnosis of CMS in the neonatal period, infancy and childhood includes spinal muscular atrophy, morphologically distinct congenital myopathies, congenital muscular dystrophies, infantile myotonic dystrophy, mitochondrial myopathy, brainstem anomaly, Mo¨bius’ syndrome, congenital fibrosis of the extraocular muscles, infantile botulism and seropositive- and seronegative-autoimmune MG. In older patients, the differential diagnosis includes motor neuron disease, limb girdle or facioscapulohumeral dystrophy, mitochondrial myopathy, chronic fatigue syndrome, and seropositive- and seronegative-autoimmune MG. Radial nerve palsy, peripheral neuropathy and syringomyelia have been incorrectly diagnosed in some cases of the SC CMS. Most entities can be excluded by careful physical examination, serologic tests and EMG studies.

Investigation of the CMS A deeper understanding of disease mechanisms and a precise classification of the CMS requires estimation of the number of AChRs per EP, light-

Engel/Ohno/Stans

98

and electron-microscopic analysis of EP morphology, and electrophysiologic assessment of EP function in vitro. Conventional microelectrode studies of EP potentials and currents readily reveal whether the transmission defect is presynaptic or postsynaptic. Patch-clamp recordings of currents flowing through single AChR channels provide precise information on the conductance and kinetic properties of the channels. All the above studies can be performed only on small bundles of muscle intact from origin to insertion. These are dissected from a larger strip of intercostal or anconeus muscle, also intact from origin to insertion, removed from the patient with minimal trauma. If the foregoing studies point to a defect in candidate gene or protein, then molecular genetic analysis becomes feasible. If a mutation is discovered in the candidate gene, then expression studies with the genetically engineered mutant molecule can be used to confirm pathogenicity and to analyze the kinetic or structural consequences of the mutation. To date, the candidate gene approach has resulted in the discovery of 18 mutations in the gene encoding the collagenic tail subunit of AChE, and close to 70 mutations in the genes coding for AChR subunits.

Presynaptic Syndromes The presynaptic origin of these syndromes has been established by electrophysiologic and morphologic methods, but the underlying defects at the protein and gene level are still not known. Paucity of Synaptic Vesicles and Reduced Quantal Release The clinical features of this rare CMS closely mimic those of autoimmune MG and the symptoms respond to anticholinesterase drugs. The paucity of synaptic vesicles in the nerve terminal reduces the number of releasable transmitter quanta, which reduces the safety margin of neuromuscular transmission [7]. The decreased synaptic vesicle density could arise from (1) a defect in the formation of synaptic vesicle precursors in the anterior horn cell, (2) a defect in the axonal transport of one or more species of precursor vesicles to the nerve terminal, (3) impaired assembly of mature synaptic vesicles from their precursors in the nerve terminal, or (4) impaired recycling of the synaptic vesicles in the nerve terminal. CMS with Episodic Apnea This rare syndrome usually presents at birth or in the neonatal period with hypotonia, variable ptosis but normal ocular ductions, severe bulbar weakness causing dysphagia, and respiratory insufficiency with cyanosis and


99

apnea. If the infant survives, the symptoms improve but the crises recur with infections, fever, excitement, vomiting or overexertion. During crises, episodes of apnea can cause sudden death or anoxic brain injury. Between crises, patients may have only mild or no myasthenic symptoms. When weakness is absent, it can be readily induced by exercise. With increasing age, the exacerbations become less frequent [6, 12–19]. The clinical features of the disease were described by Greer and Schotland [13] in 1960. In 1975, Conomy et al. [12] referred to the disease under the rubric of ‘familial infantile myasthenia’, a term that subsequently became entrenched in the medical literature. Because all CMS can be familial and because most CMS present in infancy, the term ‘familial infantile myasthenia’ has become a source of confusion [20]. We therefore refer to the disease as CMS with episodic apnea. Electron microscopy studies show no postsynaptic abnormality [6]. Stimulation of small muscle bundles at 10 Hz in vitro results in an abnormal decrease of the amplitude of the CMAP, EP potential, and miniature EP potential, pointing to impaired resynthesis or vesicular packaging of ACh [6, 21]. Treatment consists of anticholinesterase drugs given orally between crises, and parenterally, together with respiratory support, during crises. CMS Resembling the Lambert-Eaton Myasthenic Syndrome In one patient reported with this syndrome, the amplitude of the CMAP was reduced but facilitated severalfold on tetanic stimulation and the symptoms were improved by guanidine [22]. In a second patient, observed at the Mayo Clinic, quantal release by nerve impulse was very low at 1 Hz stimulation but increased markedly with stimulation rates ?10 Hz. The pre- and postsynaptic regions were structurally intact by electron microscopy and the nerve terminals harbored abundant synaptic vesicles. The patient responded only partially to combined treatment with pyridostigmine and 3,4-DAP. The defect likely resides in the presynaptic voltage-gated calcium channel or in a component of the synaptic vesicle release complex.

Synaptic AChE Deficiency The synaptic type of CMS is caused by absence or marked decrease of the asymmetric species of AChE in the synaptic basal lamina [4]. Owing to this, the synaptic currents are prolonged and evoke repetitive CMAPs. Prolonged exposure of AChR to ACh causes desensitization of AChR [23], a depolarization block at physiologic rates of stimulation [24], and an EP myopathy stemming from cationic overloading of the postsynaptic region [25]. The presynaptic terminals are abnormally small and often encased by Schwann cells, reducing

Engel/Ohno/Stans

100

a

b

Fig. 1. Schematic diagram showing domains of a ColQ strand with 18 identified ColQ mutations (a) and components of the A12 species of asymmetric AChE (b). The N-terminal region of each ColQ strand contains a PRAD that binds a homotetramer of the catalytic AChET subunit. The triple helical collagenic domain contains two positively charged heparan sulfate proteoglycan-binding domains (HSPBD) that participate in anchoring the tail subunit in the synaptic basal lamina. The C-terminal region of ColQ is essential for assembly of the triple helix of the collagen domain in a C- to N-terminal direction and may also participate in anchoring the tail subunit. Reprinted with permission [2].

the number of quanta released by nerve impulse [4, 11]. The safety margin of neuromuscular transmission is compromised by reduced quantal release, loss of AChR due to degeneration of junctional folds, and by desensitization and a depolarization block during activity. There are two major types of AChE in skeletal muscle [26, 27]: (1) globular forms consisting of monomers (G1), dimers (G2), or tetramers (G4) of the T isoform of the catalytic subunit (ACHET), and (2) asymmetric forms consisting of ACHET subunits linked to a tail subunit formed by the association of three collagen-like strands, ColQ (fig. 1). The conserved domains of the tail subunit include a proline-rich attachment domain (PRAD) in the N-terminal region of ColQ that binds an ACHET tetramer producing A4, A8 and A12 moieties [28, 29], a central collagen domain where three ColQ strands composed of GXY triplets form a triple helix, and a C-terminal region where the ColQ strands are enriched in charged residues and cysteines. The collagen domain


101

also harbors two positively charged heparan sulfate proteoglycan-binding domains implicated in binding the triple helical collagen domain to negatively charged molecules in the basal lamina [30, 31]. The function of the tail subunit is to insert asymmetric AChE into synaptic basal lamina, and asymmetric AChE is the predominant species of AChE at the EP [32, 33]. In 1998, human COLQ cDNA was cloned [34, 35] and the genomic structure of COLQ determined [34]. This, in turn, led to the discovery of 6 truncation [34] and 1 missense [35] mutations of COLQ in 7 AChE-deficient patients. One truncation mutation was upstream of PRAD, 4 were in the collagen domain, and 1 truncation and the missense mutation were in the C-terminal region. Coexpression of each COLQ mutant with wild-type ACHET in COS cells revealed that a mutation proximal to PRAD prevented the association of ColQ with ACHET; mutations distal to PRAD generated a mutant species of AChE composed of one ACHET tetramer and a truncated single and insertion-incompetent ColQ strand. Subsequent studies in our laboratory led to the discovery of 11 more ColQ mutations in 8 additional kinships [36–39]. Expression studies indicate that 4 of the new mutations prevent the formation of asymmetric AChE, but 7 others, all in the C terminal region, produce some asymmetric AChE. However, the mutant asymmetric AChEs are likely insertion incompetent because patients harboring these mutations have EP AChE deficiency.

Postsynaptic Syndromes Increased Response to ACh: The SCCMS The phenotypic consequences of the syndrome stem from prolonged opening episodes of the AChR channel (see fig. 2b) and spontaneous channel openings even in the absence of ACh. These cause (1) cationic overloading of the junctional sarcoplasm and an EP myopathy with loss of AChR from degenerating junctional folds, and (2) a depolarization block due to staircase summation of the markedly prolonged EP potentials during physiologic activity [40–42]. Eleven SCCMS mutations have been reported to date [40–48]. The different mutations occur in different AChR subunits and in different functional domains of the subunits (see table 2, fig. 2a). Each is dominant, causing a pathologic gain of function. Patch clamp studies at the EP, mutation analysis, and expression studies in human embryonic kidney fibroblast (HEK) cells identify three types of mutations. Those residing in the second transmembrane domain (M2], which lines the channel pore, decrease the free energy required for channel opening, as indicated by an increased channel opening rate even

Engel/Ohno/Stans

102

a

b

Fig. 2. a Schematic diagram of slow-channel (solid circles) and fast-channel (shaded circles) mutations reported to date. The drawing on the left shows a section through the AChR lodged in the lipid bilayer with two slow-channel mutations, aG153S and aV156M, in the extracellular domain near the ACh-binding site of the a subunit, and 3 fast channel mutations: eP121L near the ACh-binding site of the e subunit, aV156M in the M3 domain of the a subunit, and e1254ins18 in the long-cytoplasmic loop of the e subunit. The drawing on the right shows slow-channel mutations detected between the M2 and M3 domains of the a subunit, in the M2 domains of the a, b and e subunits, and in the M1 domain of the a subunit. b Examples of single-channel currents from wild-type, slow-channel and fastchannel AChRs expressed in HEK cells. Reprinted with permission [2].


103

Table 2. Distribution of 70 mutations in AChR subunit genes Mutation

Subunit genes a

Point mutations Slow-channel mutation Low affinity fast-channel mutation Fast-channel mutation with abnormal gating Null mutation Reduced expression

8

d

e

2

3 2 1 3a 10

13 2 2 3 18

1

1 1

2 1

1

17a 7a 4a

18 7 4

49

70

2

1 6

Inframe rearrangement Reduced expression Fast-channel mutation with mode-switching kinetics Premature chain termination (null mutations) Frameshifting rearrangement Splice-site mutation Nonsense mutation Total

b

15

Total

4

2

Includes 33 published and 32 unpublished CMS mutations identified in our laboratory and 5 mutations reported from other laboratories. Numbers in italics indicate recessive mutations that reduce AChR expression. a Null mutations.

in the absence of ACh, and increase the free energy required for channel closure, as indicated by a delay in channel closure [40, 42, 45]. Mutations near the ACh-binding site on the a subunit increases affinity for ACh, causing repeated channel reopenings during the prolonged ACh occupancy [41, 47]. A third type of SCCMS has features of the two preceding types and the mutations reside in the M1 or M2 domain [42, 44, 45]. Following the lead that quinidine is a long-lived open-channel blocker of AChR [49], Fukudome et al. [50] showed that clinically attainable levels of quinidine normalized the prolonged opening episodes of mutant slow-channels expressed in HEK cells. On the basis of these findings, Harper and Engel [51] treated SCCMS patients with quinidine-producing serum levels of 0.7–2.5 lg/ml (2.1–7.7 lM/l) and found that the patients were improved by clinical as well as by EMG criteria.

Engel/Ohno/Stans

104

Decreased Response to ACh: The Fast-Channel Syndromes A reduced synaptic response to ACh occurs with recessive, loss-of-function mutations of AChR that reduce the affinity for ACh, or primarily affect channel gating, or cause mode-switching kinetics (Table 2, fig. 2). Each type of mutation results in brief activation episodes (see fig. 2b) and reduces the probability of channel opening. In all three disorders, the mutated allele causing the kinetic abnormality is accompanied by a null mutation in the second allele so that the kinetic mutation dominates the clinical phenotype. Fast-channel syndrome patients respond well to combined therapy with 3,4-DAP (1 mg/kg/day given in 3–5 divided doses), which increases the number of quanta released by nerve impulse, and cholinesterase inhibitors, which increase the number of AChRs activated by each quantum. Low-Affinity Fast-Channel Syndrome. This disorder was observed in 2 patients. Both had very small MEPCs but normally abundant EP AChR and normal EP ultrastructure. Patch-clamp studies revealed infrequent and briefer than normal channel opening events, and resistance to desensitization by ACh. Each patient had two heteroallelic AChR e subunit gene mutations: a common eP121L mutation that affects ACh binding, and a signal peptide mutation (eG-8R; patient 1), and a glycosylation consensus site mutation (eS143L; patient 2). AChR expression in HEK cells was normal with eP121L but was markedly reduced with the other mutations. Therefore eP121L defines the clinical phenotype. Studies of engineered eP121L AChR revealed a markedly decreased rate of channel opening, and a reduced affinity for ACh in the open channel and desensitized states [52]. Fast-Channel Syndrome Due to a Gating Abnormality. This syndrome is caused by a kinetic and relatively low-expressor mutation in the M3 domain of the AChR a subunit, aV285I, together with a null mutation, aF233V, that unmasks the kinetic consequences of aV285I. In this CMS, as in the lowaffinity fast-channel syndrome, the duration of channel open intervals and bursts is markedly shortened, but the primary abnormality resides in the channel gating mechanism and not in affinity for ACh. Studies of genetically engineered aV285I-AChR in HEK cells revealed brief channel opening events (see fig. 2b), due to a slow opening rate constant, b, and fast closing rate constant, a, and a reduced probability of channel opening [53]. Fast-Channel Syndrome Due to Mode-Switching Kinetics. In this disorder, the kinetic abnormality is caused by an inframe duplication in the long cytoplasmic loop of e, e1254ins18, which also reduces AChR expression, plus a cysteine-loop null mutation, eC128S [54]. When expressed in HEK cells, e1254ins18 causes mode switching in the kinetics of receptor activation in which the normal high efficiency of gating is accompanied by two new modes that gate inefficiently, opening more slowly and closing more rapidly than


105

Fig. 3. Schematic diagram of 28 lowexpressor and null mutations in the e sub>Low-expressor promoter mutaunit. >low-expressor missense mutations; tions; T>truncating null mutations.

normal. At the EP, e1254ins18-AChR shows very brief activation episodes during steady-state application of ACh and appears electrically silent during the synaptic response to ACh. The phenotypic consequences are EP AChR deficiency and compensatory expression of fetal AChR harboring the c instead of the e subunit (c-AChR), which restores electrical activity at EP and rescues the phenotype [54]. Mutations Causing AChR Deficiency with or without Minor Kinetic Abnormalities CMS with severe EP AChR deficiency result from different types of homozygous or, more frequently, heterozygous recessive mutations in AChR subunit genes. The mutations are concentrated in the e subunit (see tables 2, 3, fig. 3). There are two possible reasons for this. (1) Expression of the fetal type c subunit, although at a low level, may compensate for absence of the e subunit [54–56], whereas patients harboring null mutations in subunits other than e might not survive for lack of a substituting subunit. (2) The gene encoding the e subunit, and especially the exons coding for the long cytoplasmic loop, have a high GC content that likely predisposes to DNA rearrangements [57]. Morphologic studies show an increased number of EP regions distributed over an increased span of the muscle fiber. The integrity of the junctional folds is preserved but AChR expression on the folds is patchy and faint. Some EP regions are simplified and small. The amplitude of miniature EP potentials and currents is reduced but quantal release by nerve impulse is often higher than normal. With null or low-expressor mutations in the e subunit, single channel recordings at the EP [56, 58] or immunocytochemical studies [55] reveal the presence of c-AChR at the EP that likely rescues the phenotype.

Engel/Ohno/Stans

106

Table 3. Twenty-nine low-expressor or null mutations in the AChR e subunit Mutations

Domains

Ref. No.

e-156CKT e-155GKA eG-8R eV-13D eT51P T eR64X T e59ins5 T e70insG T e1275 eC128S eS143L eR147L T e553del7 T e723delC * eP245L T eIVS7+2TKC T e760ins8 * eR311W T eIVS9-1GKC T e1012del20 T e1033delG T e1101insT T eIVS10+2TKG T e1206ins19 * e1254ins18 T e1260del23 T e1267delG T e1276delG T e1293insG

Ets binding site Ets binding site Signal peptide Signal peptide Extracellular domain Extracellular domain Extracellular domain Extracellular domain Extracellular domain Disulfide loop N-glycosylation site Extracellular domain Extracellular domain Ml domain M1 domain Link between M1 and M2 M2 domain Long cytoplasmic loop Long cytoplasmic loop Long cytoplasmic loop Long cytoplasmic loop Long cytoplasmic loop Long cytoplasmic loop Long cytoplasmic loop Long cytoplasmic loop Long cytoplasmic loop Long cytoplasmic loop Long cytoplasmic loop Long cytoplasmic loop

65 64 52 61 61 56 59 59 56 54 52 56 56, 58, 63 61 56 59 61 56 63 60 67 55 61 59 54 67 61, 62 59 55

>Promoter mutations, reduced expression; >missense mutations, reduced expression; T>premature chain termination, null mutations; *>mutation also has significant kinetic effects.

Most patients respond to anticholinesterase drugs and some derive additional benefit from 3,4-DAP. Different types of recessive mutations causing severe EP AChR deficiency have been identified (see table 2). (1) Mutations causing premature termination of the translational chain, these mutations are frameshifting [55, 56, 59–62],


107

occur at a splice site [59, 61, 63], or produce a stop codon directly [56]. (2) Point mutations in the promoter region of a subunit gene (e-155GKA [64] and e-156CKT [65]). (3) Missense mutation in a signal peptide region (eG-8R) [52]. (4) Mutations involving residues essential for assembly of the pentameric receptor. Mutations of this type were observed in the e subunit at an Nglycosylation site (eS143L) [52], in cysteine 128 (eC128S), a residue that is an essential part of the C128-C142 disulfide loop in the extracellular domain [54], and in arginine 147 (eR147L) in the extracellular domain, which lies between isoleucine 145 and threonine 150, residues that contribute to subunit assembly [56], and with a 3-codon deletion in the long cytoplasmic loop of the b subunit [66]. (5) Missense mutations affecting both AChR expression and kinetics. For example, eR311W in the long cytoplasmic loop between M3 and M4 decreases [56], whereas eP245L in the M1 domain increases [56] the open duration of channel events. In the case of eR311W and eP245L, the kinetic consequences are modest and are likely overshadowed by the reduced expression of the mutant gene.

Conclusion CMS are heterogeneous disorders caused by presynaptic, synaptic or postsynaptic defects. Full characterization of a given syndrome requires correlation of clinical and EMG data, light- and electron-microscopic examination of EP morphology, estimation of the number and distribution of AChRs at the EP, in vitro eletrophysiologic analysis of parameters of neuromuscular transmission, and molecular genetic studies when the preceding investigations point to a candidate gene or protein. Three presynaptic CMS have been recognized to date. All are recessively inherited but their molecular genetic basis remains undeciphered. One syndrome, in which a paucity of synaptic vesicles causes reduced release of transmitter quanta, closely mimics the clinical and EMG features of autoimmune MG. In another syndrome, a defect in the resynthesis or vesicular packaging of ACh causes episodic crises with apnea. In the third disorder, the electrophysiologic features resemble those of the Lambert-Eaton syndrome and the putative defect resides in the synaptic vesicle release complex. A synaptic type of CMS is caused by absence or marked deficiency of the asymmetric species of AChE in the synaptic basal lamina. The disease is now known to be caused by recessive mutations in the collagenic tail subunit of asymmetric AChE. The mutations prevent the association of the tail subunit with catalytic subunits, or the insertion of the tail subunit into the synaptic basal lamina.

Engel/Ohno/Stans

108

All postsynaptic CMS recognized thus far stem from mutations in AChR subunit genes that increase or decrease the synaptic response to ACh. An increased response occurs in the slow-channel syndromes. Here dominant mutations in different AChR subunits and in different domains of the subunits prolong the activation episodes of AChR by altering channel gating, or by increasing the affinity for ACh, or both. A decreased synaptic response to ACh occurs in the fast-channel syndromes where the mutated allele causing the kinetic abnormality is accompanied by a null mutation in the second allele, so that the mutation causing the kinetic abnormality dominates the clinical phenotype. The kinetic abnormalities reduce the affinity for ACh, or primarily affect channel gating, or cause mode-switching kinetics. In each instance, the AChR activation episodes are abnormally brief and occur at a reduced probability. Response to ACh is also reduced by low-expressor or null mutations in AChR subunit genes that result in premature termination of the translational chain or are missense mutations preventing subunit assembly or glycosylation. These mutations are concentrated in the e subunit, probably because substitution of the fetal c for the adult e subunit can rescue humans from fatal null mutations in e.

References 1 2 3 4

5

6 7

8

9

10

Engel AG: Myasthenic Syndromes; in Engel AG, Franzini-Armstrong C (eds): Myology: Basic and Clinical, ed 2. New York, McGraw-Hill, 1994, pp 1798–1835. Engel AG, Ohno K, Sine SM: Congenital myasthenic syndromes; in Engel AG (ed): Myasthenia gravis and Myasthenic Disorders. New York, Oxford University Press, 1999, pp 251–297. Rothbart HB: Myasthenia gravis. Familial occurrence. JAMA 1937;108:715–717. Engel AG, Lambert EH, Gomez MR: A new myasthenic syndrome with end-plate acetylcholinesterase deficiency, small nerve terminals, and reduced acetylcholine release. Ann Neurol 1977;1:315– 330. Engel AG, Lambert EH, Mulder DM, Torres CF, Sahashi K, Bertorini TE, Whitaker JN: A newly recognized congenital myasthenic syndrome attributed to a prolonged open time of the acetylcholineinduced ion channel. Ann Neurol 1982;11:553–569. Mora M, Lambert EH, Engel AG: Synaptic vesicle abnormality in familial infantile myasthenia. Neurology 1987;37:206–214. Walls TJ, Engel AG, Nagel AS, Harper CM, Trastek VF: Congenital myasthenic syndrome associated with paucity of synaptic vesicles and reduced quantal release. Ann NY Acad Sci 1993;681:461– 468. Engel AG, Hutchinson DO, Nakano S, Murphy L, Griggs RC, Gu Y, Hall ZW, Lindstrom J: Myasthenic syndromes attributed to mutations affecting the epsilon subunit of the acetylcholine receptor. Ann NY Acad Sci 1993;681:496–508. Vincent A, Newsom-Davis J, Wray D, Shillito P, Harrison J, Betty M, Beeson D, Mills K, Palace J, Molenaar P, Murray N: Clinical and experimental observations in patients with congenital myasthenic syndromes. Ann NY Acad Sci 1993;681:451–460. Milone M, Hutchinson DO, Engel AG: Patch-clamp analysis of the properties of acetylcholine receptor channels at the normal human endplate. Muscle Nerve 1994;17:1364–1369.


109

11

12 13 14 15 16 17 18 19 20 21

22 23 24

25

26 27

28 29

30 31 32 33 34

Hutchinson DO, Walls TJ, Nakano S, Camp S, Taylor P, Harper CM, Groover RV, Peterson HA, Jamieson DG, Engel AG: Congenital endplate acetylcholinesterase deficiency. Brain 1993;116: 633–653. Conomy JP, Levisohn M, Fanaroff A: Familial infantile myasthenia gravis: A cause of sudden death in young children. J Pediatr 1975;87:428–429. Greer M, Schotland M: Myasthenia gravis in the newborn. Pediatrics 1960;26:101–108. Fenichel GM: Clinical syndromes of myasthenia in infancy and childhood. A review. Arch Neurol 1978;35:97–103. Robertson WC, Chun RWM, Kornguth SE: Familial infantile myasthenia. Arch Neurol 1980;37: 117–119. Smit LM, Barth PG: Arthrogryposis multiplex congenita due to congenital myasthenia. Dev Med Child Neurol 1980;22:371–374. Baptist EC, Landes RV, Sturman JK: Familial infantile myasthenia gravis: A preventable cause of death. South Med J 1985;78:201–202. Gieron MA, Korthals JK: Familial infantile myasthenia gravis. Report of three cases with followup into adult life. Arch Neurol 1985;42:143–144. Zammarchi E, Donati MA, Masi S, Sarti A, Castelli S: Familial infantile myasthenia: A neuromuscular cause of respiratory failure. Childs Nerv Syst 1994;10:347–349. ¨ zdemir C: Familial infantile myasthenia: Confusion in terminology. Deymeer F, Serdaroglu P, O Neuromuscul Disord 1999;9:129–130. Hart Z, Sahashi K, Lambert EH, Engel AG, Lindstrom J: A congenital, familial, myasthenic syndrome caused by a presynaptic defect of transmitter resynthesis of mobilization (abstract). Neurology 1979;29:559. Bady B, Chauplannaz G, Carrier H: Congenital Lambert-Eaton myasthenic syndrome. J Neurol Neurosurg Psychiatry 1987;50:476–478. Katz B, Thesleff S: A study of the ‘desensitization’ produced by acetylcholine at the motor endplate. J Physiol (Lond) 1957;138:63–80. Maselli RA, Soliven BC: Analysis of the organophosphate-induced electromyographic response to repetitive nerve stimulation: Paradoxical response to edrophonium and d-tubocurarine. Muscle Nerve 1991;14:1182–1188. Salpeter MM, Kasprzak H, Feng H, Fertuck H: End-plates after esterase inactivation in vivo: Correlation between esterase concentration, functional response and fine structure. J Neurocytol 1979;8:95–115. Massoulie´ J, Pezzementi L, Bon S, Krejci E, Valette F-M: Molecular and cellular biology of cholinesterases. Prog Neurobiol 1993;41:31–91. Rotundo RL, Fambrough DM: Function and molecular structure of acetylcholinesterase; in Engel AG, Franzini-Armstrong C (eds): Myology, ed 2. New York, McGraw-Hill, 1994, pp 607– 623. Bon S, Coussen F, Massoulie´ J: Quaternary associations of acetylcholinesterase. II. The polyproline attachment domain of the collage tail. J Biol Chem 1997;272:3016–3021. Simon S, Krejci E, Massoulie´ J: A four-to-one association between peptide motifs: Four C-terminal domains from cholinesterase assemble with one proline-rich attachment domain (PRAD) in the secretory pathway. EMBO J 1998;17:6178–6187. Deprez PN, Inestrosa NC: Two heparin-binding domains are present on the collagenic tail of asymmetric acetylcholinesterase. J Biol Chem 1995;270:11043–11046. Peng HB, Xie H, Rossi SG, Rotundo RL: Acetylcholinesterase clustering at the neuromuscular junction involves perlecan and dystroglycan. J Cell Biol 1999;145:911–921. Hall ZW: Multiple forms of acetylcholinesterase and their distribution in endplate and non-endplate regions of rat diaphragm muscle. J Neurobiol 1973;4:343–361. Younkin SG, Rosenstein C, Collins PL, Rosenberry TL: Cellular localization of the molecular forms of acetylcholinesterase in rat diaphragm. J Biol Chem 1982;257:13630–13637. Ohno K, Brengman JM, Tsujino A, Engel AG: Human endplate acetylcholinesterase deficiency caused by mutations in the collagen-like tail subunit (ColQ) of the asymmetric enzyme. Proc Natl Acad Sci USA 1998;95:9654–9659.

Engel/Ohno/Stans

110

35

36

37

38

39

40

41

42

43 44

45

46

47

48

49 50 51 52

Donger C, Krejci E, Serradell P, Eymard B, Bon S, Nicole S, Chateau D, Gary F, Fardeau M, Massoulie´ J, Guicheney P: Mutation in the human acetylcholinesterase-associated gene, COLQ, is responsible for congenital myasthenic syndrome with end-plate acetylcholinesterase deficiency. Am J Hum Genet 1998;63:967–975. Ohno K, Brengman JM, Milone M, Shen X-M, Tsujino A, Anlar B, Engel AG: Congenital endplate acetylcholinesterase deficiency: Novel missense and null mutations in the collagen-like tail subunit of the asymmetric enzyme (abstract). Am J Hum Genet 1998;63:A377. Ohno K, Brengman JM, Nakano S, Walsh P, Heidenreich FR, Vincent A, Nagwane S, Engel AG: Congenital endplate acetylcholinesterase deficiency caused by nonsense, splice-site, and missense mutations in the collagenic tail subunit of asymmetric AChE (abstract). Neurology 1999;52(suppl 2): A184. Ohno K, Engel AG, Brengman JM, Harper CM, Shen X-M, Heidenreich FR, Vincent A, Milone M, Tan E, Demirci M, Walsh P, Nakano S, Akiguchi I: The spectrum of mutations causing endplate acetylcholinesterase deficiency. Ann Neurol 2000;47:162–170. Ohno K, Brengman JM, Felice KJ, Cornblath DR, Engel AG: Congenital endplate acetylcholinesterase deficiency caused by a nonsense mutation and an A-to-G splice donor site mutation at position +3 of the collagen-like tail subunit gene (COLQ): How does G at position +3 result in aberrant splicing? Am J Hum Genet 1999;65:635–644. Ohno K, Hutchinson DO, Milone M, Brengman JM, Bouzat C, Sine SM, Engel AG: Congenital myasthenic syndrome caused by prolonged acetylcholine receptor channel openings due to a mutation in the M2 domain of the e subunit. Proc Natl Acad Sci USA 1995;92:758–762. Sine SM, Ohno K, Bouzat C, Auerbach A, Milone M, Pruitt JN, Engel AG: Mutation of the acetylcholine receptor a subunit causes a slow-channel myasthenic syndrome by enhancing agonist binding affinity. Neuron 1995;15:229–239. Engel AG, Ohno K, Milone M, Wang H-L, Nakano S, Bouzat C, Pruitt JN, Hutchinson DO, Brengman JM, Bren N, Sieb JP, Sine SM: New mutations in acetylcholine receptor subunit genes reveal heterogeneity in the slow-channel congenital myasthenic syndrome. Hum Mol Genet 1996; 5:1217–1227. Ohno K, Hutchinson DO, Milone M, Nakano S, Sieb JP, Brengman JM, Engel AG: Molecular genetic basis of a slow channel syndrome (abstract). Muscle Nerve 1995;18:463. Wang H-L, Auerbach A, Bren N, Ohno K, Engel AG, Sine SM: Mutation in the M1 domain of the acetylcholine receptor alpha subunit decreases the rate of agonist dissociation. J Gen Physiol 1997;109:757–766. Milone M, Wang H-L, Ohno K, Fukudome T, Pruitt JN, Bren N, Sine SM, Engel AG: Slow-channel syndrome caused by enhanced activation, desensitization, and agonist binding affinity due to mutation in the M2 domain of the acetylcholine receptor alpha subunit. J Neurosci 1997;17:5651–5665. Gomez CM, Maselli R, Gammack J, Lasalde J, Tamamizu S, Cornblath DR, Lehar M, McNamee M, Kuncl R: A beta-subunit mutation in the acetylcholine receptor gate causes severe slow-channel syndrome. Ann Neurol 1996;39:712–723. Croxen R, Newland C, Beeson D, Oosterhuis H, Chauplanaz G, Vincent A, Newsom-Davis J: Mutations in different functional domains of the human muscle acetylcholine receptor a subunit in patients with the slow-channel congenital myasthenic syndrome. Hum Mol Genet 1997;6:767–773. Ohno K, Milone M, Brengman JM, Lo Monaco M, Evoli A, Tonali P, Engel AG: Slow-channel congenital myasthenic syndrome caused by a novel mutation in the acetylcholine receptor e subunit (abstract). Neurology 1998;50:A432. Sieb JP, Milone M, Engel AG: Effects of the quinoline derivatives quinine, quinidine, and chloroquine on neuromuscular transmission. Brain Res 1996;712:179–189. Fukudome T, Ohno K, Brengman JM, Engel AG: Quinidine normalizes the open duration of slowchannel mutants of the acetylcholine receptor. Neuroreport 1998;9:1907–1911. Harper CM, Engel AG: Quinidine sulfate therapy for the slow-channel congenital myasthenic syndrome. Ann Neurol 1998;43:480–484. Ohno K, Wang H-L, Milone M, Bren N, Brengman JM, Nakano S, Quiram P, Pruitt JN, Sine SM, Engel AG: Congenital myasthenic syndrome caused by decreased agonist binding affinity due to a mutation in the acetylcholine receptor e subunit. Neuron 1996;17:157–170.


111

53

54

55 56

57 58

59 60

61

62

63

64

65

66

67

Wang H-L, Milone M, Ohno K, Shen X-M, Tsujino A, Batocchi AP, Tonali P, Brengman JM, Engel AG, Sine SM: Acetylcholine receptor M3 domain: Stereochemical and volume contributions to channel gating. Nat Neurosci 1998;2:226–233. Milone M, Wang H-L, Ohno K, Prince RJ, Shen X-M, Brengman JM, Griggs RC, Engel AG: Mode switching kinetics produced by a naturally occurring mutation in the cytoplasmic loop of the human acetylcholine receptor e subunit. Neuron 1998;20:575–588. Engel AG, Ohno K, Bouzat C, Sine SM, Griggs RG: End-plate acetylcholine receptor deficiency due to nonsense mutations in the e subunit. Ann Neurol 1996;40:810–817. Ohno K, Quiram P, Milone M, Wang H-L, Harper CM, Pruitt JN, Brengman JM, Pao L, Fischbeck KH, Crawford TO, Sine SM, Engel AG: Congenital myasthenic syndromes due to heteroallelic nonsense/missense mutations in the acetylcholine receptor e subunit gene: Identification and functional characterization of six new mutations. Hum Mol Genet 1997;6:753–766. Krawczak M, Cooper DN: Gene deletions causing human genetic disease: Mechanisms of mutagenesis and the role of the local DNA sequence environment. Hum Genet 1991;86:425–441. Milone M, Ohno K, Pruitt JN, Brengman JM, Sine SM, Engel AG: Congenital myasthenic syndrome due to frameshifting acetylcholine receptor epsilon subunit mutation. Soc Neurosci Abstr 1996;22: 1942. ¨ zdirim E, Brengman JM, De Bleecker J, Engel AG: Myasthenic syndromes in Ohno K, Anlar B,O Turkish kinships due to mutations in the acetylcholine receptor. Ann Neurol 1998;44:234–241. Ohno K, Fukudome T, Nakano S, Milone M, Feasby TE, Tyce GM, Engel AG: Mutational analysis in a congenital myasthenic syndrome reveals a novel acetylcholine receptor epsilon subunit mutation. Soc Neurosci Abstr 1996;22:234. Middleton L, Ohno K, Christodoulou K, Brengman JM, Milone M, Neocleous V, Serdaroglu P, ¨ zdemir C, Mubaidin A, Horany K, Al-Shebab A, Mavromatis I, Mylonas I, Tsingis Deymeer F, O M, Zamba E, Pantzaris M, Kyriallis K, Engel AG: Congenital myasthenic syndromes linked to chromosome 17p are caused by defects in acetylcholine receptor e subunit gene. Neurology 1999; 53:1076–1082. Croxen R, Beeson D, Vincent A, Newsom-Davis J: Congenital myasthenic syndrome with a single nucleotide deletion at the intron/exon boundary in exon 12 of the gene encoding the acetylcholine receptor e subunit (abstract). Ann Neurol 1996;40:513. Ohno K, Engel AG, Milone M, Brengman JM, Sieb JP, Iannaccone S: A congenital myasthenic syndrome with severe acetylcholine receptor deficiency caused by heteroallelic frameshifting mutations in the epsilon subunit (abstract). Neurology 1995;45(suppl 4):A283. Ohno K, Anlar B, Engel AG: Congenital myasthenic syndrome caused by a mutation in the Etsbinding site of the promoter region of the acetylcholine receptor e subunit gene. Neuromuscul Disord 1999;9:131–135. Nichols P, Croxen R, Vincent A, Rutter R, Hutchinson M, Newsom-Davis J, Beeson D: Mutation of the acetylcholine receptor e-subunit promoter in congenital myasthenic syndrome. Ann Neurol 1999;45:439–443. Quiram P, Ohno K, Milone M, Patterson MC, Pruitt JN, Brengman JM, Sine SM, Engel AG: Acetylcholine receptor b-subunit mutations causing endplate AChR deficiency and reduced assembly with the d subunit (abstract). Neurology 1999;52(suppl 2):A185. Brengman JM, Ohno K, Shen X-M, Engel AG: Congenital myasthenic syndrome due to two novel mutations in the acetylcholine receptor e subunit gene (abstract). Muscle Nerve 1998;21(suppl 7): S120.

Andrew G. Engel, MD, Department of Neurology, Mayo Clinic, 200 First Street SW, Rochester, MN 55905 (USA) Tel. +1 507 284 5102, Fax +1 507 284 5831, E-Mail [email protected]

Engel/Ohno/Stans

112


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

Juvenile and Late-Onset Myasthenia gravis ¨ zdemir Feza Deymeer, Piraye Serdarog˘lu, Cos¸kun O Department of Neurology, University of Istanbul, Istanbul, Turkey

Myasthenia gravis (MG) is an autoimmune disease which can start at any age, predominantly affecting young women and older men [1]. In early hospitalbased studies [2, 3], onset of the disease was found to be rare in the prepubertal period; it made a peak during fertile years with a decline thereafter in women, while the peak was in late life in men. Epidemiological studies in the last 2 decades have resulted in a change of concepts about the frequency of MG in the elderly population [4], with the implication that MG may be ‘disproportionately a disease of later years’ [5], while basically confirming previously established facts for other age groups. Recently, the two extremes of age, the young and the old, have been among topics of interest in MG, the former because it may have some clinical peculiarities [6, 7] and the latter because of the ageing population [8] with a high incidence of MG [9, 10]. In this chapter, we review the literature on juvenile (JMG) and late-onset (LOMG) MG, concentrating on studies which specifically deal with young and elderly patients, and add our experience with 733 patients attending the Neuromuscular Clinic of the University of Istanbul (which will be referred to as UI).

Juvenile MG JMG is defined as an acquired autoimmune disorder, not encompassing neonatal MG and congenital myasthenic syndromes [6]. Onset is after the age of 1 in JMG, while it is usually before the age of 1 in congenital myasthenic syndromes and within the first few hours of life in neonatal MG [1, 11].

Table 1. JMG studies TXb %

Remission rate

Ocular: 3% Respiratory difficulty: 43%

60 (0)

29% (TX) 14% (no TX)

2.6

Ocular: 9% Generalized: 91%

47 (1)

38 (TX) 24 (no TX)

=15

2.4

I: 19% IIa: 17% IIb: 43 IIIc: 6 IV: 15%

60 (1)

43 % (TX)

32

q1 to p18

1.9

I: 40% IIa: 21% IIb: 11% IIcd: 28%

22 (0)

29% (TX) 12% (no TX)

Rodriguez et al. [16]

149

?1 to p16

=12 years: 2.7 q12 years: 3.7

I: 9% IIa: 16% IIb: 73% III: 2%

57

53% (TX) 30% (no TX)

Wong et al. [7]

101 (39 of 262)

?0 to =16

1.1

Ocular: 71% Generalized: 29%

Low or absent 12 (8)

17% (TX) 34% (no TX)

Andrews et al. [6]

115

?1 to =20

Pre: 1 Peri and post: 4.5

Pre: 50 Peri: 68 Post: 91

72 (1)

35% (early TX) 2% (late TX) 9% (no TX)

012 years: 1 q12 years: F1M

Crises: 33%

34

37

20% (no TX)

Authors

Number of patientsa

Age years

FMR

Classification

Millichap and Dodge [12]

35 (8 of 447)

?1 to p16

=12 years: 3.3 q12 years: 10

Seybold et al. [13]

102

?1 to p16

Ryniewicz and Badurska [14]

47

Snead et al. [15]

Pre: 20 Peri and post: 93 Unknown: 2

Positive antiAChR Ab, %

51

Anlar et al. [17]

30

p15

Lindner et al. [18]

79

q12 to 4 p18

I: 9% IIa: 45% IIb: 32% III and IV: 14%

81

82 (0)

60% (TX) 29% (no TX)

Evoli et al. [19]

133 (16 of 817)

11 to 0 20

Pre I: 26% IIa: 21% IIb: 11% III and IV: 42% Peri and post I: 16% IIa: 38% IIb: 31% III and IV: 15%

Pre: 74

Pre: 47 (0)

Pre: 0% (TX) 21% (no TX)

Peri and post: 86

Peri and Peri and post post: 84 31% (TX) (4) 0% (no TX)

Pre: 19 Peri and post: 114

Pre 1.1

Peri and post: 4.2

¨ zdemir Deymeer/Serdarog˘lu/O

114

Table 1 (continued) Authors

Number of patientsa

Age years

FMR

Classification

Positive antiAChR Ab, %

TXb %

Remission rate

UI study

140 (19 of 733)

q1 to 0 20

Pre: 0.9

Pre I: 50% IIa: 31% IIIb: 13% III and IV: 6% Peri and post I: 13% IIa: 27% IIb: 45% III and IV: 15%

Pre: 64

Pre: 0 (0)

Pre 25% (no TX)

Peri and post: 78

Peri and Peri and post: post: 55 8% (TX) (2) 28% (no TX)

Pre: 19 Peri and post: 121

Peri and post: 3

Pre, peri, and post refer to the stage of puberty. FMR>Female to male ratio, TX>thymectomy. a % of total number is given in parentheses. b Number of thymomas is given in parentheses. c Generalized weakness without ocular or bulbar signs. d Generalized disease with bulbar involvement.

In selecting JMG studies [6, 7, 12–19], compiled in table 1, we preferred those which gave some information on the whole group of juvenile patients even though the main purpose might have been to evaluate thymectomy and we refrained from including studies done with patients undergoing thymectomy only [20]. Most of the JMG studies appropriately excluded neonatal MG and congenital myasthenic syndromes. The main source of difficulty [21, 22] in the comparison of different studies was the arbitrary choice of the upper age limit for the juvenile period, ranging from 15 to 19. The juvenile period is heterogenous with pubertal stages which have different characteristics. When the juvenile period is considered as a unity, the uniqueness of each stage is lost. Unfortunately, studies on JMG used different cutoff ages to separate prepuberty from postpuberty and most of them confined themselves to giving the female to male ratio (FMR) for the different stages. Seybold et al. [13] were among the few who treated prepuberty separately. Andrews et al. [6, 23] analyzed the different stages in more detail, applying Tanner’s criteria [24] to their patient population and divided it into three stages: prepuberty (08.9 years in females and09.3 years in males), postpuberty (115.3 years in females and 116.5 years in males) and peripuberty in between these ages. Like Evoli et al. [19], we used these age brackets in our study for the three stages. Demographic Features We analyzed demographic data in three categories: (1) studies dealing specifically with JMG patients (table 1), (2) large hospital-based studies [2, 3,

Juvenile and Late-Onset Myasthenia gravis

115

25–29], (3) population-based (epidemiological) studies with incidence rates [9, 10, 30–34]. This separation was at times artificial and some studies, analyzed within one category, actually belonged to more than one category. JMG Studies (table 1). In Caucasian series, JMG made up less than 20% of the total patient population. Prepubertal MG constituted 2–4% of the entire group of patients [19] (UI). Females were affected more frequently than males, with this predominance being much less during prepuberty as compared to later juvenile years. In some studies, the two sexes were almost equally affected during prepuberty. In black prepubertal children, the incidence of MG was higher than in comparable white children and FMR was 2 both in pre- and postpubertal stages [6, 35]. The Chinese exhibited unique features in relation to puberty [7], the highest incidence for both sexes being in the 1st decade, with a peak at 2–3 years of age. Hospital-Based Studies. There are several large hospital-based studies (listed above) which give an idea of the age- and sex-related characteristics of MG even though the main focus is not any specific age group. In these studies, prepubertal patients made up the smallest portion of the whole group, only paralleled by the oldest age brackets. A review of the literature [36] revealed that in 4.3% of all cases the onset was before the age of 10 and in 24% before the age of 20. Again, female predominance was evident, less so in prepuberty than in postpuberty. There were even some studies in which males predominated during prepuberty [2, 29]. Population-Based (Epidemiological ) Studies. More precise demographic information was obtained in population-based studies (listed above), which are not flawed by the selection bias inherent in hospital-based studies. Ageand sex-specific incidence rates of some of these studies are compared in table 2. Overall, it can be said that the incidence rate was one of the lowest of all age groups in the 1st decade, where female preponderance was not very striking. The rate increased for both sexes in the 2nd decade, but much more so for females than for males. In the Japanese [37], the highest incidence for both sexes was in the 1st decade of life, similar to the Chinese [7]. Females and males were equally affected. The apparent discrepancy in the FMR between Caucasian and ChineseJapanese populations in the juvenile period is probably just a function of the over-representation of prepubertal patients in the Chinese and the Japanese studies and tends to disappear when prepuberty is considered separately in Caucasian studies. Onset Symptoms It is well-known that ocular presentation is very high in prepuberty in the Chinese and the Japanese, being 71 and 90%, respectively, in the two


116

Table 2. Age- and sex-specific incidence rates per million population per year Age, years Storm-Mathisen [30]

010 10–19 20–29 30–39 40–49 50–59 60–69 70–79 180 a

Giagheddu et al. [31]

Ferrari and Lovaste Christensen et al. [34] [9]

females

males

females

males

females

0.53 4.58 8.34 7.21 4.61 4.34 4.69 6.44 1.35

0.10 0.66 2.09 2.33 2.99 2.63 7.02 3.48 2.99

0.5 4.7 4.4 5.7 5.2 3.6 5.9 2.2 a

1.0 0.9 1.1 4.0 3.3 2.9 0.9 a

3.1 14.8 13.1 10.2 3.6 20.1 14.1 a

males 4.1

9.2 6.9 7.7 10.1

females

males

1.5 4.0 7.0 6.1 5.1 7.3 9.7 11.7 1.3

0.7 1.2 2.2 2.1 2.1 5.2 15.1 14.1 8.6

Over 70 years of age.

studies [7, 37]. JMG also commonly presents with ocular symptoms in Caucasians [12, 15, 17, 38]. Information provided in the study by Bundey [36] for onset symptoms lends itself to the separation of pubertal stages [6]. Seybold et al. [13] also give detailed information on onset symptoms for 35 prepubertal patients. Two interesting features emerge for Caucasians from these two studies together with the UI study: the striking predominance of ocular presentation in prepuberty and the presence of a peculiar onset symptom, lower extremity weakness, becoming most evident in peri- and postpuberty. Prepuberty. Presentation was with purely ocular symptoms in prepuberty in 64% [36] and 89% (UI) of the patients. In the study of Seybold et al. [13] which, unlike the other two studies, includes many patients over the age of 10, ocular presentation was seen in 47%. Ptosis was more commonly the onset symptom as compared to diplopia [36] (UI). The second mode of presentation was leg weakness. Onset with bulbar weakness was rare, not being present in any of the patients of Bundey [36] and UI. Peri- and Postpuberty. When these two stages are considered together, presentation was with ocular (about 40%), bulbar (about 20%) and lower extremity symptoms (about 20%), followed by extremity (both or upper extremities) and combined symptoms [36] (UI). The importance of leg weakness as an onset symptom in these stages was particularly noteworthy. Disease started with leg weakness in 24% of the prepubertal patients of Seybold et al. [13]; 88% of these were children with onset at 10–14 years. In the other two studies, of all juvenile patients with leg weakness onset 73% [36] and 96% (UI) were


117

in peri- and postpubertal stages. Rarely, difficulties in smiling or in closing the eyes accompanied leg weakness. Leg weakness was sometimes so severe as to result in frequent falls. In some of our patients, this was the only symptom for a fairly long time and predominated even when other symptoms were added so that differential diagnosis from muscular dystrophies which can also present at this age became an important issue. Oosterhuis [39] explains the frequent occurrence of leg weakness, which he notes as an onset symptom in myasthenics below the age of 30, on the basis of overactivity in young people putting an extra burden upon the lower extremities. Anti-Acetylcholine Receptor Antibodies and HLA Associations The seropositivity rate and the titer of anti-acetylcholine receptor (antiAChR) antibodies were found to be low in prepubertal children [19, 23]. Interestingly, some seronegative children became seropositive at or after puberty. In the postpubertal period, the seropositivity rate was not different from that in myasthenic adults [23]. Not differing substantially from Caucasians in the behavior of antibodies [7, 40], the Chinese showed a strong association with HLA-DR9 and Bw46 [41] whereas Caucasians showed no specific HLA association in the juvenile period. Thymoma Thymoma associated with MG was found to be very rare in the juvenile period, particularly in prepuberty [19, 42]. Isolated cases, usually in postpuberty, were reported (table 1). Associated Autoimmune Conditions Thyroid disease was the most frequently associated autoimmune disease in this age group [7, 12, 18] (UI). Rheumatoid arthritis, lupus erythematosus, diabetes mellitus, asthma and vitiligo were other cited diseases in the JMG studies. MG was not usually associated with other autoimmune conditions during prepuberty [19]. In the UI series, all 8 patients with thyroid disease and the 1 patient with rheumatoid arthritis were in peri- and postpuberty. Severity and Outcome Proper comparison of different studies is hampered by both assessment and selection biases. Different classifications and improvement measures [20, 21] or seemingly the same measures with different meanings attributed to similar categories have been used [14, 15]. Remission was defined by some as a state of total eradication of symptoms with no medications [6, 12, 14–16, 19] (UI) and by others as the same with medications [18]; others still [13] accepted


118

minor symptoms as part of remission. Another major problem was the different criteria in the selection for thymectomy [12, 22], such as exclusion of very young children or patients responding to steroids. In the UI study, 75% of thymectomized patients during peri- and postpuberty were moderately or severely involved while the corresponding figure was only 36% in nonthymectomized patients so that any comparison became meaningless. In addition, our more severely involved patients were usually kept on a low-dose, alternateday steroid maintenance treatment making it impossible to judge remissions. The confounding effect of immunosuppressants starting prior to thymectomy further complicated evaluations [43]. Seybold [22] found only 5 studies [6, 12, 13, 16, 20] which allowed proper assessment of the value of thymectomy in juvenile patients. We did include remission rates of thymectomized and nonthymectomized patients for all studies in table 1, but the reader should evaluate them with these reservations in mind. Prepuberty. Prepubertal MG was not always found to have a benign progression although it was associated with two benign features: a high percentage of purely ocular (Osserman class I) MG and long-lasting spontaneous remissions. While about three quarters of Chinese [7] and Japanese [37] children with MG had ocular symptoms only, this percentage was less but still high (about one quarter to one half ) in some Caucasian series [19] (UI). Spontaneous remissions were more frequent in prepuberty [6, 16] and lasted as long as 12–13 years [13] (UI). Patients with ocular MG were more likely to have spontaneous remissions [11]. The high remission rate made the evaluation of all treatment modalities difficult. When the disease progressed mildly, anticholinesterases were sufficient to keep it under control [15]. However, respiratory difficulties occurred in as many as 40% of the patients [12, 19], a fact which has to be kept in mind even when the disease appears to be mild. Also, an acute fulminating form has been described [11]. Immunosuppressant drugs and thymectomy have to be considered when the disease assumes a severe form. Some studies [14, 20, 44, 45] with a relatively large number of prepubertal patients found thymectomy beneficial, while others [6, 16] reported less favorable results as compared to peri- and postpuberty. A review of the results of several studies showed a lower remission rate after thymectomy as compared to spontaneous remission for children under 5 [6]. It is unlikely that more definite conclusions will emerge since the number of prepubertal children is small, but it is useful to know that thymectomy in very young children has no major deleterious effect on the immune system [22]. Overall, with immunosuppressants and/ or thymectomy, where necessary, the outcome is good in most patients [6, 12, 46]. With purely ocular MG, low-dose, alternate-day steroids can resolve the symptoms [35].


119

Peri- and Postpuberty. These patients are similar to myasthenics with early onset [19] with minor differences. In some studies, males were found to be less severely affected than females [6, 17] (UI). Thymectomy, done much more frequently at this stage, was considered to be beneficial by most authors [6, 20, 44, 47–49]. Despite the absence of randomized studies, probably unethical at this point, thymectomy has a firm place in the treatment of peri- and postpubertal JMG [39]. Remissions were more frequent with early thymectomy, done within 1–2 years of disease onset [6, 16], although thymectomy done later also produced satisfactory results [13, 14]. Again, the outcome is good with different modalities of treatment including immunosuppressants [6, 18] and is possibly superior to that in adults [12]. Conclusions Prepubertal MG differs from MG in other age groups in sex prevalence, racial characteristics, seropositivity, clinical severity and progression. Males are often as frequently affected as females at this stage. The predominant presentation is ocular and purely ocular forms are often seen. Both the percentage and the titer of positive anti-AChR antibodies are low. Thymoma associated with MG is extremely rare. A high remission rate is noteworthy, making evaluation of all treatment modalities very difficult. This benign feature has to be weighed against the fact that a fairly high percentage may have respiratory difficulties so that thymectomy and immunosuppressants have to be seriously considered. Although thymectomy appears to be effective in isolated cases, its value continues to be disputed among authors and the issue will be difficult to resolve because of the small number of patients. With appropriate treatment, the outcome is good. The Chinese and the Japanese show striking differences from the Caucasians in prepuberty. It is a period with the highest incidence of MG for both sexes and it is predominantly purely ocular. With the advent of puberty, the characteristics of MG are very similar to those of later fertile years. An important onset symptom of the peri- and postpubertal years is leg weakness and distinction from muscular dystrophies is of great importance. Only isolated cases of thymoma have been reported. The beneficial effect of thymectomy in peri- and postpuberty, particularly if done early, is stressed by many. The outcome is good with appropriate treatment including immunosuppressants.

Late-Onset MG Previous literature accepted the age of 40 as the beginning of the older age group [50]. Somnier et al. [33], based on epidemiological data, suggested


120

that onset age for LOMG should be set at 50 rather than earlier. Aarli [4] recently defined LOMG as MG without evidence of thymoma, occurring after the age of 50. In our attempt to adhere to this definition in selecting the relevant papers dealing with elderly myasthenics [8, 51–56], compiled in table 3, we encountered several difficulties: neither was the age limit uniformly defined, varying from 50 to 65 in the studies, nor was thymoma considered separately within the elderly population. Demographic Features The same design and the same references (for hospital-based and epidemiological studies) as in the JMG section were used. LOMG Studies (table 3). When larger studies were considered, patients over 50 years of age made up about 30% of the total patient population and those over 60 years about 20%. Males outnumbered females or FMR was 1. Hospital-Based Studies. Oosterhuis [50], compiling information from several large hospital-based studies, reported that 36–47% of the patients had disease onset at or above the age of 40 except in a Japanese study with only 22%. In the hospital-based studies listed in the JMG section, LOMG was more frequent in men as compared to women. Peak age of onset was in the 5th to 7th decades in men and in the 2nd to 3rd decades in women. In some European studies, there was a relatively constant distribution of onset age for men after the 1st decade [29] or the peak was at a younger age similar to women [25, 57]. Population-Based (Epidemiological) Studies. Most population-based studies of Caucasians showed that the highest incidence rates of all age groups occurred in the elderly (table 2). They confirmed what was already known for men, but the results for women were very surprising. Either the highest incidence rate for women was found to fall within older age brackets or a bimodal distribution was present with a second peak late in life. FMR was 1 in the elderly population. Chinese [7] and Japanese [37] populations were completely different from the Caucasians in that they did not show a late peak. Onset Symptoms The most frequent presenting symptoms were ocular, occurring as an isolated symptom in over 50% of the patients in the majority of the studies. Onset with bulbar symptoms followed this type of presentation. In the UI study, bulbar onset was more common in women as compared to men. Presentation with weakness in the extremeties only was uncommon in this age group. An interesting onset symptom, easily confused with other etiologies in this age group, was weakness in the neck (head drop) [55], occurring in 6% in the UI study.


121

Table 3. LOMG studies Authors

Number of Age years patientsa

FMR Classification

Positive anti- TXb AChR Ab % %

Good outcome (remission plus improvement)

q60

0.9

Evoli et al. [52]

37 160 (13 of 278)

0.6


94

27 (5)

78%

Donaldson et al. [8]

55 q50 (33 of 165)

0.4


82

38 (10)

87%

Antonini et al. [53]

25 160 (20 of 122)

0.7

Ocular: 8% 86 Generalized: 92%

4 (1)

92%

Schon et al. [54]

13 (59 of 22)

q60

0.4

85

8 (1)

Bille´-Turc et al. [55]

34 q65 (17 of 200)

0.6

Ocular: 15% 94 Generalized: 85%

6 (2)

68%

Slesak et al. [56]

113 (30 of ?)

1

Ocular: 32%

13 (10)

87–93

UI study

183 q50 (25 of 733)

0.7


16 (18)

81%

Herishanu et al. [51] 15 (45 of 33)

?60

0

86

FMR>Female to male ratio, TX>thymectomy. a % of total number is given in parentheses. b Number of thymomas is given in parentheses.

Anti-AChR Antibodies and HLA Associations That LOMG may be a different disease from early-onset MG has been suggested by several immunological and HLA antigen studies. HLA-A3 was found to be more frequent in older patients [58] and HLA-B8 in young female patients [59, 60]. A comparison of the characteristics of nonthymoma patients below and above the age of 40 [61] showed that the older patients had a unique profile: association with HLA-A3, B7, DRw2, lower anti-AChR antibody


122

titers, higher antistriated muscle antibodies, higher autoantibodies other than those known to be associated with MG and higher immune diseases other than MG. Although anti-AChR antibody titers were found to be low, the percentage of positive anti-AChR antibodies was not any lower in the elderly than in other subgroups [61–64], with values at or above 85% in the majority of the studies on LOMG (table 3). Thymoma Thymomas have the highest prevalence in older men and women [39]. The percentage of elderly patients with thymoma was about 5–10 % in the studies on LOMG (table 3). The incidence in older women was twice as much as that in older men in some studies [39] (UI). Associated Autoimmune Conditions Thyroid disease, particularly hypothyroidism, was the most common autoimmune disease associated with LOMG [8, 55]. Hashimoto thyroiditis was present in 6% of the UI patients. Rheumatoid arthritis, systemic lupus erythematosus, diabetes mellitus, pernicious anemia, ankylosing spondylitis and idiopathic thrombocytopenic purpura were other cited autoimmune disorders. In the UI study, there were 3 patients with asthma, 2 patients with pernicious anemia and 1 each with systemic lupus erythematosus, idiopathic thrombocytopenic purpura, pemphigus and psoriasis. Association with all autoimmune diseases was more frequent in those with the onset around the age of 60 or later (UI). Severity and Outcome About 10–30 of the patients were classified as ocular MG (Osserman class I) in the majority of the LOMG studies. Myasthenics with mild symptoms (Osserman classes 1 and 2a) made up about 30–50% of the patients. On the other hand, about 30% had severe MG. Thus, LOMG can be mild, but one has to be aware of the fact that it is a potentially very severe disease. There was no consensus about the effect of thymectomy in this age group although some authors reported a benefit [65, 66]. Thymectomy was performed only in a minority of the cases unless the patient had a thymoma so that in most studies remission and improvement can easily be attributed to the effect of immunosuppressants. With immunosuppressants, mainly steroids, 68–92% of the patients improved, including a minority who went into remission. We have noted as did Bille´-Turc et al. [55] that low-dose steroids can be very effective in selected mildly affected patients. Yet, high doses are necessary in potentially severe cases.


123

Complications of steroids in the elderly, particularly cataracts and infection [8], have been stressed by some [52], while others have not found the side effects to be different from those in young people [53]. Because of these reservations, azathioprine is increasingly used as an adjunct to steroids in the elderly [8, 56] (UI). Azathioprine has fewer side effects than those of steroids [39, 67]. Our experience with azathioprine in the elderly is also favorable. Conclusions Onset of MG in the elderly is more common than previously thought. While it is well known that the peak age of onset is in the older age groups in men, recent epidemiological studies have revealed a similar tendency in women or at least a bimodal distribution with a second peak late in life. FMR is close to the 1 in the older age group. The disease often starts with ocular symptoms. Onset with weakness in the extremeties is uncommon. An interesting onset symptom in LOMG is weakness in the neck. Although anti-AChR titers are low in the elderly, the seropositivity rate is similar to that in early-onset MG. A unique antigen and HLA profile suggests that LOMG may be a different form of the disease. The disease is mild in half the cases; however, it can be severe in a third. Treatment is in many ways similar to that in the other age groups. The possibility of response to low-dose steroids, yet the necessity to give high doses in potentially severe cases and the possible beneficial effects of thymectomy in selected cases should be considered. Azathioprine is increasingly used in the elderly. The prognosis is good with appropriate treatment.

References 1 2 3 4 5 6 7 8 9

Engel AG, Franzini-Armstrong C: Myology. New York, McGraw-Hill, 1994. Schwab R, Leland C: Sex and age in myasthenia gravis as critical factors in incidence and remission. JAMA 1953;153:1270–1273. Perlo V, Poskanzer D, Schwab R, Viets H, Osserman K, Genkins G: Myasthenia gravis: Evaluation of treatment in 1355 patients. Neurology 1966;16:431–439. Aarli JA: Late-onset myasthenia gravis: A changing scene. Arch Neurol 1999;56:25–27. Phillips L 2d, Torner J, Anderson M, Cox G: The epidemiology of myasthenia gravis in central and western Virginia. Neurology 1992;42:1888–1893. Andrews PI, Massey JM, Howard JF Jr, Sanders DB: Race, sex, and puberty influence onset, severity, and outcome in juvenile myasthenia gravis. Neurology 1994;44:1208–1214. Wong V, Hawkins BR, Yu YL: Myasthenia gravis in Hong Kong Chinese. 2. Paediatric disease. Acta Neurol Scand 1992;86:68–72. Donaldson DH, Ansher M, Horan S, Rutherford RB, Ringel SP: The relationship of age to outcome in myasthenia gravis. Neurology 1990;40:786–790. Christensen PB, Jensen TS, Tsiropoulos I, Sorensen T, Kjaer M, Hojer-Pedersen E, Rasmussen MJ, Lehfeldt E, de Fine Olivarius B: Incidence and prevalence of myasthenia gravis in western Denmark: 1975 to 1989. Neurology 1993;43:1779–1783.


124

10

11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31

32 33 34 35 36

Emilia-Romagna Study Group on Clinical and Epidemiological Problems in Neurology: Incidence of myasthenia gravis in the Emilia-Romagna region: A prospective multicenter study. Neurology 1998;51:255–258. Fenichel GM: Clinical syndromes of myasthenia in infancy and childhood. A review. Arch Neurol 1978;35:97–103. Millichap J, Dodge P: Diagnosis and treatment of myasthenia gravis in infancy, childhood, and adolescence: A study of 51 patients. Neurology 1960;10:1007–1014. Seybold ME, Howard FM Jr, Duane DD, Payne WS, Harrison EG Jr: Thymectomy in juvenile myasthenia gravis. Arch Neurol 1971;25:385–392. Ryniewicz B, Badurska B: Follow-up study of myasthenic children after thymectomy. J Neurol 1977; 217:133–138. Snead OC 3d, Benton JW, Dwyer D, Morley BJ, Kemp GE, Bradley RJ, Oh SJ: Juvenile myasthenia gravis. Neurology 1980;30:732–739. Rodriguez M, Gomez MR, Howard FM Jr, Taylor WF: Myasthenia gravis in children: Long-term follow-up. Ann Neurol 1983;13:504–510. ¨ zdirim E, Renda Y, Yalaz K, Aysun S, Topçu M, Topalog˘lu H: Myasthenia gravis in Anlar B, O childhood. Acta Paediatr 1996;85:838–842. Lindner A, Schalke B, Toyka KV: Outcome in juvenile-onset myasthenia gravis: A retrospective study with long-term follow-up of 79 patients. J Neurol 1997;244:515–520. Evoli A, Batocchi AP, Bartoccioni E, Lino MM, Minisci C, Tonali P: Juvenile myasthenia gravis with prepubertal onset. Neuromuscul Disord 1998;8:561–567. Adams C, Theodorescu D, Murphy EG, Shandling B: Thymectomy in juvenile myasthenia gravis. J Child Neurol 1990;5:215–218. Cavanagh N: The role of thymectomy in childhood myasthenia. Dev Med Child Neurol 1980;22: 668–674. Seybold ME: Thymectomy in childhood myasthenia gravis. Ann NY Acad Sci 1998;841:731–741. Andrews PI, Massey JM, Sanders DB: Acetylcholine receptor antibodies in juvenile myasthenia gravis. Neurology 1993;43:977–982. Tanner J, Davies P: Clinical longitudinal standards for height and height velocity for North American children. J Pediatr 1985;107:317–329. Simpson J: An evaluation of thymectomy in myasthenia gravis. Brain 1958;81:112–144. Simpson JF, Westerberg MR, Magee KR: Myasthenia gravis. An analysis of 295 cases. Acta Neurol Scand 1966;42:(suppl 23):21–27. Osserman KE, Genkins G: Studies in myasthenia gravis: Review of a twenty-year experience in over 1200 patients. Mt Sinai J Med 1971;38:497–537. Grob D, Brunner N, Namba T: The natural course of myasthenia gravis and effect of therapeutic measures. Ann NY Acad Sci 1981;377:652–669. Oosterhuis HJ: Observations of the natural history of myasthenia gravis and the effect of thymectomy. Ann NY Acad Sci 1981;377:678–690. Storm-Mathisen A: Epidemiology of myasthenia gravis in Norway. Acta Neurol Scand 1984;70: 274–284. Giagheddu M, Puggioni G, Sanna G, Tamburini G, Marrosu F, Rachele MG, Murgia B, Rosati G: Epidemiological study of myasthenia gravis in Sardinia, Italy (1958–1986). Acta Neurol Scand 1989;79:326–333. Sorensen T, Holm E-B: Myasthenia gravis in the county of Viborg, Denmark. Eur Neurol 1989; 29:177–179. Somnier FE, Keiding N, Paulson OB: Epidemiology of myasthenia gravis in Denmark. A longitudinal and comprehensive population survey. Arch Neurol 1991;48:733–739. Ferrari G, Lovaste MG: Epidemiology of myasthenia gravis in the province of Trento (northern Italy). Neuroepidemiology 1992;11:135–142. Andrews PI: A treatment algorithm for autoimmune myasthenia gravis in childhood. Ann NY Acad Sci 1998;841:789–802. Bundey S: A genetic study of infantile and juvenile myasthenia gravis. J Neurol Neurosurg Psychiatry 1972;35:41–51.


125

37

38 39 40

41 42 43

44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59

60 61 62

Fukuyama Y, Hirayama Y, Osawa M: Epidemiological and clinical features of childhood myasthenia gravis in Japan; in Satyoshi E (ed): Myasthenia gravis – Pathogenesis and Treatment. Tokyo, University of Tokyo Press, 1981, pp 19–27. Teng P, Osserman KE: Studies in myasthenia gravis: Neonatal and juvenile types. J Mt Sinai Hosp 1956;23:711–727. Oosterhuis HJGH: Myasthenia gravis. Groningen, Groningen Neurological Press, 1997. Chiu HC, Vincent A, Newsom-Davis J, Hsieh KH, Hung T: Myasthenia gravis: Population differences in disease expression and acetylcholine receptor antibody titers between Chinese and Caucasians. Neurology 1987;37:1854–1857. Chen WH, Chiu HC, Hseih RP: Association of HLA-Bw46DR9 combination with juvenile myasthenia gravis in Chinese. J Neurol Neurosurg Psychiatry 1993;56:382–385. Furman WL, Buckley PJ, Green AA, Stokes DC, Chien LT: Thymoma and myasthenia gravis in a 4-year-old child. Case report and review of the literature. Cancer 1985;56:2703–2706. Lindberg C, Andersen O, Larsson S, Oden A: Remission rate after thymectomy in myasthenia gravis when the bias of immunosuppressive therapy is eliminated. Acta Neurol Scand 1992;86:323– 328. Keynes G: Surgery of the thymus gland. Second (and third) thoughts. Lancet 1954;i:1197–1202. Hansson O, Johansson L, Stalberg E, Westerholm C: Thymectomy in juvenile myasthenia gravis. Neuropa¨diatrie 1972;3:429–436. Sarnat HB, McGarry JD, Lewis JE Jr: Effective treatment of infantile myasthenia gravis by combined prednisone and thymectomy. Neurology 1977;27:550–553. Fonkalsrud E, Herrmann C, Mulder D: Thymectomy for myasthenia gravis in children. J Pediatr Surg 1970;5:157–165. Youssef S: Thymectomy for myasthenia gravis in children. J Pediatr Surg 1983;18:537–541. Campbell J, Bisio J, Harrison M, Campbell T: Surgical treatment of myasthenia gravis in childhood. J Pediatr Surg 1983;18:857–861. Oosterhuis H: Myasthenia gravis. New York, Churchill Livingstone, 1984. Herishanu Y, Abramsky O, Feldman S: Myasthenia gravis in the elderly. J Am Geriatr Soc 1976; 24:228–231. Evoli A, Tonali P, Scoppetta C, David P: Myasthenia gravis in the elderly: Report of 37 cases. J Am Geriatr Soc 1983;31:352–355. Antonini G, Morino S, Gragnani F, Fiorelli M: Myasthenia gravis in the elderly: A hospital based study. Acta Neurol Scand 1996;93:260–262. Schon F, Drayson M, Thompson R: Myasthenia gravis and elderly people. Age Ageing 1996;25: 56–58. Bille´-Turc F, Padovani R, Pouget J, Serratrice G, Bille´ J: La myastheńie de re´ve´lation tardive: 34 cas chez des patients de plus de 65 ans. Rev Neurol (Paris) 1997;153:129–134. Slesak G, Melms A, Gerneth F, Sommer N, Weissert R, Dichgans J: Late-onset myasthenia gravis. Follow-up of 113 patients diagnosed after age 60. Ann NY Acad Sci 1998;841:777–780. Hokkanen E: Myasthenia gravis. A clinical analysis of the total material from Finland with special reference to endocrinological and neurological disorders. Ann Clin Res 1969;1:94–108. Fritze D, Herrman C, Naeim F, Smith GS, Walford RL: HLA antigens in myasthenia gravis. Lancet 1974;i:240–242. Feltkamp T, Van Den Berg-Loonen P, Nijenhuis L, Oosterhuis H, Engelfriet C, Van Rossum A, Van Loghem J: Relations between HL-A antigens, myasthenia gravis and autoantibodies; in Kunze K, Desmedt J (eds): Studies on Neuromuscular Diseases. Proc Int Symp Giessen 1973. Basel, Karger, 1975, pp 180–185. Pirskanen R: On the significance of HL-A and LD antigens in myasthenia gravis. Ann NY Acad Sci 1976;274:451–460. Compston DA, Vincent A, Newsom-Davis J, Batchelor JR: Clinical, pathological, HLA antigen and immunological evidence for disease heterogeneity in myasthenia gravis. Brain 1980;103:579–601. Lindstrom JM, Seybold ME, Lennon VA, Whittingham S, Duane DD: Antibody to acetylcholine receptor in myasthenia gravis. Prevalence, clinical correlates, and diagnostic value. Neurology 1976; 26:1054–1059.


126

63

64

65 66 67

Morel E, Raimond F, Goulon-Goeau C, Berrih S, Vernet-Der-Garabedian B, Harb J, Bach JF: Le dosage des anticorps anti-rećepteurs de l’ace´tylcholine dans la myastheńie. Etude de 329 se´rums. Nouv Presse Me´d 1982;11:1849–1864. Vincent A, Newsom-Davis J: Acetylcholine receptor antibody as a diagnostic test for myasthenia gravis: Results in 153 validated cases and 2967 diagnostic assays. J Neurol Neurosurg Psychiatry 1985;48:1246–1252. Olanow CW, Lane RJ, Roses AD: Thymectomy in late-onset myasthenia gravis. Arch Neurol 1982; 39:82–83. Monden Y, Nakahara K, Fujii Y, Hashimoto J, Ohno K, Masaoka A, Kawashima Y: Myasthenia gravis in elderly patients. Ann Thorac Surg 1985;39:433–436. Drachman DB: Myasthenia gravis. N Engl J Med 1994;330:1797–1810.

Feza Deymeer, MD, Department of Neurology, University of Istanbul, C ¸ apa 34390, Istanbul (Turkey)


127


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

Hereditary Peripheral Neuropathies Peter De Jonghe a, b, Vincent Timmerman a, Eva Nelis a a

b

Flanders Interuniversity Institute for Biotechnology (VIB), Born-Bunge Foundation (BBS), Department of Biochemistry, University of Antwerp (UIA) and Department of Neurology, University Hospital Antwerp (UZA), Antwerp, Belgium

At the end of the 1960s Dyck et al. [1] classified the inherited neuropathies of the peripheral nervous system based on mode of inheritance, clinical features, neuropathological and electrophysiological findings. Dyck and coworkers distinguished three large groups, i.e. hereditary motor and sensory neuropathies (HMSN), hereditary motor neuropathies (HMN) and hereditary sensory neuropathies (HSN) or hereditary sensory and autonomic neuropathies (HSAN). Each category is further subdivided into several types. Molecular genetic studies have confirmed this extensive heterogeneity. Currently, 22 inherited peripheral neuropathy loci have been mapped (table 1). However, the genetic loci for many types still remain to be found and one can safely predict that there must exist between fifty and one hundred genetically distinct types. This huge number of loci stands in sharp contrast to the small number of only four genes that, so far, have been found to be involved in the inherited peripheral neuropathies. These genes are: peripheral myelin protein 22 gene (PMP22) at chromosome 17p11.2, myelin protein zero gene (MPZ, P0) at chromosome 1q22-q23, connexin 32 gene (Cx32,GJB1) at chromosome Xq13 and the early growth response element 2 gene (EGR2) at chromosome 10q21.1q22.1. At present, 332 different mutations within these four genes have been detected, leading to a wide variety of clinical phenotypes [2]. A database, containing all published and a number of unpublished mutations, can be consulted at http://molgen-www.uia.ac.be/CMTmutations/. The results of functional studies of the mutated proteins in human biopsy specimens, cellular and animal models have been the subject of recent review papers and will not be discussed here [3, 4]. We will focus on the inherited neuropathies in which molecular genetics have provided new insights.

Clinical Phenotypes Most patients affected by an inherited peripheral neuropathy have a clinical phenotype closely resembling the patients originally described by Charcot and Marie [5] and Tooth [6] in 1886. The eponym Charcot-Marie-Tooth disease or CMT is still commonly used to designate this phenotype [7]. The disease usually starts in the 2nd decade of life with a progressive paresis of the peroneal muscles. Several years later some weakness of the intrinsic hand muscles also develops. Sensory symptoms are usually discrete or absent and tendon reflexes are often abolished. Skeletal abnormalities such as pes cavus or scoliosis are frequently observed. The severity of the disease is variable. Most patients are able to live an independent life even in old age while other patients become wheelchair-bound due to the involvement of the proximal muscles of the legs. Additional features may be present such as deafness, pupillary abnormalities, vocal cord paralysis, and poorly healing ulcers [8]. A second phenotype is the De´jerine-Sottas syndrome (DSS) [9]. Some authors have proposed to reserve this eponym for patients with a severe, early onset demyelinating neuropathy [10]. They further specified that nerve conduction velocities (NCV) should be severely reduced to less than 12 m/s and the protein content of the cerebrospinal fluid should be increased. This syndrome is usually considered to be inherited as an autosomal recessive (AR) trait although most of the patients do not have affected relatives [7]. In the recent literature, DSS is often used to designate a severe, early onset inherited peripheral neuropathy without any reference to pathology, NCVs or mode of inheritance. The difference between DSS and severe CMT is therefore mainly a matter of taste. A third, rare phenotype is congenital hypomyelination (CH). CH patients have a congenital onset of the disease with breathing and feeding problems. The evolution is often rapidly progressive with a fatal outcome in infancy. CH is, by definition, a demyelinating neuropathy. Molecular genetic studies have shown that each of these phenotypes can be caused by mutations in different genes and therefore should be considered as syndromes and not as separate disease entities [2].

Classification of HMSN The HMSN are a heterogeneous group of disorders in which both motor and sensory nerves are affected. The HMSNs are subdivided into seven subtypes based on clinical phenotype, mode of inheritance, electrophysiological and neuropathological characteristics [7]. We will only discuss types I, II and III since molecular genetics have made progress in these types. HMSN type I, also called CMT1, and HMSN type III or DSS are demyelinating neuropathies

Hereditary Peripheral Neuropathies

129

Table 1. Loci for CMT and related peripheral neuropathies Locus

Chromosome Mutation/gene

Autosomal dominant Charcot-Marie-Tooth type 1 (CMT1 – HMSN type I ) CMT1A 17p11.2 1.5-Mb tandem duplication/ dosage of PMP22 CMT1A 17p11.2 PMP22 mutations CMT1B 1q22-23 MPZ mutations CMT1C ? ? CMT1 10q21.1-22.1 EGR2 mutation Autosomal recessive Charcot-Marie-Tooth type 1 (CMT1 – HMSN type I ) CMT4A 8q13-21 ? CMT4B 11q23 ? CMT4 5q31-33 ? HMSN-L 8q24 ? Dominant X-linked Charcot-Marie-Tooth type 1 (CMT1 – HMSN type I ) CMT1X Xq13.1 Cx32 mutations Autosomal dominant Charcot-Marie-Tooth type 2 (CMT2 – HMSN type II ) CMT2A 1p35-36 ? CMT2B 3q13-22 ? CMT2C ? ? CMT2D 7p14-15 ? Autosomal recessive Charcot-Marie-Tooth type 2 (CMT2 – HMSN type II ) CMT2 1q21.2-21.3 ? Axonal motor-sensory neuropathy with deafness and mental retardation CMT2X Xq24-26 ? De´jerine-Sottas Syndrome (DSS – HMSN type III ) DSS 17p11.2 DSS 1q22-23 DSS 10q21.1-22.1 AD-DSS 8q23-24

PMP22 mutations MPZ mutations EGR2 mutations ?

Congenital hypomyelination (CH ) CH CH CH

MPZ mutation EGR2 mutations PMP22 mutations

1q22-23 10q21.1-22.1 17p11.2

Distal hereditary motor neuropathy (Distal HMN ) Distal HMN II 12q24 Distal HMN V 7p Congenital non-progressive distal HMN 12q23-24 Recessive distal HMN 9p12-21

De Jonghe/Timmerman/Nelis

? ? ? ?

130

Table 1 (continued) Locus

Chromosome Mutation/gene

Hereditary sensory neuropathy (HSN ) HSN-I

9q22

?

Congenital cataracts facial dysmorphism neuropathy (CCFDN ) CCFDN 18qter ? Hereditary neuropathy with liability to pressure palsies (HNPP ) HNPP 17p11.2 1.5-Mb deletion/dosage of PMP22 HNPP 17p11.2 PMP22 mutations Hereditary neuralgic amyotrophy (HNA ) or familial brachial plexus neuropathy HNA 17q25 ?

that show signs of de- and remyelination on nerve biopsy studies. CMT1 presents with a classical CMT phenotype and can be inherited as an autosomal dominant (AD), AR or X-linked trait. HMSN type III patients have the more severe DSS phenotype. Since many of the DSS patients have normal parents, it was suggested that DSS is mainly inherited as an AR trait. HMSN type II or CMT2 is an axonal neuropathy showing loss of nerve fibers and signs of regeneration without overt signs of de- and remyelination. CMT2 patients have a classical CMT phenotype and cannot be distinguished from CMT1 patients on clinical examination only. Electrophysiological studies demonstrate that CMT2 patients have normal or slightly reduced NCVs while CMT1 patients have severely reduced NCVs, usually less than 38 m/s for the motor median nerve [11]. In most families all patients can be diagnosed as either CMT1 or CMT2. However, some families seem to have a puzzling mixture of CMT1 and CMT2 patients and for these families the term ‘intermediate CMT’ was proposed without gaining wide acceptance [12].

CMT1 AD CMT1 AD CMT1 represents the most common form of the inherited peripheral neuropathies. Molecular genetic studies have identified mutations in the genes coding for peripheral myelin protein 22 (PMP22) at chromosome 17p11.2 (CMT1A) [13,14], myelin protein zero (MPZ, P0) at chromosome 1q22-q23 (CMT1B) [15] and the early growth response element 2 gene (EGR2) at chromo-


131

some 10q21.1-q22.1 (the name of this locus is still pending) [16] as causes for the majority of AD CMT1 cases. Some AD CMT1 families have been excluded for linkage to the known AD CMT1 loci (preliminarily designated CMT1C) [17]. CMT1A. The majority of AD CMT1 families showed linkage to chromosome 17p11.2 (CMT1A) [18]. In 1991, two research groups independently demonstrated that some CMT1A patients have a 1.5-Mb tandem duplication in one of their chromosome 17 homologues [13, 14]. This duplication contains the PMP22 gene [19, 20]. Occasionally alternatively sized duplications are found but they always contain the PMP22 gene [21]. The detection of a PMP22 point mutation in Trembler mice provided solid evidence that PMP22 is indeed the CMT1A gene [22]. Epidemiological studies showed a high prevalence of the 1.5-Mb CMT1A tandem duplication, reaching 71% in very rigorously selected unrelated CMT1 families [23]. The CMT1A duplication is therefore, by far, the most frequent mutation in CMT1. An unexpected finding was the high rate of de novo CMT1A duplications [24]. Diagnosing a CMT1A de novo mutation has important consequences for genetic counseling since it will be transmitted as an AD trait to the next generations. Some CMT1A families do not have the CMT1A duplication but carry mutations in the PMP22 gene [25, 26]. These PMP22 mutations are present in the heterozygous state and are thus AD-inherited. Many of these PMP22 mutations, resulting in a severe DSS phenotype, are de novo mutations [27]. Recently, a recessive PMP22 mutation was reported in DSS siblings born to normal consanguineous parents who were heterozygous for this mutation [28]. So far 39 distinct PMP22 mutations have been observed. These include three polymorphisms, 29 mutations leading to CMT1A or DSS and seven mutations associated with hereditary neuropathy with liability to pressure palsies (HNPP), a disorder that will be discussed later in this chapter. CMT1A duplication patients have a classical CMT phenotype. The severity varies between families, within families and even between monozygous twins [29]. Most patients are able to lead an independent life even at an old age. In general, CMT1A with the duplication can be considered to be mild to moderately severe. A few patients homozygous for the CMT1A duplication have been described [13, 30]. They usually have a more severe phenotype. All patients, regardless of the severity of the clinical phenotype, have slowed NCVs, which is thus a completely penetrant trait. Therefore, NCV testing is a 100% reliable test to confirm or to rule out the disease in CMT1A families. As we will discuss later this finding cannot be extrapolated to CMT1B and CMT1X. NCVs are severely slowed to 20–30 m/s. However, a few of our CMT1A duplication patients had a NCV in the motor median nerve of 42 m/s. These exceptional cases would be misdiagnosed using a strict cutoff value of 38 m/s


132

to differentiate between CMT1 and CMT2. Follow-up studies show that NCVs do not dramatically change over a long period of time [31]. Also, there is a rather poor correlation between severity of the disease and NCV slowing [32]. CMT1A patients with PMP22 mutations usually have a more severe phenotype and several patients with a DSS phenotype have been reported. Neuropathological studies in CMT1A, due to the CMT1A duplication or PMP22 mutations, show loss of large myelinated fibers and the presence of classical onion bulbs consisting of concentric layers of Schwann cell processes. Recently, biopsies of two family members with a peculiar PMP22 mutation showed loosening of the compaction of myelin sheaths resembling the abnormalities seen in some patients with MPZ mutations [33]. CMT1B. CMT1B is caused by mutations in the myelin protein zero gene (MPZ, P0) localized at chromosome 1q22-23. So far, 69 distinct diseasecausing mutations and 3 polymorphisms have been reported in MPZ. All the disease-causing mutations behave like AD mutations and lead to clinical symptoms when present in the heterozygous state. A few homozygous patients, born to consanguineous CMT1B parents, have been reported. The parents were asymptomatic or had a mild phenotype while the homozygous children were severely affected [34, 35]. Recently, a case of germ-line mosaicism was reported for a MPZ mutation. The mutation was present in 2 affected children and absent in the DNA extracted from lymphocytes of both parents [36]. MPZ mutations are associated with a variety of clinical phenotypes, i.e. classical CMT1, DSS, CH and surprisingly a CMT2-like phenotype [2]. Initially, MPZ mutations were detected in families with a classical CMT1 phenotype and this is still the most common phenotype associated with MPZ mutations. On average, CMT1B patients are somewhat more severely affected than CMT1A patients with the 1.5-Mb CMT1A duplication. Some MPZ mutations result in a DSS phenotype. These DSS patients usually represent de novo MPZ mutations since reproductive fitness is decreased in these severely affected individuals. Also, CH patients due to MPZ mutations have been described [34]. All these CMT1B, DSS and CH patients with MPZ mutations have severely slowed NCVs to less than 38 m/s. Two distinct neuropathological phenotypes have been described in CMT1B patients [37]. Some biopsies show classical onion bulbs consisting of concentric layers of myelin sheaths while decompaction of the myelin sheaths is a striking feature in others. Surprisingly, MPZ mutations can also be associated with a CMT2 like phenotype, i.e. slightly reduced to normal NCVs and neuropathological features of an axonal neuropathy. The first two CMT1B families with a CMT2 phenotype were actually reported for other reasons and their peculiar phenotype remained almost unnoticed [34, 35]. In both instances, these patients were the consanguineous parents of severely affected children who were homozygous for an


133

MPZ mutation. In 1998, Marrosu et al. [38] reported a large Italian family with a CMT2 phenotype due to a MPZ mutation. Subsequently, other MPZ mutations with a similar phenotype have been described, including several families affected by the peculiar Thr124Met mutation [39, 40]. Interestingly, all families with the Thr124Met mutation have a similar CMT2 phenotype characterized by late onset age, severe gait disturbances, lancinating pains, pupillary abnormalities and deafness [39]. Neuropatholocial examination of sural nerve biopsy specimens showed clusters of regenerating fibers in the absence of myelin abnormalities. NCVs ranged from normal values in asymptomatic young mutation carriers to severely reduced values in older patients. EGR2. Recently, Warner et al. [16], using a candidate gene approach, detected three mutations in the early growth response 2 (EGR2) gene localized at chromosome 10q21.1-q22.1, in patients with a CMT1 and CH phenotype. Some patients were heterozygous for a mutation while others were homozygous. Subsequently a fourth EGR2 mutation was reported in a DSS patient [41]. At present six distinct EGR2 mutations have been reported [2]. X-Linked CMT1 The X-linked form of CMT1 (CMT1X) is caused by mutations in the Cx32 gene localized at chromosome Xq13 [42]. So far, 221 distinct diseasecausing mutations and 7 polymorphisms in the coding region have been reported [2]. In two families, pathogenic mutations in the promoter region of Cx32 have been found. Most CMT1X patients have a positive family history, but a genuine de novo mutation has recently been described [43]. All Cx32 mutations seem to result in a CMT phenotype although marked variability in severity exists between and within families. In general, male patients are more severely affected than females but female patients can be severely affected and become wheelchair-dependent. At least in male patients, CMT1X is more severe than the AD forms of CMT1. A subclinical involvement of central nervous system (CNS) pathways has been detected by brainstem auditory evoked potentials CMT1X families [44]. It has been suggested that the detection of these CNS abnormalities by brainstem auditory evoked potentials can help to select families for mutation screening of the Cx32 gene. Surprisingly Cx32 mutations are the second most common mutation in CMT1 only surpassed by the CMT1A duplication. The explanation for this discrepancy is the fact that the presence of severely affected females can easily obscure the X-linked inheritance pattern. Only the absence of a male-to-male transmission then points to a possible X-linked mode of inheritance. NCV studies in large families with Cx32 mutations have finally solved some controversies in the classification of CMT types. Male CMT1X patients have reduced motor and sensory NCVs but values higher than 38 m/s for the motor median nerve are


134

common. NCVs in female patients or female mutation carriers can be normal or slightly reduced. Based on a cutoff value of 38 m/s for the motor median nerve, some patients (especially males) will be diagnosed as CMT1 and others (mainly females) as CMT2 [45]. In retrospect, most of the intermediate CMT families turned out to be X-linked families with Cx32 mutations. Neuropathological studies in CMT1X have shown a mixture of axonal pathology and the presence of small onion bulbs [46]. AR CMT1 AR CMT1 is rare in outbred populations but large pedigrees have been studied in populations with a high rate of consanguineous marriages such as North Africa. An AR CMT1 type has been described in Gypsies (HMSN-L), an ethnically isolated group. AR CMT1 patients are, in general, more severely affected than AD CMT1 patients. Additional features such as deafness and scoliosis seem to be an integral part of the phenotype in some AR CMT1 types. All patients with AR CMT1 have severely slowed NCVs. The neuropathology of AR CMT1 seems to be more complex than in AD CMT1. Besides classical onion bulbs consisting of concentric Schwann cell lamellae, other, previously unrecognized alterations of the myelin sheaths have been described such as myelin outfoldings and basal lamina onion bulbs. Four loci for AR CMT1 have been mapped but the genes involved are not known. To complicate matters, the designation CMT4 is used for the AR CMT loci. The AR CMT1 loci are: CMT 4A at chromosome 8q13-q21, CMT4B at chromosome 11q23, a locus at chromosome 5q31-33 the name of which is pending and HMSN-L at chromosome 8q24. CMT4A. The first AR CMT1 locus (CMT4A) was mapped to chromosome 8q13-q21 in inbred Tunisian families [47]. The clinical phenotype is severe early onset CMT. Many patients in these families became wheelchairdependent. Nerve biopsy studies showed the presence of classical onion bulbs. CMT4B. The CMT4B locus has been mapped to chromosome 11q23 in a large inbred Italian family [48]. Patients have a severe early onset CMT. Nerve biopsy examination shows irregular outfoldings of the myelin sheaths. AR CMT1 with myelin outfoldings is a genetically heterogeneous disorder since some families with this phenotype have been excluded for linkage to the CMT4B locus [49]. AR CMT1 with Basal Lamina Onion Bulbs. LeGuern et al. [50] mapped a third AR CMT1 locus to chromosome 5q23-33 in Algerian inbred families. They refrained from giving a name to this new locus. The clinical phenotype in these families is very peculiar. Patients develop a severe scoliosis as the presenting sign of the disease. Only later they develop signs of a peripheral neuropathy. Onion bulbs consisting of basal laminae are found in nerve biopsies [51].


135

HMSN-L. This AR CMT1 locus has been described in Bulgarian Gypsy families and maps to chromosome 8q24 [52]. To make HMSN classifications even more confusing, the name of this new locus refers to the town Lom were these families live. Patients have classical CMT but hearing loss is a constant additional feature.

HMSN Type III or DSS HMSN type III or DSS is a genetically heterogeneous group of disorders. There is a tendency in the recent literature to use the terminology of DSS as a mere synonym of early onset severe CMT. We have already discussed that mutations in PMP22, MPZ and EGR2 can lead to DSS. These patients are usually isolated and represent de novo cases of dominant mutations in these genes. Occasionally, DSS patients are homozygous for the CMT1A duplication or mutations in MPZ [2]. Recently, the first example of an MPZ germ-line mosaicism was reported in a DSS sib [36]. A locus for AD DSS was assigned to 8q23-q24 in a single pedigree [53]. No mutations in the known CMT genes were found in a small AD DSS family that could also be excluded for linkage to the known loci [54]. This implies further genetic heterogeneity in this syndrome.

CMT2 CMT2 is usually inherited in an AD or AR mode [7]. An X-linked CMT2 type has been described but additional features, such as hearing loss and mental retardation, were also present [55]. Molecular genetic studies in CMT2 have been much less productive than in CMT1. One of the reasons, for which we have no obvious explanation, is that large CMT2 families suitable for linkage studies seem to be much rarer than extended CMT1 pedigrees. A major disadvantage for linkage studies in CMT2 is the lack of a reliable biological marker. In CMT1 families, NCV studies can score each individual, including asymptomatic young persons, as either affected or unaffected since this trait is 100% penetrant in most CMT1 variants. In CMT2, NCVs can still be normal in clinically affected individuals. Molecular genetic studies have demonstrated that CMT2 is very heterogeneous. AD CMT2 Three AD CMT2 loci have been mapped: CMT2A at chromosome 1p35p36 [56], CMT2B at chromosome 3q21–q22 [57] and CMT2D at chromosome 7p14 [58]. CMT2C is reserved for the locus of AD CMT2 with vocal cord


136

paralysis [59]. These CMT2C families have been excluded for linkage to the AD CMT2 loci. No genes for CMT2 have been found so far. For the clinician it is very important to keep in mind that the rigorous use of a cutoff value of 38 m/s to differentiate between CMT1 and CMT2 will classify some CMT1A, CMT1B and CMT1X patients as CMT2 as we have already mentioned. Therefore, we screen our CMT2 patients for MPZ and Cx32 (if no male-to-male transmission is present) mutations [39, 60]. CMT2A. The first AD CMT2 locus (CMT2A) was mapped to chromosome 1p35-36 [56] in 1993. CMT2A is associated with a classical CMT phenotype. CMT2B. In 1995, Kwon et al. [57] reported linkage to chromosome 3q13q22 in a family with an axonal neuropathy and both sensory and motor deficits. They designated this new locus CMT2B. However, sensory abnormalities were much more pronounced than one normally observes in CMT2 patients. Several patients in the family of Kwon et al. had poorly healing ulcers that necessitated amputations of distal parts of the limbs. Linkage to the CMT2B locus has been confirmed in a second family with a similar phenotype [61]. CMT2D. The CMT2D locus was mapped to chromosome 7p14 [58]. A striking feature in these CMT2D families is the unusual evolution and distribution of the motor symptoms. Unlike other CMT types, which invariably start with weakness in the feet, CMT2D begins in the hands where weakness still predominates later in the evolution of the disease. This unusual phenotype was observed in the families that were originally used to map the CMT2D locus. However, it might be that also families with classical CMT2 map to the same locus. AR CMT2 Recently, the first locus for AR CMT2 was mapped to chromosome 1q21.2-q21.3 in a large consanguineous Moroccan family [62]. Patients had a severe neuropathy sometimes with proximal involvement. X-Linked CMT2 A locus for X-linked CMT2 has also been mapped to chromosome Xq2426 [55]. The phenotype, however, is not a pure axonal peripheral neuropathy since additional features such as deafness and mental retardation are also present, suggesting a concomitant involvement of the CNS.

Distal Hereditary Motor Neuropathies The distal HMN mainly present with a classical CMT phenotype. The main discriminative clinical factor between CMT1/CMT2 and distal HMN is


137

the absence of sensory symptoms and signs in the latter. Sensory signs, however, are often discrete in CMT1 and CMT2 and, therefore, testing of the sensory NCVs is obligatory to rule out subclinical sensory involvement. Neuropathological findings in distal HMN are scarce and demonstrate an axonal neuropathy of the motor nerves while the sensory nerves remain intact. Motor NCVs in distal HMN are normal or slightly reduced when severe muscle atrophy is present. Sensory NCVs are normal and the sensory nerve action potentials have normal amplitudes. Concentric needle EMG shows chronic neurogenic alterations. This latter observation is important since some distal myopathies can mimic distal HMN. Distal HMN is clinically and genetically very heterogeneous and a classification into seven subtypes has been made based on mode of inheritance, age at onset, severity and the presence of additional symptoms [63]. This already extensive classification, however, is not complete and some clinical entities were not included or have been described later on [64–66]. Distal HMN is usually inherited in an AD or AR mode. X-linked families have not been described so far. Molecular genetic studies have recently confirmed that these clinical distinct phenotypes represent separate genetic entities. Distal HMN Type II The locus for distal HMN type II also known as spinal CMT was mapped to chromosome 12q24 in a single Belgian family [67, 68]. The disease starts between the age of 15–20 years and presents with a classical, though rather severe CMT phenotype since some older patients become wheelchairbound. Distal HMN Type V A locus for distal HMN type V was localized on chromosome 7p14 in a single large Bulgarian family [69]. The linkage region shows a large overlap with the CMT2D region. Interestingly, distal HMN type V also starts in the hands where it continues to predominate. Families with a similar phenotype have been excluded from this region indicating genetic heterogeneity [70]. Congenital Nonprogressive Distal HMN This congenital form of distal HMN was not included in the original distal HMN classification. Its locus was recently mapped to chromosome12q23-q24. Although distal HMN type II also maps to chromosome 12q24 there is no overlap between these two linkage regions [71]. This dominantly inherited disorder presents at birth and is associated with arthrogryposis. The course is benign with almost no progression [65, 66].


138

Recessive Distal HMN The first locus for AR distal HMN was mapped to chromosome 9p21p12 in Jordanian families from the Jerash region [72]. Clinical features include pyramidal signs within the early stages of the disease pointing to an additional involvement of CNS pathways.

Hereditary Sensory Neuropathies The HSN also called HSAN are rare diseases [73]. Clinically they present with prominent sensory abnormalities sometimes leading to poorly healing ulcers that result in osteomyelitis and eventually necessitate the amputation of distal parts of the limbs. Good foot care can largely prevent these complications. Varying degrees of autonomic disturbances may be present. The HSNs are mostly axonal neuropathies. Electrophysiologically, the sensory nerve action potentials have reduced amplitudes or are not obtainable. A classification in several subtypes has been proposed based on clinical and genetic data. Both AD and AR variants have been reported. AD HSN Type I Molecular genetic linkage studies have mapped the gene for AD HSN type I to chromosome 9q22.1-q22.3 in Australian families [74]. These families originate from England and the ancestors were deported as convicts. Patients in these chromosome 9-linked HSN type I families have striking sensory abnormalities and occasionally develop poorly healing ulcers in the distal parts of the limbs. However, most of the patients also have some weakness of the distal muscles of the legs. Therefore, HSN type I is not an exclusively sensory neuropathy. There must exist a third locus for AD HSN or AD CMT2 with prominent sensory signs since a large family with an ulceromutilating neuropathy was excluded for linkage to the CMT2B and HSN I locus [75].

Hereditary Recurrent Neuropathies Hereditary Neuropathy with Liability to Pressure Palsies HNPP is an AD-inherited peripheral neuropathy characterized by recurrent palsies of individual nerve trunks [76, 77]. The palsies are typically painless and both motor and sensory nerves can be involved. Peroneal, ulnar and radial nerve palsies are common in HNPP. Palsies usually follow minor trauma to the affected nerve such as prolonged pressure at common entrapment sites. Recovery is often complete within minutes or months although some patients develop residual


139

deficits. Molecular genetic studies have shown that HNPP is a genetically homogeneous disorder linked to the CMT1A locus at chromosome 17p11.2. Most HNPP patients have a 1.5-Mb deletion in the region that is duplicated in CMT1A patients. Thus, most HNPP patients have only one copy of the PMP22 gene [78]. Only a few HNPP patients have a point mutation in PMP22. These mutations are either nonsense mutations leading to truncated protein or frameshift mutations that result in a severely altered and probably nonfunctional PMP22 protein [2]. Initially, it was suggested that HNPP is genetically heterogeneous since some families were excluded for linkage to chromosome 17p11.2 [79]. However, reanalysis of the data showed that two different peripheral neuropathies were segregating in a single family. Family members with HNPP were subsequently shown to harbor a PMP22 point mutation [70]. Extensive clinical and electrophysiological studies of large groups of patients, diagnosed as HNPP based on DNA testing, have recently been reported [80]. Four clinical phenotypes emerge. Most patients have a typical HNPP phenotype with recurrent palsies. However, almost 30% of mutation carriers remain asymptomatic. We have also seen patients who almost immediately developed paraesthesias after slight pressure but never had prolonged motor or sensory deficits. Finally, some HNPP patients have a progressive sensorimotor neuropathy that closely resembles CMT. Electrophysiological studies have clearly demonstrated that there exists a widespread involvement in HNPP. This is an important diagnostic tool to differentiate HNPP from acquired mononeuropathies. More than 90% of HNPP patients have prolonged terminal latencies of the motor median nerve and reduced sural nerve conductions even in the absence of clinical signs of damage to these nerves. Nerve biopsies in HNPP show focal myelin thickening called ‘tomacula’. HNPP patients with PMP22 point mutations tend to have more severe clinical, electrophysiological and neuropathological abnormalities although there exists a large overlap with HNPP due to the common 1.5-Mb deletion [81]. Hereditary Neuralgic Amyotrophy Hereditary neuralgic amyotrophy (HNA) is a rare disorder characterized by recurrent episodes of brachial plexus palsies [77]. In contrast to HNPP, these HNA episodes are always preceded by intense pain in the affected arm. Intriguingly, HNA episodes are often triggered by recent viral infections, immunizations or parturition suggesting some autoimmune disease mechanism. Motor symptoms predominate but minor sensory symptoms may be present. Occasionally the lumbar plexus, the phrenic nerve or individual cranial nerves can be involved. Symptoms usually recover within weeks to months. Recovery is usually good but can be incomplete. The number of episodes ranges from a single episode to more then ten attacks. The attacks can be uni- or bilateral. HNA is inherited as an AD trait. Dysmorphic features such


140

as epicanthus, hypotelorism, cleft palate and short stature have been described in some HNA families but these features usually do not cosegregate with the HNA trait [77]. The HNA gene was recently mapped to chromosome 17q25 [82, 83]. The HNA gene, however, is not yet found. Most HNA families show linkage to the chromosome 17q25 HNA locus but some families have been excluded. HNA is thus a genetically heterogeneous disorder.

Conclusions Molecular genetic investigations had a direct impact on the study of inherited peripheral neuropathies by mapping the disease loci and identifying causative gene mutations. This paved the way for DNA diagnosis that can now be offered to patients and their relatives. DNA diagnosis is essential for prenatal and preimplantation diagnoses [84, 85]. Indirectly, these molecular genetic studies have greatly enhanced our knowledge about the variability in clinical, electrophysiological and neuropathological features. Initially, the need for large families or homogeneous groups of families, suitable for linkage studies, has forced researchers to carefully document the phenotypes in their patients and families. Later on, genotype-phenotype correlations in patients with known mutations or families with proven linkage to a certain locus made it possible to delineate new disease entities.

Acknowledgments Our research is funded by grants of the Fund for Scientific Research – Flanders (FWO), the Geneeskundige Stichting Koningin Elisabeth (GSKE), a Special Research Fund of the University of Antwerp, the Association Française contre les Myopathies (AFM, France), and the Muscular Dystrophy Association (MDA, USA). V.T. and E.N. are research assistants of the FWO.

References 1

2 3 4

Dyck PJ: Inherited neuronal degeneration and atrophy affecting peripheral motor, sensory and autonomic neurons; in Dyck PJ, Thomas PK, Lambert EH (eds): Peripheral Neuropathy. Philadelphia, Saunders, 1984, pp 1600–1642. Nelis E, Haites N, Van Broeckhoven C: Mutations in the peripheral myelin genes and associated genes in inherited peripheral neuropathies. Hum Mutat 1999;13:11–28. Nelis E, Timmerman V, De Jonghe P, Van Broeckhoven C, Rautenstrauss B: Molecular genetics and biology of inherited peripheral neuropathies: A fast moving field. Neurogenetics 1999;2:137–148. Suter U, Snipes GJ: Biology and genetics of hereditary motor and sensory neuropathies. Annu Rev Neurosci 1995;18:45–75.


141

5 6 7

8

9 10

11 12 13

14

15

16

17

18

19

20

21 22 23

24

Charcot J-M, Marie P: Sur une forme particulie`re d’atrophie musculaire progressive, souvent familiale, de´butant par les pieds et les jambes et atteignant plus tard les mains. Rev Me´d 1886;6:97–138. Tooth HH: The Peroneal Type of Progressive Muscular Atrophy. London, Lewis, 1886. Dyck PJ, Chance P, Lebo R, Carney JA: Hereditary motor and sensory neuropathies; in Dyck PJ, Thomas PK, Griffin JW, Low PA, Podulso JF (eds): Peripheral Neuropathy. Philadelphia, Saunders, 1993, pp 1094–1136. Thomas PK, Marques W Jr, Davis MB, Sweeney MG, King RH, Bradley JL, Muddle JR, Tyson J, Malcolm S, Harding AE: The phenotypic manifestations of chromosome 17p11.2 duplication. Brain 1997;120:465–478. Dejerine J, Sottas J: Sur la ne´vrite interstitielle, hypertrophique et progressive de l’enfance. C R Soc Biol 1893;45:63–96. Ouvrier RA, McLeod JD, Conchin TE: The hypertrophic forms of hereditary motor and sensory neuropathy. A study of hypertrophic Charcot-Marie-Tooth disease (HMSN type I) and DejerineSottas disease (HMSN type III) in childhood. Brain 1987;110:121–148. Harding AE, Thomas PK: The clinical features of hereditary motor and sensory neuropathy types I and II. Brain 1980;103:259–280. Madrid R, Bradley WG, Davis CJF: The peroneal muscular atrophy syndrome. Part 2. J Neurol Sci 1977;32:91–122. Lupski JR, Montes de Oca-Luna R, Slaugenhaupt S, Pentao L, Guzzetta V, Trask BJ, SaucedoCardenas O, Barker DF, Killian JM, Garcia CA, Chakravarti A, Patel PI: DNA duplication associated with Charcot-Marie-Tooth disease type 1A. Cell 1991;66:219–239. Raeymaekers P, Timmerman V, Nelis E, De Jonghe P, Hoogendijk JE, Baas F, Barker DF, Martin J-J, de Visser M, Bolhuis PA, Van Broeckhoven C, HMSN Collaborative Research Group: Duplication in chromosome 17p11.2 in Charcot-Marie-Tooth neuropathy type 1a (CMT 1a). Neuromuscul Disord 1991;1:93–97. Hayasaka K, Himoro M, Sato W, Takada G, Uyemura K, Shimizu N, Bird T, Conneally PM, Chance PF: Charcot-Marie-Tooth neuropathy type 1B is associated with mutations of the myelin P0 gene. Nat Genet 1993;5:31–34. Warner LE, Mancias P, Butler IJ, McDonald CM, Keppen L, Koob G, Lupski JR: Mutations in the early growth response 2 (EGR2) gene are associated with hereditary myelinopathies. Nat Genet 1998;18:382–384. Chance PF, Bird TD, O’Connell P, Lipe HP, Lalouel J-M, Leppert M: Genetic linkage and heterogeneity in type I Charcot-Marie-Tooth disease (hereditary motor and sensory neuropathy type I). Am J Hum Genet 1990;47:915–925. Vance JM, Barker D, Yamaoka LH, Stajich JM, Loprest L, Hung W-J, Fischbeck K, Roses AD, Pericak-Vance MA: Localization of Charcot-Marie-Tooth disease type 1a (CMT1a) to chromosome 17p11.2. Genomics 1991;9:623–628. Patel PI, Roa BB, Welcher AA, Schoener-Scott R, Trask BJ, Pentao L, Snipes GJ, Garcia CA, Francke U, Shooter EM, Lupski JR, Suter U: The gene for the peripheral myelin protein PMP-22 is a candidate for Charcot-Marie-Tooth disease type 1A. Nat Genet 1992;1:159–165. Timmerman V, Nelis E, Van Hul W, Nieuwenhuijsen BW, Chen KL, Wang S, Ben Othman K, Cullen B, Leach RJ, Hanemann CO, De Jonghe P, Raeymaekers P, van Ommen G-JB, Martin J-J, Mu¨ller HW, Vance JM, Fischbeck KH, Van Broeckhoven C: The peripheral myelin protein gene PMP-22 is contained within the Charcot-Marie-Tooth disease type 1A duplication. Nat Genet 1992;1:171–175. Valentijn LJ, Baas F, Zorn I, Hensels GW, de Visser M, Bolhuis PA: Alternatively sized duplication in Charcot-Marie-Tooth disease type 1A. Hum Mol Genet 1993;2:2143–2146. ¨ zcelik T, Snipes G, Kosaras B, Francke U, Billings-Gagliardi S: Trembler Suter U, Welcher AA, O mouse carries a point mutation in a myelin gene. Nature 1992;356:241–244. Nelis E, Van Broeckhoven C, et al: Estimation of the mutation frequencies in Charcot-Marie-Tooth disease type 1 and hereditary neuropathy with liability to pressure palsies: A European collaborative study. Eur J Hum Genet 1996;4:25–33. Hoogendijk JE, Hensels GW, Gabreels-Festen AAWM, Gabreels FJM, Janssen EAM, De Jonghe P, Martin J-J, Van Broeckhoven C, Valentijn LJ, Baas F, de Visser M, Bolhuis PA: De-novo mutation in hereditary motor and sensory neuropathy type 1. Lancet 1992;339:1081–1082.


142

25

26

27 28 29

30

31

32

33

34

35

36

37

38

39

40

41

42

Roa BB, Garcia CA, Suter U, Kulpa DA, Wise CA, Mueller J, Welcher AA, Snipes GJ, Shooter EM, Patel PI, Lupski JR: Charcot-Marie-Tooth disease type 1A. Association with a spontaneous point mutation in the PMP22 gene. N Engl J Med 1993;329:96–101. Valentijn LJ, Baas F, Wolterman RA, Hoogendijk JE, van den Bosch NHA, Zorn I, Gabree¨lsFesten AAWM, de Visser M, Bolhuis PA: Identical point mutations of PMP-22 in Trembler-J mouse and Charcot-Marie-Tooth disease type 1A. Nat Genet 1992;2:288–291. Roa BB, Dyck PJ, Marks HG, Chance PF, Lupski JR: Dejerine-Sottas syndrome associated with point mutation in the PMP22 gene. Nat Genet 1993;5:269–273. Parman Y, Plante´-Bordeneuve V, Guiochon-Mantel A, Eraksoy M, Said G: Recessive inheritance of a new point mutation of the PMP22 gene in Dejerine-Sottas disease. Ann Neurol 1999;45:518–522. Garcia CA, Malamut RE, England JD, Parry GS, Liu P, Lupski JR: Clinical variability in two pairs of identical twins with the Charcot-Marie-Tooth disease type 1A duplication. Neurology 1995; 45:2090–2093. LeGuern E, Gouider R, Mabin D, Tardieu S, Birouk N, Parent P, Bouche P, Brice A: Patients homozygous for the 17p11.2 duplication in Charcot-Marie-Tooth type 1A disease. Ann Neurol 1997;41:104–108. Garcia CA, Combarros O, Calleja J, Berciano J: Charcot-Marie-Tooth disease type 1A with 17p duplication in infancy and early childhood: A longitudinal clinical and electrophysiologic study. Neurology 1998;50:1061–1067. Kaku DA, Parry GJ, Malamut R, Lupski JR, Garcia CA: Nerve conduction studies in CharcotMarie-Tooth polyneuropathy associated with a segmental duplication of chromosome 17.Neurology 1993;43:1806–1808. Fabrizi GM, Cavallaro T, Taioli F, Orrico D, Morbin M, Simonati A, Rizzuto N: Myelin uncompaction in Charcot-Marie-Tooth neuropathy type 1A with a point mutation of peripheral myelin protein22. Neurology 1999;53:846–851. Warner LE, Hilz MJ, Appel SH, Killian JM, Kolodny EH, Karpati G, Watters GV, Nelis E, Van Broeckhoven C, Lupski JR: Clinical phenotypes of different MPZ (P0) mutations may include CharcotMarie-Tooth 1B, Dejerine-Sottas and congenital hypomyelination. Neuron 1996;17:451–460. Ikegami T, Nicholson G, Ikeda H, Ishida A, Johnston H, Wise G, Ouvrier R, Hayasaka K: A novel homozygous mutation of the myelin Po gene producing Dejerine-Sottas disease (hereditary motor and sensory neuropathy type III). Biochem Biophys Res Commun 1996;222:107–110. Takashima H, Nakagawa M, Kanzaki A, Yawata Y, Horikiri T, Matsuzaki T, Suehara M, Izumo S, Osame M: Germline mosaicism of MPZ gene in Dejerine-Sottas syndrome (HMSN III) associated with hereditary stomatocytosis. Neuromuscul Disord 1999;9:232–238. Gabree¨ls-Festen AAWM, Hoogendijk JE, Meijerink PHS, Gabree¨ls FJM, Bolhuis PA, van Beersum SEC, Kulkens T, Nelis E, Jennekens FGI, de Visser M, van Engelen BGM, Van Broeckhoven C, Mariman ECM: Two divergent types of nerve pathology in patients with different P0 mutations in Charcot-Marie-Tooth disease. Neurology 1996;47:761–765. Marrosu MG, Vaccargiu S, Marrosu G, Vannelli A, Cianchetti C, Muntoni F: Charcot-MarieTooth disease type 2 associated with mutation of the myelin protein zero gene. Neurology 1998; 50:1397–1401. De Jonghe P, Timmerman V, Ceuterick C, Nelis E, De Vriendt E, Lo¨fgren A, Vercruyssen A, Verellen C, Van Maldergem L, Martin J-J, Van Broeckhoven C: The Thr124Met mutation in the peripheral myelin protein zero (MPZ) gene is associated with a clinically distinct Charcot-MarieTooth phenotype. Brain 1999;122:281–290. Chapon F, Latour P, Diraison P, Schaeffer S, Vandenberghe A: Axonal phenotype of CharcotMarie-Tooth disease associated with a mutation in the myelin protein zero gene. J Neurol Neurosurg Psychiatry 1999;66:779–782. Timmerman V, De Jonghe P, Ceuterick C, De Vriendt E, Lo¨fgren A, Nelis E, Warner LE, Lupski JR, Martin J-J, Van Broeckhoven C: Novel missense mutation in the early growth response 2 gene associated with Dejerine-Sottas syndrome phenotype. Neurology 1999;52:1827–1832. Bergoffen J, Scherer SS, Wang S, Oronzi Scott M, Bone LJ, Paul DL, Chen K, Lensch MW, Chance PF, Fischbeck KH: Connexin mutations in X-linked Charcot-Marie-Tooth disease. Science 1993; 262:2039–2042.


143

43

44

45 46 47

48

49

50

51

52

53 54 55 56

57

58

59 60

Meggouh F, Benomar A, Rouger H, Tardieu S, Birouk N, Tassin J, Barhoumi C, Yahyaoui M, Chkili T, Brice A, LeGuern E: The first de novo mutation of the connexin 32 gene associated with X-linked Charcot-Marie-Tooth disease. J Med Genet 1998;35:251–252. Nicholson G, Corbett A: Slowing of central conduction in X-linked Charcot-Marie-Tooth neuropathy shown by brain stem auditory evoked responses. J Neurol Neurosurg Psychiatry 1996;61: 43–46. Nicholson G, Nash J: Intermediate nerve conduction velocities define X-linked Charcot-MarieTooth neuropathy families. Neurology 1993;43:2558–2564. Hahn AF: Hereditary motor and sensory neuropathy: HMSN type II (neuronal type) and X-linked HMSN. Brain Pathol 1993;3:147–155. Ben Othmane K, Hentati F, Lennon F, Ben Hamida C, Blel S, Roses AD, Pericak-Vance MA, Ben Hamida M, Vance JM: Linkage of a locus (CMT4A) for autosomal recessive Charcot-Marie-Tooth disease to chromosome 8q. Hum Mol Genet 1993;2:1625–1628. Bolino A, Brancolini V, Bono F, Bruni A, Gambardella A, Romeo G, Quattrone A, Devoto M: Localization of a gene responsible for autosomal recessive demyelinating neuropathy with focally folded myelin sheaths to chromosome 11q23 by homozygosity mapping and haplotype sharing. Hum Mol Genet 1996;5:1051–1054. Gambardella A, Bolino A, Muglia M, Bono F, Valentino P, Oliveri RL, Sabatelli M, Brancolini C, Van Broeckhoven C, Romeo G, Devoto M, Quattrone A: Genetic heterogeneity in autosomal recessive hereditary motor and sensory neuropathy with focally folded myelin sheats (CMT4B). Neurology 1998;50:799–801. LeGuern E, Guibot A, Kessali M, Ravise´ N, Tassin J, Maisonobe T, Grid D, Brice A: Homozygosity mapping of an autosomal recessive form of demyelinating Charcot-Marie-Tooth disease to chromosome 5q23–q33. Hum Mol Genet 1996;5:1685–1688. Kessali M, Zemmouri R, Guilbot A, Maisonobe T, Brice A, LeGuern E, Grid D: A clinical, electrophysiologic, neuropathologic, and genetic study of two large Algerian families with an autosomal recessive demyelinating form of Charcot-Marie-Tooth disease. Neurology 1997;48:867– 873. Kalaydjieva L, Hallmayer J, Chandler D, Savov A, Nikolova A, Angelicheva D, King RHH, Ishpekova B, Honeyman K, Calafell F, Shmarov A, Petrova J, Turnev I, Hristova A, Moskov M, Stancheva S, Petkova I, Bittles AH, Geogieva V, Middleton L, Thomas PK: Gene mapping in Gypsies identifies a novel demyelinating neuropathy on chromosome 8q24. Nat Genet 1996;14: 214–217. Ionasescu VV, Kimura J, Searby CC, Smith WL Jr, Ross MA, Ionasescu R: A Dejerine-Sottas neuropathy family with a gene mapped on chromosome 8. Muscle Nerve 1996;19:319–323. Lynch DR, Hara H, Yum SW, Chance PF, Scherer SS, Bird SJ, Fischbeck KH: Autosomal dominant transmission of Dejerine-Sottas disease (HMSN III). Neurology 1997;49:601–603. Priest JM, Fischbeck KH, Nouri N, Keats BJB: A locus for axonal motor-sensory neuropathy with deafness and mental retardation maps to Xq24-q26. Genomics 1995;29:409–412. Ben Othmane K, Middleton LT, Loprest LJ, Wilkinson KM, Lennon F, Rozear MP, Stajich JM, Gaskell PC, Rosed AD, Pericak-Vance MA, Vance JM: Localization of a gene (CMT2A) for autosomal dominant Charcot-Marie-Tooth disease type 2 to chromosome 1p and evidence of genetic heterogeneity. Genomics 1993;17:370–375. Kwon JM, Elliott JL, Yee WC, Ivanovich J, Scavarda NJ, Moolsintong PJ, Goodfellow PJ: Assignment of a second Charcot-Marie-Tooth type II locus to chromosome 3q. Am J Hum Genet 1995; 57:853–858. Ionasescu VV, Searby C, Sheffield VC, Roklina T, Nishimura D, Ionasescu R: Autosomal dominant Charcot-Marie-Tooth axonal neuropathy mapped on chromosome 7p (CMT2D). Hum Mol Genet 1996;5:1373–1375. Yoshioka R, Dyck PJ, Chance PF: Genetic heterogeneity in Charcot-Marie-Tooth neuropathy type 2. Neurology 1996;46:569–571. Timmerman V, De Jonghe P, Spoelders P, Simokovic S, Lo¨fgren A, Nelis E, Vance J, Martin J-J, Van Broeckhoven C: Linkage and mutation analysis of Charcot-Marie-Tooth neuropathy type 2 families with chromosomes 1p35-p36 and Xq13. Neurology 1996;46:1311–1318.


144

61

62

63 64 65 66 67

68

69

70

71

72

73

74

75

76 77 78

79

De Jonghe P, Timmerman V, FitzPatrick D, Spoelders P, Martin J-J, Van Broeckhoven C: Mutilating neuropathic ulcerations in a chromosome 3q13-q22 linked Charcot-Marie-Tooth disease type 2B family. J Neurol Neurosurg Psychiatry 1997;62:570–573. Bouhouche A, Benomar A, Birouk N, Mularoni A, Meggouh F, Tassin J, Grid D, Vandenberghe A, Yahyaoui M, Chkili T, Brice A, LeGuern E: A locus for an axonal form of autosomal recessive Charcot-Marie-Tooth disease maps to chromosome 1q21.2-q21.3. Am J Hum Genet 1999;65:722– 727. Emery AEH: Review: The nosology of the spinal muscular atrophies. J Med Genet 1971;8:481–495. Boylan KB, Cornblath DR, Glass JD, Alderson K, Kuncl RW, Kleyn PW, Gilliam TC: Autosomal dominant distal spinal muscular atrophy in four generations. Neurology 1995;45:699–704. Fleury P, Hageman G: A dominantly inherited lower motor neuron disorder presenting at birth with associated arthrogryposis. J Neurol Neurosurg Psychiatry 1985;48:1037–1048. Frijns CJ, Van Deutekom J, Frants RR, Jennekens FG: Dominant congenital benign spinal muscular atrophy. Muscle Nerve 1994;17:192–197. Timmerman V, Raeymaekers P, Nelis E, De Jonghe P, Muylle L, Ceuterick C, Martin J-J, Van Broeckhoven C: Linkage of distal hereditary motor neuropathy type II (distal HMN II) in a single pedigree. J Neurol Sci 1992;109:41–48. Timmerman V, De Jonghe P, Simokovic S, Lo¨fgren A, Beuten J, Nelis E, Ceuterick C, Martin J-J, Van Broeckhoven C: Distal hereditary motor neuropathy type II (distal HMN II): Mapping of a locus to chromosome 12q24. Hum Mol Genet 1996;5:1065–1069. Christodoulou K, Kyriakides T, Hristova AH, Georgiou DM, Kalaydjieva L, Yshpekova B, Ivanova T, Weber JL, Middleton LT: Mapping of a distal form of spinal muscular atrophy with upper limb predominance to chromosome 7p. Hum Mol Genet 1995;4:1629–1632. De Jonghe P, Timmerman V, Van Broeckhoven C, and workshop participants: 2nd workshop of the European CMT consortium: 53rd ENMC International Workshop on Classification and Diagnostic Guidelines for Charcot-Marie-Tooth Type 2 (CMT2-HMSN II) and Distal Hereditary Motor Neuropathy (distal HMN – spinal CMT) 26–28 September 1997, Naarden, The Netherlands. Neuromuscul Disord 1998;8:426–431. van der Vleuten AJW, van Ravenswaaij-Arts CMA, Frijns CJM, Smits APT, Hageman G, Padberg GW, Kremer H: Localisation of the gene for a dominant congenital spinal muscular atrophy predominantly affecting the lower limbs to chromosome 12q23-q24. Eur J Hum Genet 1998;6: 376–382. Middleton LT, Christodoulou K, Mubaidin A, Zamba E, Tsingis M, Kyriacou K, Abu-Sheikh S, Kyriakides T, Neocleous V, Georgiou DM, El-Khateeb M, Al-Qudah A, Horany K: Distal hereditary motor neuronopathy of the Jerash type. Ann NY Acad Sci 1999;883:65–68. Dyck PJ: Neuronal atrophy and degeneration predominantly affecting peripheral sensory and autonomic neurons; in Dyck PJ, Thomas PK, Griffin JW, Low PA, Podulso JF (eds): Peripheral Neuropathy. Philadelphia, Saunders, 1993, pp 1065–1093. Nicholson GA, Dawkins JL, Blair IP, Kennerson ML, Gordon MJ, Cherryson AK, Nash A, Bananis T: The gene for hereditary sensory neuropathy type I (HSN-I) maps to chromosome 9q22.1-q22.3. Nat Genet 1996;13:101–104. Auer-Grumbach M, Wagner K, Timmerman V, De Jonghe P, Hartung HP: Ulcero-mutilating neuropathy in an Austrian kinship without linkage to HMSN IIB and HSN I loci. Neurology 2000; 54:45–52. DeJong JGY: Over families met hereditaire dispositie tot het optreden van neuritiden, gecorreleerd met migraine. Psychiat Neurol Bl Amst 1947;50:60–76. Windebank AJ: Inherited recurrent focal neuropathies; in Dyck PJ, Thomas PK, Griffin JW, Low PA, Podulso JF (eds): Peripheral Neuropathy. Philadelphia, Saunders, 1993, pp 1137–1148. Chance PF, Alderson MK, Leppig KA, Lensch MW, Matsunami N, Smith B, Swanson PD, Odelberg SJ, Distsche CM, Bird TD: DNA deletion associated with hereditary neuropathy with liability to pressure palsies. Cell 1993;72:143–151. Mariman ECM, Gabree¨ls-Festen AAWM, van Beersum SEC, Jongen PJH, van de Looij E, Baas F, Bolhuis PA, Ropers HH, Gabree¨ls FJM: Evidence for genetic heterogeneity underlying hereditary neuropathy with liability to pressure palsies. Hum Genet 1994;93:151–156.


145

80

81

82

83

84

85

Gouider R, LeGuern E, Gugenheim M, Tardieu S, Maisonobe T, Le´ger JM, Vallat J-M, Agid Y, Bouche P, Brice A: Clinical, electrophysiologic, and molecular correlations in 13 families with hereditary neuropathy with liability to pressure palsies and chromosome 17p11.2 deletion. Neurology 1995;45:2018–2023. Lenssen PPA, Gabre{e:}ls-Festen AA, Valentijn LJ, Jongen PJH, van Beersum SEC, van Engelen BGM, van Wensen PJM, Bolhuis PA, Gabree¨ls FJ, Mariman ECM: Hereditary neuropathy with liability to pressure palsies: Phenotypic differences between patients with the common deletion and a PMP22 frame shift mutation. Brain 1998;121:1451–1458. Pellegrino JE, Rebbeck TR, Brown MJ, Bird TD, Chance PF: Mapping of hereditary neuralgic amyotrophy (familial brachial plexus neuropathy) to distal chromosome 17q. Neurology 1996;46: 1128–1132. Meuleman J, Kuhlenbaümer G, Schirmacher A, Wehnert A, De Jonghe P, De Vriendt E, Young P, Airaksinen E, Pou-Serradell A, Prats J-M, Ringelstein EB, Sto¨gbauer F, Van Broeckhoven C, Timmerman V: Genetic refinement of the hereditary neuralgic amyotrophy (HNA) locus at chromosome 17q25. Eur J Hum Genet 1999;7:920–927. Navon R, Timmerman V, Lo¨fgren A, Liang P, Nelis E, Zeitune M, Van Broeckhoven C: Prenatal diagnosis of Charcot-Marie-Tooth disease type 1A (CMT1A) using molecular genetic techniques. Prenat Diagn 1995;15:633–640. De Vos A, Sermon K, Van de Velde H, Joris H, Vandervorst M, Lissens W, Mortier G, De Sutter P, Lo¨fgren A, Van Broeckhoven C, Liebaers I, Van Steirteghem A: Pregnancy after preimplantation genetic diagnosis for Charcot-Marie-Tooth disease type 1A. Mol Hum Reprod 1998;4:978–984.

Peter De Jonghe, Laboratory of Molecular Genetics, Department of Biochemistry, University of Antwerp (UIA), Universiteitsplein 1, B–2610 Antwerp (Belgium) E-Mail [email protected]


146


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

Acute Inflammatory Demyelinating Polyradiculoneuropathy Isabelita R. Bella Department of Neurology, UMass Memorial Health Care, Worcester, Mass., USA

Acute inflammatory demyelinating polyradiculoneuropathy (AIDP) is an immunologically mediated disorder producing multifocal demyelination of peripheral nerves, nerve roots and cranial nerves. In recent years, neurologists have gained a better understanding of the pathophysiology of this disorder that has lead to advances in therapeutic modalities. The eponym, Guillain-Barre´ syndrome (GBS), has been used synonymously with AIDP, in recognition of the authors who provided descriptions of this condition. In recent years, the term GBS has come to encompass a number of disorders of different pathophysiological patterns comprised of demyelinating and axonal variants. AIDP is the most common subtype in developed countries, while axonal variants – acute motor axonal neuropathy (AMAN) and acute motor sensory axonal neuropathy (AMSAN) – are found more commonly in northern China [1]. AIDP is the most common cause of rapidly progressive weakness with an annual incidence of 0.6–1.9 cases per 100,000 population [2]. It occurs at all ages [2]. The potential to cause respiratory failure and autonomic nervous system instability make this disorder a potential neurologic emergency.

Clinical Features The major clinical features of GBS are progressive weakness and areflexia. Weakness evolves rapidly (usually over days) and is heralded by paresthesias of the feet and hands. The legs are often affected first (approximately 50% of cases) with subsequent spread to the arms. Weakness may, however, start in the cranial nerves or arms and descend to the legs (approximately 14% of

cases), or start simultaneously in the arms and legs (approximately one third of cases) [2]. Unlike typical axonal neuropathies, proximal muscles of the arms and legs are affected early in the course of the disease. Paresthesias described as ‘pins and needles’ and ‘tingling’ sensations often occur in the distal musculature and ascend as the disease progresses. Moderate to severe deep aching pain in the back and leg simulating sciatica and/or dysesthesias of the limbs was seen in approximately 50% of patients at presentation in one series [3]. Pain may be exacerbated by straight leg raising, suggesting that the pain may be due to traction of an inflamed nerve root [3]. On examination, there is symmetric proximal and distal muscle weakness and loss or attenuation of reflexes. Objective signs of sensory loss are usually mild, often involving only diminished vibratory sensation despite complaints of significant pain and dysesthesias. Widespread and profound sensory loss however has been described as a variant of GBS [4]; presumably the brunt of the pathology falls on sensory nerves. Respiratory muscle involvement may occur, with up to one third of patients requiring mechanical ventilation [2] within 18 days after onset of symptoms [5]. Bilateral facial and oropharyngeal weakness are seen in more than half the patients with AIDP [6] while ophthalmoparesis occurs in less than 20% [2]. Ophthalmoplegia is rare but may be seen in association with ataxia and areflexia in the Miller-Fisher variant [2]. Pupillary abnormalities have been described primarily in those with severe ophthalmoparesis and ptosis, presumably secondary to involvement of postganglionic parasympathetic and sympathetic nerves. Rarely, papilledema with pseudotumor cerebri occurs in the setting of markedly elevated cerebrospinal fluid (CSF) protein (?200 mg/dl) [2]. Autonomic nervous system abnormalities are potentially lethal and may be seen in as many as 65% of patients [2]. They are more frequent in patients with severe motor dysfunction and respiratory failure. Manifestations include cardiac arrhythmias (sinus tachycardia, bradycardia, ventricular tachycardia, atrial flutter, atrial fibrillation, and asystole), orthostatic hypotension, and hypertension [2, 7]. Transient bladder paralysis, sweating abnormalities, and paralytic ileus may also occur [2]. Weakness typically does not progress for longer than 4 weeks; 50% of patients will reach the nadir of their clinical course in 2 weeks, and 90% in 4 weeks [4]. Weakness progressing longer than 4 weeks may be seen in a variant of GBS [4] but progression of a demyelinating polyneuropathy for more than 2 months suggests chronic inflammatory demyelinating polyradiculoneuropathy (CIDP) [8–10]. CIDP shares similar clinical, electrophysiological, and pathological features to AIDP but differs in its temporal evolution, course, and response to treatment [11]. Two to five percent of patients have recurrent GBS [12].

Bella

148

The severity of GBS is variable. Symptoms may vary from mild face and limb weakness to quadriplegia and requirement for mechanical ventilation within a few days of onset. Variations to the typical clinical picture of GBS include pure motor GBS, pure sensory GBS, Miller-Fisher variant, pandysautonomia, and axonal forms (see below).

Axonal Variants Until recently, GBS has been used interchangeably with AIDP, referring to its predominant pathological features of demyelination and inflammation. Axonal degeneration, when encountered, was felt to be secondary to intense inflammation – a ‘bystander’ effect [13, 14]. The recognition of primary axonal forms of GBS, AMSAN and AMAN, has broadened the clinical spectrum of GBS. In 1986, Feasby et al. [14] reported 5 patients with severe motor and sensory neuropathy that fulfilled the clinical criteria for GBS. These patients differed clinically in that the time to peak severity was much shorter (1 week) and symptoms were more severe with more than half requiring mechanical ventilation; all had inexcitable motor nerves, and all but 1 had poor recovery. Autopsy of 1 patient revealed an acute axonal neuropathy without evidence of demyelination, suggesting that the primary insult was to the axon. Griffin et al. [1] described similar cases of GBS in northern China and introduced the term AMSAN to describe these patients. They also discovered another distinct pattern of GBS prominent in northern China – that of an acute AMAN occurring primarily in the summer months among rural children and young adults [15]. Autopsy studies showed wallerian-like degeneration of motor fibers. Recovery was good, similar to that of traditional AIDP, suggesting that degeneration primarily affects the motor nerve terminals and intramuscular axons [16].

Antecedent Events In approximately two thirds of patients, there is an acute flu-like illness 1–3 weeks prior to the onset of the neuropathy, most commonly with upper respiratory symptoms, or a gastroenteritis. A number of infectious agents have been associated with GBS [2, 17] including cytomegalovirus, Epstein-Barr virus, hepatitis, influenza, mycoplasma, herpes virus, and Campylobacter jejuni. C. jejuni, in particular, has been been found to be an important antecedent infectious agent. Culture or serological evidence of C. jejuni occurs in 26–40% of sporadic GBS in developed countries [18, 19]. In contrast, 66–74% of GBS

Acute Inflammatory Demyelinating Polyradiculoneuropathy

149

patients in China, where the motor variant is more common, are seropositive for C. jejuni [20, 21]. Although C. jejuni is found in both axonal and demyelinating forms of GBS in addition to the Fisher variant, some studies show it occurs more frequently in the motor axonal form (AMAN) [19, 22]. There are conflicting reports as to whether C. jejuni infection correlates with a more severe neuropathy with a poor prognosis [18, 19]. Because the convalescent excretion of C. jejuni lasts less than 3 weeks, serology may be more sensitive than stool culture in identifying infection in a patient who later develops GBS [22]. Visser et al. [23] suggest that GBS related to CMV infection produces a different clinical pattern compared to GBS patients with C. jejuni infection or GBS patients without either infection. They found CMV-associated GBS patients were younger, developed severe sensory loss, more often developed cranial nerve involvement (most commonly facial paresis), and more often required mechanical ventilation [23]. Other antecedent events include immunization, renal transplantation and surgery. GBS may also occur in the setting of an underlying systemic disease such as systemic lupus erythematosus, Hodgkin’s disease, and in early HIV infection. Rarely, Lyme polyradiculoneuropathy may mimic GBS. Epidemiological studies determined that there was an increased risk of developing GBS up to 6 weeks after immunization with the 1976–1977 swine flu vaccine [24]. Recent influenza vaccination programs (1992–1993 and 1993– 1994) were associated with only a small risk of GBS (1–2 additional cases per 1 million vaccinated persons) [25]. Many physicians are understandably concerned about administering influenza vaccinations to GBS patients. It appears reasonable to follow the recommendation of Hughes et al. [26] of delaying immunization for 1 year after the onset of GBS, to prevent the small chance of relapse of symptoms with vaccination [26]. Patients should avoid reimmunization if the initial vaccination precipitated GBS, especially with tetanus toxoid [26].

Laboratory Features Albuminocytologic dissociation (elevated CSF protein without pleocytosis), electrodiagnostic studies revealing evidence of demyelination, and nerve biopsy revealing inflammation and demyelination are the characteristic laboratory findings in GBS. Routine Laboratory Studies Hematologic studies are typically normal but may show a mild increase in erythrocyte sedimentation and white blood cell count [2]. Creatine kinase

Bella

150

levels are often mildly elevated in those with severe pain, perhaps reflecting myonecrosis [2]. Hyponatremia may be seen on occasion due to inappropriate secretion of antidiuretic hormone possibly due to a resetting of the osmoreceptor response [17]. Cerebrospinal Fluid Albuminocytologic dissociation in the CSF is characteristic of GBS. A breakdown in the blood-CSF barrier may account for these abnormalities [2]. In the first 48 h after symptom onset, CSF protein may be normal, but after the 1st week of symptoms, CSF protein is elevated [4], peaks in 3–4 weeks, at times reaching as high as 1,800 mg/dl [2]. Rarely, no rise in CSF protein is seen even after several weeks. The presence of oligoclonal bands has also been described in GBS [2]. The CSF is typically acellular with 10 or fewer mononuclear leukocytes/mm3; however, 11–50 mononuclear leukocytes/mm3 may rarely be encountered [4]. CSF pleocytosis raises the possibility that GBS may be occurring in the setting of HIV or Lyme disease. Autoantibodies to Glycoconjugates A number of antiglycoconjugate antibodies have been described in GBS. The strongest association is between IgG anti-GQ1b and the Miller-Fisher syndrome [21]. Approximately 90% of Miller-Fisher syndrome patients have high titers of GQ1b antibodies in the acute phase which disappear with clinical recovery [27]. The association between other ganglioside antibodies and GBS, however, is variable. Ganglioside antibodies which have been described in GBS include LM1, GM1, GD1b, GD1a and GT1b [28]. Anti-GM1 antibodies have been reported to occur in 25–60% of GBS patients [18, 21]. The significance of these antibodies is unclear. Several studies suggest anti-GM1 antibodies occur more frequently in the pure motor form of GBS [21, 29], in those with axonal variants [29, 30], and in those with a poor prognosis [29], while others have found no correlation between the presence of anti-GM1 antibodies and outcome or pattern of disease (axonal vs. demyelinating) [18, 19]. Kuwabara et al. [30] shed light into the prognostic value of anti-GM1 antibodies. They found the presence of anti-GM1 antibodies to correlate with the acute motor axonal pattern of GBS. However, there were two patterns of clinical recovery: those who improved quickly over 2 weeks and those who had a slower recovery than the anti-GM1-negative patients, requiring several months for recovery and for whom prognosis was poor. The rapid recovery of the first group of patients suggests that a mechanism other than axonal degeneration must be in place. Perhaps reversible immune-mediated changes at the nodes of Ranvier in motor fibers [31] produce conduction block and


151

subsequent weakness. Anti-GM1 antibodies have been reported to affect sodium and potassium currents [32] and to block conduction, lending support to this mechanism [31]. Degeneration and regeneration of intramuscular motor nerve terminals may be another mechanism that could account for rapid recovery in AMAN patients [31]. Poor outcomes in the second group of patients suggest an axonal pathogenesis. More recently, IgG anti-GD1a antibodies have been strongly associated with the AMAN subtype of GBS found in China [21]. Electrodiagnostic Studies In AIDP. Electrodiagnostic studies in AIDP patients reveal evidence of a demyelinating neuropathy. Conduction velocity is reduced (080% of lower limit of normal) and sensory and motor distal latencies are prolonged [4]. Characteristic findings of segmental demyelination such as partial conduction block and temporal dispersion due to differential slowing help differentiate acquired from hereditary demyelinating disorders [33]. Similar to CSF protein abnormalities, routine electrical studies may be normal early in the disease; in these cases, F wave responses are often prolonged [33] presumably due to proximal nerve or nerve root involvement. The compound motor action potential amplitude may be reduced due to axon loss or distal conduction block. Needle electromyography may only show reduced recruitment early in course of GBS, but after several weeks, fibrillation potentials and positive sharp waves may be seen if there has been axon loss. In Axonal Variants. In the severe axonal form of GBS, AMSAN, motor and sensory nerves may be electrically inexcitable [14]. In AMAN patients, electrodiagnostic studies disclose low or absent motor responses with normal conduction velocities and normal sensory responses [1]. Needle examination in both axonal variants may reveal denervation potentials [34], within 3–4 weeks of onset of symptoms [35].

Pathogenesis It now appears that the varied histopathological patterns seen in GBS may stem from different pathogenetic mechanisms. All are immune-mediated but the immune response may be targeting different portions of the peripheral nerve to account for the various presentations of GBS. In AIDP the immune attack appears to be directed to epitopes on the Schwann cell while in axonal variants, the axon is preferentially targeted. The exact antigenic determinants to which the immune response is directed, however, are not known [2]. Gangliosides or glycoconjugates, located at the abaxonal surface of the Schwann

Bella

152

cell in AIDP or at the motor nodes of Ranvier and internodal axolemma in AMAN, have been proposed as the targets of the immune attack [36, 37]. In AIDP Both cellular and humoral components of the immune system appear to play a role in the demyelinating lesion of GBS. Observations in the animal model of GBS called experimental autoimmune neuritis (EAN) lend support to a cellular-mediated immune response in the pathogenesis of GBS [38]. Waksman and Adams [39] injected rabbit nerve/ganglia and Freund’s adjuvant into rabbits which produced quadriplegia within 2 weeks. Transfer of specific T cell lines reactive to P2/PO produced the classical clinical, electrophysiological, and pathological features of EAN in rats [38]. Further evidence of T cell activation comes from the finding of elevated levels of soluble IL-2 receptors in GBS [40]. Equally important findings support a humoral mediated immune response in the pathogenesis of GBS. Koski et al. [41] found anti-peripheralnerve myelin antibodies in the sera of GBS patients with clinical improvement correlating with a reduction of the anti-peripheral-nerve myelin titer. Sera from EAN animals and GBS patients injected intraneurally or subperineurally in animals have also produced demyelination [42, 43]. Lastly, the effectiveness of plasmapheresis in GBS supports a major role for humoral factors [44]. Recent evidence suggest that the target of the immune attack in AIDP is the Schwann cell. Hafer-Macko et al. [36] found complement activation markers (C3d and C5b-9) along the outer surface of the Schwann cell, supporting a complement-mediated immune mechanism directed towards the Schwann cell [36]. They speculate that complement-fixing antibodies bind to epitopes on the outer surface of the Schwann cell, thereby activating complement and leading to myelin vesiculation. In Axonal Forms Evidence that axonal constituents are the target of the immune attack in axonal forms of GBS comes from work by Hafer-Macko et al. [37]. They examined peripheral nerves from 7 AMAN patients who came to autopsy. They found IgG and C3d (a complement activation product) bound to the nodal and internodal axolemma in motor fibers which had not yet begun wallerian-like degeneration [37]. Macrophages were also found in internodal periaxonal space displacing the axon from the Schwann cell and myelin sheath. This is in contrast to AIDP, in which complement activation markers were found on the Schwann cell itself. The above findings suggest that axonal forms of GBS may be complement- and IgG-mediated, and that the immune attack is directed to epitopes of the axolemma. The specific epitope again is not known, but the presence of high titers of anti-GD1a antibodies in AMAN


153

but not AIDP patients suggests that a GD1a-like epitope may be the target of the immune attack [21]. As mentioned earlier, the rapid improvement seen in AMAN patients may be due to reversible changes at the node. In their study using rat nerve fibers, Takigawa et al. [32] found that anti-GM1 antibodies, in the presence of complement, decreases Na+ current and induces membrane leakage. This may lead to blocking of Na+ channels and prevent further impulse conduction. Molecular Mimicry The presence of antiganglioside antibodies, particularly anti-GM1 antibodies, in both demyelinating and axonal forms of GBS and the finding of ganglioside-like epitopes on some strains of C. jejuni have led to the concept of ‘molecular mimicry’ [21, 45], in which an immune attack occurs on the epitope shared by the nerve fiber and infectious organism [46], as a possible mechanism for C. jejuni-associated GBS. Yuki et al. [45] demonstrated the presence of a terminal tetrasaccharide located in lipopolysaccharides of C. jejuni serotype (PEN 19) to be identical to that of GM1 [45]. The authors speculate that infection by C. jejuni induces production of antiGM1 antibodies which may bind to motor nerve terminals causing nerve inexcitability and subsequent weakness [45]. Anti-GQ1b-like epitopes have also been demonstrated in C. jejuni isolates associated with the Fisher syndrome [47]. Infection with a C. jejuni strain containing the same ganglioside-like epitopes shared with nerve fibers and the presence of sera containing antiganglioside antibodies against these specific ganglioside-like epitopes are not in itself sufficient to cause GBS [46]. Perhaps differences in host susceptibility (genetic predisposition, intercurrent illness) or differences in organisms may account for the varied immune responses [46].

Pathology The predominant pathological features of AIDP are a mononuclear inflammatory infiltrate of the endoneurium – often in a perivenular distribution, and segmental demyelination [13]. The inflammatory changes are present throughout the peripheral nervous system, affecting sensory and motor nerve roots, spinal nerves, plexuses, proximal and distal peripheral nerve trunks, and even intramuscular nerve twigs [13]. Areas of segmental demyelination – characterized by retraction of the myelin sheath in the node of Ranvier in early lesions and denuded axons in more developed lesions – are found adjacent to inflammatory areas [13]. Nerve biopsy, however, is rarely indicated as the

Bella

154

clinical, electrodiagnostic, and CSF abnormalities of AIDP reliably support the diagnosis.

Differential Diagnosis Table 1 contains a number of conditions which cause rapidly progressive weakness and must be differentiated from AIDP.

Management and Treatment GBS patients should be observed in the hospital, preferably in an intensive care unit, for potential respiratory and autonomic nervous system compromise (see table 2). Patients with neck flexor weakness (correlates with diaphragmatic muscle weakness), in particular, require close monitoring. Forced vital capacity and maximum inspiratory pressure should be monitored around the clock as weakness may rapidly progress to involve respiratory muscles. Obtaining an arterial blood gas is helpful in determining adequate oxygenation and possible hypercapnia. Clinically, however, patients may become restless, tachycardic, tachypneic, and sedated before changes in blood gases are detected. A normal forced vital capacity is 65 ml/kg; at a level of 30 ml/kg, a poor cough with accumulation of secretions may be seen which can be managed with chest physical therapy and supplemental oxygen. At a level of 25 ml/kg, the sigh mechanism is compromised producing atelectasis and hypoxia; incentive spirometry and deep breathing can minimize atelectasis [48]. Criteria for intubation have been suggested by Ropper and Kehne [49] (see table 1). Tracheostomy should be delayed for 7–10 days as up to one-third of patients may improve rapidly and can be extubated after the first few days [49]. Signs of autonomic nervous system dysfunction should be monitored by measuring blood pressure, fluid status, and cardiac rhythm. Hypotension can be managed by infusion of fluids; hypertension treated only if persistent and severe, using short-acting a-adrenergic blocking agents, and bradyarrhythmias with atropine [7]. Care should also be taken to avoid complications of a bedridden patient, such as deep venous thrombosis and pulmonary embolism, compression neuropathies, and pressure sores. Frequent turning of the patient, physical therapy, insulating pads over potential compression sites, and subcutaneous heparin can help prevent these complications. Patients with GBS may be treated with either plasmapheresis (PE) or intravenous immunoglobulin (IVIG). Several large, multicenter controlled


155

Table 1. Conditions that may mimic GBS [adapted from 64, with permission] Disorder

Major distinguishing features

Myasthenia gravis

Reflexes are spared Ocular weakness predominates Positive response to edrophonium EMG: decremental motor response Botulism Predominant bulbar involvement Autonomic abnormalities (pupils) EMG: normal velocities, low amplitudes, incremental response (with HFRNS) Tick paralysis Rapid progression (l–2 days) Tick present Shellfish poisoning Rapid onset (face, finger, toe numbness) Follows consumption of mussels/clams Toxic neuropathies EMG: usually axon loss Organophosphorus toxicity Acute cholinergic reaction Porphyric neuropathy Mental disturbance Abdominal pain Diphtheritic neuropathy Prior pharyngitis Slower evolution Palatal/accommodation paralysis Myocarditis Poliomyelitis Weakness, pain and tenderness Preserved sensation CSF: protein and cell count elevated Periodic paralysis Reflexes normal Cranial nerves and respiration spared Abnormal serum [K+] Critical illness neuropathy Sepsis and multiorgan failure ?2 weeks EMG: axon loss Acute myopathy of Tetraparesis and areflexia intensive care Follows prolonged treatment with NMBA and corticosteroids Trauma; status asthmaticus; and organ transplantationassociated Clinical and EMG features of myopathy Vasculitic neuropathy Systemic signs Asymmetry in sensory and motor loss Reflexes reduced proportionate to weakness Rabies Fever Asymmetric sensory and motor signs Muscle spasms, agitation, abnormal mental status EMG>Electromyogram; HFRNS>high-frequency repetitive nerve stimulation; NMBA> neuromuscular blocking agent.

Bella

156

Table 2. Management of GBS [adapted from 65, with permission] Clinical status

Management

Very mild GBS Ambulatory without assistance and no ventilatory compromise

Admit to ward Monitor VC every 8 h Observation Consider PE or IVIG1, 2

Ambulatory only with assistance and no ventilatory compromise

Admit to ICU if possible or to floor with frequent monitoring of VC Check ABG IVIG1 or PE (4 exchanges)

Not ambulatory and mild ventilatory compromise

Admit to ICU Monitory VC frequently Check ABG Intubate if [49] VC =12–15 ml/kg Falling VC over 4–6 h Oropharyngeal paresis with aspiration Respiratory fatigue with VC of 15 ml/kg IVIG1 or PE (4 exchanges)

Not ambulatory and requires assisted ventilation

Admit to ICU Monitor for autonomic nervous system dysfunction [7] Treat hypotension with infusion of fluids Use short-acting a-adrenergic blocking agents to treat persistent hypertension Treat bradyarrhythmias with atropine Frequent turns to avoid pressure sores Avoid compression neuropathies Physical therapy IVIG1 or PE (4 exchanges)

ABG>Arterial blood gases. 1 IVIG is preferable if there is poor vascular access or cardiovascular instability. The dose administered is 400 mg/kg/day for 5 consecutive days. 2 One study [53] demonstrated improvement with 2 plasma exchanges compared to none in mild GBS patients. There has been no controlled study showing benefit of IVIG in mild GBS patients. Because efficacy of IVIG and plasma exchange has been demonstrated in patients with more severe GBS, IVIG would appear to be just as helpful for mild GBS.

trials [50, 51] demonstrated the beneficial effect of PE when used within the first 2 weeks of disease onset, even in those with poor prognostic signs [52]. On average, patients treated with PE were able to walk 1 month earlier than untreated patients; respiratory-dependent patients treated with PE walked 3 months earlier than those untreated [50]. The GBS study group recommends


157

exchanging 200–250 ml/kg in 3–5 sessions over 7–14 days [50]. PE, until recently, was utilized in patients who were not ambulatory, had evidence of respiratory compromise, or had bulbar difficulties. Results from the French Cooperative Group on Plasma Exchange in Guillain-Barre´ Syndrome study revealed that mildly affected patients who remain ambulatory also benefit from 2 plasma exchanges while moderate to severely affected patients (those not ambulatory or requiring mechanical ventilation) benefit from 4 plasma exchanges; severely affected patients did not benefit from 2 additional exchanges [53]. Spontaneous relapse may occur in a small number of patients (3–10%) within days to weeks after improvement; additional treatment with plasma exchange benefits these patients [54]. PE requires good venous access and is available in major medical centers. Patients who have cardiovascular instability or autonomic dysfunction may tolerate this poorly due to its potential for inducing hypotension. These limitations prompted a search for alternative treatments. The favorable results of several uncontrolled studies suggested IVIG may be a therapeutic alternative to PE. A large randomized trial was subsequently performed by Dutch investigators [55] and found that IVIG given in the first 2 weeks of the disease was as efficacious as PE. Following this study, two small uncontrolled studies [56, 57] reported a high relapse rate of more than 50% compared to 10% seen in PE-treated patients [54]. Subsequently, a large randomized controlled trial conducted by The Plasma Exchange/Sandoglobulin GuillainBarre´ Syndrome Trial Group confirmed the equivalence of IVIG and PE; there was not a substantial benefit in using a combination of PE followed by IVIG although there was a nonsignificant trend favoring the use of combination treatment in some outcome measures [58]. Minor side effects of IVIG including flulike symptoms, headache, nausea and fatigue occur in less than 10% of patients [59]. Care must be taken to avoid administering IVIG to those with IgA deficiency or renal disease as it may cause anaphylaxis in the former [59] and renal failure in the latter [60]. Because of its relative ease of administration, IVIG has increasingly become the initial treatment in AIDP. The use of corticosteroids in AIDP is generally ineffective. The results of a pilot study suggest that treatment with IVIG and methylprednisolone may be more advantageous than treatment with IVIG alone [61]. Larger, controlled trials are needed to confirm this.

Outcome The majority of patients have a good outcome. Recovery typically occurs over weeks to months and patients are usually able to return to normal life

Bella

158

within 6–12 months [2]. The length of time to full recovery is related to the severity of the illness. It may take 1.5–2 years of intense rehabilitation to attain maximal improvement [2]. Several factors are associated with a poor outcome: age greater than 60 years, rapidly progressive weakness in less than 7 days, need for mechanical ventilation, and the most powerful predictor – a mean distal motor amplitude of 20% of normal or less [52]. The presence of all four factors is associated with a less than 20% probability of walking unassisted at 6 months without treatment. Treatment with plasmapheresis, however, increases this probability to 40% [52]. Approximately 75% of patients make a good recovery; of these, less than 15% have no residual deficits and the remainder are able to return to work with mild residual deficits manifesting as paresthesias or minimal distal weakness [2]. Severe residual deficits impairing ambulation occur in approximately 10% of patients despite therapy [62]. Although mortality has improved with the institution of mechanical ventilation and respiratory care, 2–12% of patients with AIDP die from the disorder. Causes include sepsis, adult respiratory distress syndrome, autonomic dysfunction, pulmonary embolism, and most commonly, ventilator-associated pneumonia [63].

Conclusion GBS comprises a group of disorders of different pathophysiological patterns – AIDP, AMSAN, and AMAN. Patients present similarly with rapidly progressive weakness, areflexia, and albuminocytologic dissociation in the CSF. In AIDP, evidence of a primary demyelinating neuropathy is seen on electrodiagnostic studies. Although rarely indicated, nerve biopsies reveal endoneurial mononuclear inflammatory infiltrates along with evidence of demyelination. The majority of patients make a good recovery, especially with IVIG or PE treatment. In the axonal variants, AMSAN and AMAN, a primary axonal process is evident on electrodiagnostic and pathological studies. AMSAN patients differ from AIDP by their more severe course and longer time to recovery. AMAN patients, on the other hand, lack sensory abnormalities typically present in AIDP, while their prognosis is good. Recent work has suggested that molecular mimicry may be a possible pathogenetic mechanism for some patients with GBS. Targeting epitopes located in different portions of the peripheral nerve (Schwann cell in AIDP and axolemma in axonal variants) may account for the various presentations of GBS.


159

References 1

2 3 4 5 6 7 8 9 10 11 12 13 14 15

16

17

18 19

20

21

22 23

Griffin JW, Li CY, Ho TW, Xue P, Macko C, Gao CY, Yang C, Tian M, Mishu B, Cornblath DR, McKhann GM, Asbury AK: Guillain-Barre´ syndrome in northern China: The spectrum of neuropathological changes in clinically defined cases. Brain 1995;118:577–595. Ropper AH, Wijdicks EFM, Truax BT: Guillain-Barre´ Syndrome. Philadelphia, Davis, 1991. Moulin DE, Hagen N, Feasby TE, Amireh R, Hahn A: Pain in Guillain-Barre´ syndrome. Neurology 1997;48:328–331. Asbury AK, Cornblath DR: Assessment of current diagnostic criteria for Guillain-Barre´ syndrome. Ann Neurol 1990;27(suppl):S21–S24. Andersonn T, Siden A: A clinical study of the Guillain-Barre´ syndrome. Acta Neurol Scand 1982; 66:316–327. Ropper AH: The Guillain-Barre´ syndrome. N Engl J Med 1992;326:1130–1136. Lichtenfeld P: Autonomic dysfunction in the Guillain-Barre´ syndrome. Am J Med 1971;50:772–780. Barohn RJ, Kissel JT, Warmolts JR, Mendell JR: Chronic inflamatory demyelinating polyradiculoneuropathy. Arch Neurol 1989;46:878–884. Dyck PJ, Lais AC, Ohta M, Bastron JA, Okazaki H, Groover RV: Chronic inflammatory polyradiculoneuropathy. Mayo Clin Proc 1975;50:621–637. McCombe PA, Pollard JD, McLeod JG: Chronic inflammatory demyelinating polyradiculoneuropathy: A clinical and electrophysiological study of 92 cases. Brain 1987;110:1617–1630. Bouchard C, Lacroix C, Plante´ V, Adams D, Chedru F, Guglielmi JM, Said G: Clinicopathologic findings and prognosis of chronic inflammatory demyelinating polyneuropathy. Neurology 1999;52:498–503. Grand’Maison F, Feasby TE, Hahn AF, Koopman WJ: Recurrent Guillain-Barre´ syndrome: Clinical and laboratory features. Brain 1992;115:1093–1106. Asbury AK, Arnason BG, Adams RD: The inflammatory lesion in idiopathic polyneuritis: Its role in pathogenesis. Medicine 1969;48:173–215. Feasby TE, Gilbert JJ, Brown WF, Bolton CF, Hahn AF, Koopman WF, Zochodne DW: An acute axonal form of Guillain-Barre´ polyneuropathy. Brain 1986;109:1115–1126. McKhann GM, Cornblath DR, Griffin JW, Ho TW, Li CY, Jian Z, Wu HS, Zhaori G, Liu Y, Jou LP, Liu TC,Gao CY, Mao JY, Blaser MJ, Mishu B, Asbury AK: Acute motor axonal neuropathy: A frequent cause of acute flaccid paralysis in China. Ann Neurol 1993;33:333–342. Ho TW, Hsieh ST, Nachamkin I, Willison HJ, Sheikh K, Kiehlbauch J, Flanigan K, McArthur JC, Cornblath DR, McKhann GM, Griffin JW: Motor nerve terminal degeneration provides a potential mechanism for rapid recovery in acute motor axonal neuropathy after Campylobacter infection. Neurology 1997;48:717–724. Arnason BGW, Soliven B: Acute inflammatory demyelinating polyradiculoneuropathy, in Dyck PJ, Thomas PK, Griffin JW, Low PA, Poduslo JF (eds): Peripheral Neuropathy. Philadelphia, Saunders, 1993, pp 1437–1497. Rees JH, Gregson NA, Hughes RAC: Anti-ganglioside GM1 antibodies in Guillain-Barre´ syndrome and their relationship to Campylobacter infection. Ann Neurol 1995;38:809–816. Enders U, Karch H, Toyka KV, Michels M, Zielasek J, Pette M, Heesemann J, Hartung HP: The spectrum of immune responses to Campylobacter jejuni and glycoconjugates in Guillain-Barre´ syndrome and in other neuroimmunological disorders. Ann Neurol 1993;34:136–144. Ho TW, Mishu B, Li Cy, Gao CY, Cornblath DR, Griffin JW, Asbury AK, Blaser MJ, McKhann GM: Guillain-Barre´ syndrome in northern China: Relationship to Campylobacter jejuni infection and anti-glycolipid antibodies. Brain 1995;118:597–605. Ho TW, Willison HJ, Nachamkin I, Li CY, Veitch J, Ung H, Wang GR, Liu RC, Cornblath DR, Asbury AK, Griffin JW, McKhann GM: Anti-GD1a antibody is associated with axonal but not demyelinating forms of Guillain-Barre´ syndrome. Ann Neurol 1999;45:168–173. Blaser MJ: Epidemiologic and clinical features of Campylobacter jejuni infections. J Infect Dis 1997; 176(suppl 2):S103–105. Visser LH, van der Meche´ FGA, Meulstee J, Rothbarth PP, Jacobs BC, Schmitz PIM, van Doorn PA, Dutch Guillain-Barre´ Study Group: Cytomegalovirus infection and Guillain-Barre´ syndrome: The clinical, electrophysiologic, and prognostic features. Neurology 1996;47:668–673.

Bella

160

24

25

26

27

28

29

30 31 32

33 34

35

36

37

38 39 40

41 42 43 44

Safranek TJ, Lawrence DN, Kurland LT, Culver DH, Wiederholt WC, Hayner NS, Osterholm MT, O’Brien P, Hughes JM, the Expert Neurology Group: Reassessment of the association between Guillain-Barre´ syndrome and receipt of swine influenza vaccine in 1976–1977: Results of a twostate study. Am J Epidemiol 1991;133:940–951. Lasky T, Terracciano GJ, Magder L, Koski CL, Ballesteros M, Nash D, Clark S, Haber P, Stolley PD, Schonberger, LB, Chen RT: The Guillain-Barre´ syndrome and the 1992–1993 and 1993–1994 influenza vaccines. N Engl J Med 1998;339:1797–1802. Hughes RAC, Choudhary PP, Osborn M, Rees JH, Sanders EACM: Immunization and risk of relapse of Guillain-Barre´ syndrome or chronic inflammatory demyelinating polyradiculoneuropathy. Muscle Nerve 1996;19:1230–1231. Chiba A, Kusunoki S, Obata H, Machinami R, Kanazawa I: Serum anti-GQ1b IgG is associated with ophthalmoplegia in Miller-Fisher syndrome and Guillain-Barre´ syndrome: Clinical and immunohistochemical studies. Neurology 1993;43:1911–1917. Ilyas AA, Mithen FA, Dalakas MC, Chen ZW, Cook SD: Antibodies to acidic glycolipids in Guillain-Barre´ syndrome and chronic inflammatory demyelinating polyneuropathy. J Neurol Sci 1992;107:111–121. van den Berg LH, Marrink J, de Jager AEJ, de Jong HJ, van Imhoff GW, Latov N, Sadiq SA: AntiGM1 antibodies in patients with Guillain-Barre´ syndrome. J Neurol Neurosurg Psychiatry 1992; 55:8–11. Kuwabara S, Asahina M, Koga M, Mori M, Yuki N, Hattori T: Two patterns of clinical recovery in Guillain-Barre´ syndrome with IgG anti-GM1 antibody. Neurology 1998;51:1656–1660. Ho TW, Li C, Cornblath DR, Gao CY, Asbury AK, Griffin JW, McKhann GM: Patterns of recovery in the Guillain-Barre´ syndromes. Neurology 1997;48:695–700. Takigawa T, Yasuda H, Kikkawa R, Shigeta Y, Takahiko S, Kitasato H: Antibodies against GM1 ganglioside affect K+ and Na+ currents in isolated rat myelinated nerve fibers. Ann Neurol 1995; 37:436–442. Albers JW: AAEM case report #4: Guillain-Barre´ syndrome. Muscle Nerve 1989;12:705–711. Visser LH, Van der Mech? FGA, Van Doorn PA, Meulstee J, Jacobs BC, Oomes PG, Kleyweg RP, Dutch Guillain-Barre´ Study Group: Guillain-Barre´ syndrome without sensory loss (acute motor neuropathy): A subgroup with specific clinical, electrodiagnostic and laboratory features. Brain 1995;118:841–847. Feasby TE, Hahn AF, Brown WF, Bolton CF, Gilbert JJ, Koopman WJ: Severe axonal degeneration in acute Guillain-Barre´ syndrome: Evidence of two different mechanisms? J Neurol Sci 1993;116: 185–192. Hafer-Macko CE, Sheikh KA, Li CY, Ho TW, Cornblath DR, McKhann GM, Asbury AK, Griffin JW: Immune attack on the Schwann cell surface in acute inflammatory demyelinating polyneuropathy. Ann Neurol 1996;39:625–635. Hafer-Macko C, Hsieh S, Li CY, Ho TW, Sheikh K, Cornblath DR, McKhann GM, Asbury AK, Griffin JW: Acute motor axonal neuropathy: An antibody-mediated attack on axolemma. Ann Neurol 1996;40:635–644. Hartung HP, Pollard JD, Harvey GK, Toyka KV: Immunopathogenesis and treatment of the Guillain-Barre´ syndrome. Part I. Muscle Nerve 1995;18:137–153. Waksman BH, Adams RD: Allergic neuritis: Experimental disease of rabbits induced by peripheral nervous tissue and adjuvants. J Exp Med 1955;102:213–235. Hartung HP, Hughes RAC, Taylor WA, Heininger K, Reiners K, Toyka KV: T cell activation in Guillain-Barre´ syndrome and in MS: Elevated serum levels of soluble IL-2 receptors. Neurology 1990;40:215–218. Koski CL, Gratz E, Sutherland J, Mayer RF: Clinical correlation with anti-peripheral-nerve myelin antibodies in Guillain-Barre´ syndrome. Ann Neurol 1986;19:573–577. Feasby TE, Hahn AF, Gilbert JJ: Passive transfer of demyelinating activity in Guillain-Barre´ polyneuropathy. Neurology 1980;30:363. Saida T, Saida K, Silberberg DH, Brown MJ: Transfer of demyelination by intraneural injection of experimental allergic neuritis serum. Nature 1978;272:639–641. Dyck PJ, Kurtzke JF: Plasmapheresis in Guillain-Barre´ syndrome. Neurology 1985;35:1105–1107.


161

45

46

47

48

49 50 51 52

53 54 55

56 57 58

59 60 61 62 63 64 65

Yuki N, Taki T, Inagaki F, Kasama T, Takahashi M, Saito K, Handa S, Miyatake T: A bacterium lipopolysaccaride that elicits Guillain-Barre´ syndrome has a GM1 ganglioside-like structure. J Exp Med 1993;178:1771–1775. Sheikh KA, Nachamkin I, Ho TW, Willison HJ, Veitch J, Ung H, Nicholson M, Li CY, Wu HS, Shen BQ, Cornblath DR, Asbury AK, McKhann GM, Griffin JW: Campylobacter jejuni lipopolysaccarides in Guillain-Barre´ syndrome. Molecular mimicry and host susceptibility. Neurology 1998;51:371–378. Yuki N: Molecular mimicry between gangliosides and lipopolysaccharides of Campylobacter jejuni isolated from patients with Guillain-Barre´ syndrome and Miller-Fisher syndrome. J Infect Dis 1997; 176 (suppl 2):S150–S153. Ropper AH: ICU management of acute inflammatory-postinfectious polyneuropathy (LandryGuillain-Barre´-Strohl syndrome); in Ropper AH, Kennedy SK (eds): Neurological and Neurosurgical Intensive Care, ed 2. Rockville, Aspen, 1988, pp 253–268. Ropper AH, Kehne SM: Guillain-Barre´ syndrome: Management of respiratory failure. Neurology 1985;35:1662–1665. The Guillain-Barre´ Syndrome Study Group: Plasmapheresis and acute Guillain-Barre´ syndrome. Neurology 1985;35:1096–1104. French Cooperative Group on Plasma Exchange in Guillain-Barre´ Syndrome: Efficiency of plasma exchange in Guillain-Barre´ syndrome: Role of replacement fluids. Ann Neurol 1987;22:753–761. McKhann GM, Griffin JW, Cornblath DR, Mellits ED, Fisher RS, Quaskey SA, The GuillainBarre´ Syndrome Study Group: Plasmapheresis and Guillain-Barre´ syndrome: Analysis of prognostic factors and the effect of plasmapheresis. Ann Neurol 1988;23:347–353. The French Cooperative Group on Plasma Exchange in Guillain-Barre´ Syndrome: Appropriate number of plasma exchanges in Guillain-Barre´ syndrome. Ann Neurol 1997;41:298–306. Ropper AH, Albers JW, Addison R: Limited relapse in Guillain-Barre´ syndrome after plasma exchange. Arch Neurol 1988;45:314–315. van der Meche FGA, Schmitz PIM, Dutch Guillain-Barre´ Study Group: A randomized trial comparing intravenous immune globulin and plasma exchange in Guillain-Barre´ syndrome. N Engl J Med 1992;326;1123–1129. Irani DN, Cornblath DR, Chaudhry V, Borel C, Hanley DF: Relapse in Guillain-Barre´ syndrome after treatment with human immune globulin. Neurology 1993;43:872–875. Castro LHM, Ropper AH: Human immune globulin infusion in Guillain-Barre´ syndrome: Worsening during and after treatment. Neurology 1993;43:1034–1036. Plasma Exchange/Sandoglobulin Guillain-Barre´ Syndrome Trial Group: Randomised trial of plasma exchange, intravenous immunoglobulin, and combined treatments in Guillain-Barre´ syndrome. Lancet 1997;349:225–230. Dalakas MC: Mechanism of action of intravenous immunoglobulin and therapeutic considerations in the treatment of autoimmune neurologic diseases. Neurology 1998;51(suppl 5):S2–S8. Tan E, Hajinazarian M, Bay W, Neff J, Mendell JR: Acute renal failure resulting from intravenous immunoglobulin therapy. Arch Neurol 1993;50:137–139. The Dutch Guillain-Barre´ Study Group: Treatment of Guillain-Barre´ syndrome with high-dose immune globulins combined with methylprednisolone: A pilot study. Ann Neurol 1994;35:749–752. French Cooperative Group on Plasma Exchange in Guillain-Barre´ Syndrome: Plasma exchange in Guillain-Barre´ syndrome: One-year follow-up. Ann Neurol 1992;32:94–97. Lawn ND, Wijdicks EFM: Fatal Guillain-Barre´ syndrome. Neurology 1999;52:635–638. Bella IR, Chad DA: Guillain-Barre´ Syndrome; in Irwin RS, Cerra FB, Rippe JM (eds): Intensive Care Medicine, ed 4. Philadelphia, Lippincott-Raven, 1999, p 2117. Bella IR, Chad DA: Neuromuscular disorders and acute respiratory failure. Neurol Clin 1998;16:408.

Isabelita R. Bella, MD, Department of Neurology, UMass Memorial Health Care, 55 Lake Avenue North, Worcester, MA 01655 (USA) Tel. +1 508 856 4147, Fax +1 508 856 6778

Bella

162


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

Proximal Spinal Muscular Atrophy of Childhood Irena Hausmanowa-Petrusewicz a, Jacek Zaremba b a b

Neuromuscular Unit, Medical Research Centre, Polish Academy of Sciences and Department of Genetics, Institute of Psychiatry and Neurology, Warsaw, Poland

History Proximal childhood spinal muscular atrophy (SMA) is one of the most common and severe autosomal recessive diseases of children. In all spinal atrophies, dysfunction and loss of anterior horn cells leads to muscle atrophy and weakness sparing the sensory system and the central nervous system. The first description of SMA is attributed to the paper by Sevestre [1] in Paris. The affected child was symptomatic from birth, and severely paralyzed. In 1891, an Austrian neurologist Werdnig [2] described two brothers who had weak limbs from the age of 10 months and abnormal breathing; the legs were more affected than the arms. Werdnig also described the postmortem examination of 23 muscles, peripheral nerves and central nervous system. In 1893, Hoffmann [3], a neurologist in Heidelberg, described two families with similar symptoms which started 2 months after birth and led to death at the ages of 1, 2, 3 or 5 years. Hoffmann noticed general floppiness and more severely affected legs than arms. As opposed to the case of Sevestre both Werdnig’s and Hoffmann’s infantile cases had a more chronic course but were equally severe. The introduction of clinical electromyography (EMG) after World War II extended the concepts of SMA: the ‘dystrophy with fasciculations’ was proved to be juvenile spinal atrophy (Kugelberg and Welander syndrome) [4]. Subsequently there was a vivid debate between ‘lumpers’ and ‘splitters’ whether SMA represented two (or even three) different diseases. This uncertainty was later resolved by molecular geneticists.

Table 1. Three forms of proximal childhood SMA

Form 1 1a 1b

Onset

Survival

in utero (overt at birth) within 6 months

=2 years 2–4 years 8–10% of these children survive much longer but do not improve

Form 2

=18 months

several years 90% up to 10 years

Form 3 3a 3b

=3 years 3–17 years

normal life expectancy normal life expectancy

Clinical picture: form 1a infants: floppy, frog-like position, legs more paralyzed than arms, areflexia, abdominal breathing, weak cry, finger tremor, alert; form 1b: similar symptoms but progress slower than in group 1a; unable to lift the head and sit without support; form 2 (intermediate): proximal limb weakness and atrophy, joint contractures, scoliosis, chest deformities; achieve unaided sitting but cannot stand; form 3: proximal weakness, mostly in the legs; form 3a: slow progress leading to wheelchair after 5–7 years, and form 3b: ambulatory for a very long time.

Classifications of Childhood SMA and Clinical Features The main criteria used to classify SMA are age at onset of symptoms, course of the disease and age at death. We recognize three forms of proximal childhood SMA [5, 6] (table 1). Dubowitz [7] distinguished ten classes of varieties within each form; he also indicated the possibility of an SMA form 0: the most severe with a history of reduced fetal movement in utero, with asphyxia and severe universal muscle weakness at birth (including facial weakness) [8]. The inclusion and exclusion criteria of childhood SMA were approved by the International SMA Consortium at the European Neuromuscular Center [9]. The inclusion criteria correspond to clinical features described above and confirmed by neurogenic EMG and biopsy. Exclusion criteria summarized signs and symptoms considered incompatible with SMA such as involvement of diaphragm or extraocular muscles, mental retardation, marked slowing of nerve conduction velocity (CV), sensory deficit, congenital joint contractures, metabolic abnormalities and high CK activity.

Hausmanowa-Petrusewicz/Zaremba

164

Epidemiology The current incidence varies between 10 and 15 per 100,000 live births, which is higher than in the past because the detectability has improved. On the basis of these estimates the frequency of heterozygosity for autosomal recessive SMA would be 1:50. The prevalence of a chronic, milder form of SMA3 is about 1:83,000 of the general population [10]. In some countries consanguineous marriages are more frequent and the incidence of SMA is very high, e.g. in some Muslim countries. An exceptionally high incidence of infantile SMA was found in the Karaite community in Israel [11] – the prevalence of SMA there is 1:400, and carrier frequency 1:10.

Laboratory Findings No biochemical markers of SMA are known. Generally CK activity in SMA children is in the normal range. In small infants with SMA1, electromyographic studies (EMG) show fibrillations and fasciculations which are not very prominent and a typical spontaneous activity which is motor unit firing at a rate of 5–15 Hz [12]. During slight contraction, two populations of motor unit action potentials were found – one short and low and the second long and high [13]. There are no complex potentials. In SMA2 and 3, the spontaneous activity includes fibrillations, long positive waves and fasciculations. Quantitative EMG is characterized by long, high motor unit action potentials which are often complex (up to 75%) [13]. Nerve conduction studies in SMA2 and 3 have shown normal motor and sensory CV; after a few years it may slightly slow down. However, motor CV is slow in SMA1: the delay in maturation of CV in SMA1 children is obvious [14]. Morphological investigations of muscle are needed to recognize the neurogenic character of the weakness. In spite of the progress in molecular genetics muscle biopsy is necessary in the non-deleted cases or in some exceptional cases. Light microscopy in SMA1 shows clusters of small muscle fibers of 5to 8-lm diameter as well as normal and/or hypertrophic fibers. The proportion of small fibers is higher in severe cases [15]. They are different from the small atrophic denervated fibers in other neurogenic disorders: in SMA they are round, with a well-preserved architecture, centrally placed single nuclei and have a high activity of desmin, fetal and neonatal myosin, and are surrounded by N-CAM [16] (fig. 1). The fetal-like character of these changes is confirmed by electron microscopy which shows myotube-like fibers with large centrally placed nuclei and scanty sarcoplasm, and by the presence of immature fibers

Proximal Spinal Muscular Atrophy of Childhood

165

1

2 Fig. 1. SMA1b: musculus quadriceps femoris. Round small muscle fibers and a group of fibers of normal diameter. HE.¶420. Fig. 2. SMA1a: musculus quadriceps femoris. Myotube-like cell.¶9,400.

consisting of 2–3 cells at different stages of maturation surrounded by a common basal lamina [16] (fig. 2). Undifferentiated histochemical fiber types persist beyond the myotube stage. In SMA1, apoptosis of muscle is observed, which does not occur in the muscles of normal neonates. Nucleus and sarcoplasm show condensation, fragmentation and formation of apoptotic bodies [17].


166

Fig. 3. SMA3: musculus quadriceps femoris. Muscle fibers of normal diameter among the hypertrophic and atrophic fibers. HE.¶390.

Intramuscular nerves also reveal some signs of immaturity with multiaxonal and premyelinated fibers. The histopathological pattern in SMA3 is different and resembles the chronic denervation of mature muscle. The process in SMA3 begins in the perinatal period or later. Small mononucleated muscle fibers are among the normal and hypertrophic ones. The atrophic fibers are irregular, often angular; sometimes, only groups of nuclei are seen. Splitting and other secondary changes are superimposed. Target or targetoid fibers can be seen (fig. 3). In peripheral nerves, demyelination and wallerian degeneration or thin myelin sheaths are found. A higher percentage of thin fibers and a uniform length of their internodes regardless of fiber diameter [18] are described. Changes in ventral roots are similar to those in peripheral nerves [19]. Motoneurons of the spinal cord and motor nuclei of VI–XII cranial nerves show both atrophic and degenerative changes mostly expressed in lumbar and cervical parts of the spinal cord. Degenerating cells are shrunken, angulated and darkly stained. In electron microscopy, they show the presence of apoptosis. Some investigators suggest that immaturity of muscle, motoneurons, or both in SMA [20–22] is evidenced by the myotube-like appearance of muscle, undifferentiated metabolically, small muscle fibers, the multiaxonal character of intramuscular innervation, delay of nerve maturation, excessive death of motoneurons and apoptosis seen in muscles of SMA babies.


167

The hypothesis of a disturbed interaction between nerve and muscle in SMA was proposed by Vrbova and Lowrie [23]. They suggested that improper nerve-muscle interactions in SMA inhibit the conversion of motoneurons from growing to transmitting cells. They cannot cope with functional demands and are activated too early, before maturation, which leads to death. This happens extensively in the most severe form of SMA. If a motoneuron has more time for maturation before it becomes active, survival is better. As an alternative hypothesis to the arrest of maturation, Morrison [20] discussed the slow progressive death of neurons beginning in utero and continuing until death. In spite of many pieces of evidence that in nerve-muscle interaction the motoneuron defect is primary, some data suggest a possible primary role of the muscle in the SMA pathology [17, 22, 24]; thus, the problem is still unsolved.

Genetics The locus of all three forms of SMA is 5q11.2–13.3 on the basis of linkage studies [25]. In 1995, Melki et al. [26] detected deletions in SMA individuals within 5q13 and identified a candidate gene, the survival motor neuron (SMN) gene. Simultaneously, a Canadian group identified another candidate gene, the neuronal apoptosis inhibitory protein (NAIP) gene [27]. The region encompassing both SMN and NAIP contains an inverted duplication; the genes and polymorphic markers within this region have two copies – telomeric and centromeric (fig. 4). The region is particularly unstable, prone to deletions and other rearrangements. Another gene mapped to the 5q13 region is p44, which is a subunit of a basal transcription factor TFIIH [28]. The SMN gene contains eight exons that span 20 kb; the telomeric SMNt and centromeric SMNc (now named SMN1 and SMN2, respectively) differ only by five base pairs. SMN1 and SMN2 produce a transcript encoding 294 amino acids. Lefebvre et al. [26] found that 98.6% of patients had a deletion of SMN1 with the loss of exons 7 and 8 or exon 7 alone; the remaining patients had a single base and other small mutations. It is now a well-established fact that mutations in the SMN1 gene are the major determinants of the SMA phenotype. The NAIP gene containing 16 exons is extending over 60 kb and also has two copies. In the series of Roy et al. [27] 45% of SMA1 and 18% of SMA2 and SMA3 patients had a deletion of the NAIP gene. NAIPc and NAIPt differ, NAIPc does not contain exon 5 (present in NAIPt) and it is a pseudogene (unable to transcribe). The telomeric copy of the p44 gene is deleted in SMA at a frequency similar to that observed for the NAIP gene. Both the NAIP gene and the p44


168

Fig. 4. SMA duplicated chromosomal region according to Lefebvre et al. [33]. Cen> Centromere; Tel>telomere.

gene do not seem to be critical for the development of SMA although they are detected more frequently in the most severe form of SMA1, and NAIP is a good candidate for a modifier because of its antiapoptotic function. Scharf et al. [29] recently found a novel gene H4F5 situated very close to SMN1 and deleted in over 90% of SMA1. Therefore, the authors proposed that H4F5 may be a good candidate for a phenotypic modifier in SMA. There is now evidence that SMA2 and SMA3 may not be due to deletions but to gene conversion events in which SMN1 is replaced by SMN2. Normal individuals usually have two copies of SMN2 and two copies of SMN1 (one copy of each gene per chromosome). Conversion of SMN1 to SMN2 provides an increase of copies of SMN2 [30–32]. McAndrew et al. [30] found three (instead of two) copies of SMN2 in 1.9% of normals, 19% of SMA1 carriers and 34.5% of SMA2 and SMA3 carriers. Thus, the SMN2 product appears to partly compensate the absence of SMN1 [33]. However, an increased number of SMN2 copies have also been found in the severe form of SMA, and this can indicate that apart from SMN2 there are other factors which may modulate the SMA phenotype [33a]. The SMN protein is located in nuclear structures called ‘gems’ which are found in vicinity of coiled bodies (hence the term gems: gemini of coiled bodies) [34]. Interactions were shown of SMN protein with RNA-binding protein and with antiapoptotic protein Bcl-2 suggesting an antiapoptotic role of SMN [35]. It was also shown that the SMN protein was associated and colocalized in the cytoplasm and in gems with a novel SMN-interactive protein 1. The SMN-interactive protein 1 complex is involved in the cytoplasmic assembly of small nuclear ribonucleoproteins to spliceosomal RNAs and with the nuclear import of the small nuclear ribonucleoprotein complex [36]. These facts provide evidence of a likely part played by the SMN protein in the metabolism of RNA [reviewed in 24, 33, 37]. Immunohistochemical analysis revealed a marked deficiency of SMN protein in motor neurons in SMA1 and a reduction in SMA3 [38].


169

The role of the SMN protein in the pathogenesis of SMA is not known. An important clue to this problem may be the finding of the SMN protein self-oligomerization defect in SMA and evidence that the severity of SMA correlates with the intracellular concentration of oligomerization-competent SMN proteins [39]. In spite of a difference of five nucleotides between SMN1 and SMN2 genes, they encode an identical protein. The question arose, therefore, why SMN2 was unable to fully compensate the loss of SMN1. The answer was provided only recently by Lorson et al. [40] who found that it was due to one nucleotide exchange – CKT at codon 280 – a change which dictates exon 7 alternative splicing and production of SMN2-defective mRNA which lacks exon 7. This was a very important step forward in the elucidation of SMA pathogenesis and a finding which according to the authors may be helpful in designing a gene therapy targeted at the repair of the SMN2 gene transcript. The above data show remarkable progress in the studies on the molecular basis of SMA. There remains, however, a number of phenomena which seem to be also related to epigenetic factors, i.e. those altering the activity of the SMN gene without changing its structure. These are in particular incomplete penetrance of the mutated genes and gender influence, which are characteristic mainly of the mildest form of SMA. Investigations based on traditional segregation analysis showed a predominance of males in the mildest SMA3 group and revealed incomplete penetrance of the mutated gene, particularly in females after an age at onset of 8 years. These findings could suggest the existence of a ‘female sparing factor’ possibly connected with some milestones of female hormonal development [41, 42]. More recent investigations not only confirmed the predominance of males in the mildest forms of SMA [43] but also provided molecular evidence of incomplete penetrance of the mutated gene in some families: unaffected individuals were found in SMA families among carriers of homozygous deletions of exons 7 and 8 of the SMN gene, and the majority of them were females [44, 45]. It is worthwhile noting that the influence of sex is also characteristic of some other motor neuron diseases, such as the distal form of SMA [46] and amyotrophic lateral sclerosis [47, 48]. It is tempting to speculate that some sex steroids, possibly owing to a mechanism involving their nuclear receptors, may be capable of upregulating transcription of SMN2 and/or antiapoptotic genes, particularly in the mildest SMA3 form of the disease. Obviously other mechanisms of the positive influence of sex steroids on the clinical picture of SMA are also conceivable.


170

Differential Diagnosis (A) With other diseases: The differential diagnosis of SMA1 includes all conditions that cause the floppy baby syndrome: general (e.g. system disorders), cerebellar, congenital neuromuscular diseases such as myasthenia gravis, nemaline myopathy, myotubular myopathy, congenital mitochondrial myopathies, congenital myotonic dystrophy, congenital neuropathies. In SMA3, the differentiation with muscular dystrophy or polymyositis is most important. (B) The differentiation between SMA and related (apparently similar) neurogenic disorders is more difficult. This large group includes: (1) monomelic forms of SMA, e.g. Ryukyuan disease – still not mapped [49], a juvenile distal form of Hirayama et al. [50], an asymmetric, nongenetic syndrome, (2) distal SMA – a heterogeneous group [51], particularly important is diaphragmatic SMA [52] with a characteristic involvement of the diaphragm; it does not map to 5q, and (3) bulbar forms, including the Fazio-Londe syndrome [53, 54]. (C) SMA-plus: Cases not mapped to chromosome 5q with some classical clinical features of SMA but also with some symptoms considered as exclusion criteria in the diagnosis of SMA, e.g. SMA with mental retardation, with olivopontocerebellar features [55, 56], with ophthalmoplegia or oculo-pharyngeal syndrome [57]. Most important, however, are the descriptions of disorders presenting the same mutations as in classical SMA but entirely distinct phenotypes considered until recently to be incompatible with SMA. SMN deletions have been found in some cases of congenital axonal neuropathies with extremely slow CV [58] or in some cases of arthrogryposis [59, 60] which is against the formerly accepted exclusion criteria [9]. Obviously, not all cases of congenital neuropathy or arthrogryposis map to chromosome 5q. The neurogenic form of arthrogryposis has been considered by some authors a variant of SMA but contractures are not found in typical SMA (except minor contractures of some joints). Moreover, SMA is familial while even severe arthrogryposis is mostly sporadic and unlike SMA improves with time. Infrequent cases of familial arthrogryposis (autosomal recessive or X-linked) with fractures of the long bones were previously diagnosed as SMA [61]. The severe subset of arthrogryposis seems to be allelic with SMA. Some cases of congenital heart disease coexisting sometimes with classical SMA show SMN deletion [62].

Treatment In clinical practice, attention is given to ventilatory support, physical therapy, orthopedic devices and surgery. Physical and orthopedic treatment


171

maintains late ambulation and prevents contractures. The scoliosis sometimes requires surgery (fusion). Some neurotrophic factors enhance motoneuron survival in vitro and therapeutic trials are being conducted. Until now no clinical application has been tried. Regarding the gene therapy of SMA, some hope may be attached to already established molecular characteristics of SMA and to the role played by the modifying genes. Understanding basic molecular features of SMN2 which are alternative splicing of exon 7 [40] and special ability of SMN2 to convert to SMA1 seems to make SMN2 the best candidate and a target for gene therapy. Finding ways of suppressing apoptosis of the motoneurons could be a possible therapeutic approach, e.g. through activation of NAIP and/or other antiapoptotic genes, such as Bcl-2. Disclosing molecular mechanisms of the lack of clinical expression in homozygous carriers of exon 7 and 8 deletion of SMN1 [44, 45] might provide a clue for the future approach to gene therapy of SMA.

Genetic Counselling and Prenatal Diagnosis SMA conforms to the autosomal recessive pattern of inheritance, which makes estimation of the genetic risk a simple matter. The genetic risk value 1 in 4 should be given, although in SMA3 it was found to be smaller, particularly for females due to the incomplete penetrance of the mutated gene and to the sex influence [41, 42]. The risk for a heterozygous healthy sibling of an affected homozygous patient of having an affected child is 2/3¶1/50¶1/4>1/300, and the risk of having such a child for a midly affected or unaffected (nonmanifesting) individual with a homozygous mutation of the SMN1 gene is 1:100 (assuming a frequency of heterozygotes in the general population of 1:50). Sometimes the genetic risk is not high even for the couples who previously gave birth to an affected child, since 2% of the cases are due to new mutations of the SMN1 gene in one of the parents [63]. Detection of such cases is more complicated than simply finding a homozygous deletion, because it is based on linkage analysis in unaffected siblings of the proband. Moreover, one should bear in mind that even if we find in a healthy child the same haplotype as in an affected one, there is still a possibility of germinal mosaicism in the mother. The prenatal diagnosis is based on the test which identifies homozygous deletion of exon 7 of SMN1 gene [see commentary, 64]. False-negative results occur in ~5% of the cases. Most of them must be compound heterozygotes (with deletion of SMN1 in one chromosome and with another mutation in the other) and the remaining, very infrequent cases are homozygotes for small mutations in the SMN1 gene [65].


172

The prenatal test should be offered to all couples having a genetic risk of 1/4, but should also be available to those with an increased but much lower risk – up to 1/300. One important condition, however, is a previous identification of a homozygous deletion in the affected individual from the same family. This condition does not have to be treated as absolute, if there are good grounds for assuming that the clinical diagnosis of SMA in the proband – whose DNA is not available – was correct. Absence of homozygous deletion in such cases reduces the risk of SMA from 25 to 2% [65]; the 2% risk is due to the infrequent occurrence of small mutations and, possibly, a wrong clinical diagnosis of SMA in some cases. Detection of carriers and compound heterozygotes is feasible [30, 66] but difficult. Therefore, the SMA Consortium at a meeting in 1997 acknowledged the fact that prenatal diagnosis in the absence of homozygous deletion of SMN1 in the proband is hazardous and advised to refrain from prenatal tests in such cases [64].

References 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

Sevestre M: Paralysie flasque des quatre membres et des muscles du tronc (sauf le diaphragme) chez un nouveau-ne´. Bull Soc Pe´diatr Paris 1899;1:7–13. Werdnig G: Zwei fru¨hinfantile heredita¨re Fa¨lle von progressiver Muskelatrophie unter dem Bilde der Dystrophie, auf neurotischer Grundlage. Arch Psychiatr Nervenkr 1891;22:437–481. ¨ ber chronische spinale Muskelatrophie im Kindesalter, auf familia¨rer Basis. Dtsch Hoffmann J: U Z Nervenheilkd 1893;3:427–470. Kugelberg E, Welander L: Heredofamilial juvenile muscular atrophy simulating muscular dystrophy. Arch Neurol Psychiatr 1956;75:500–509. Emery AEH: Diagnostic Criteria for Neuromuscular Disorders. Baarn, ENMC, 1994, pp 48–54. Zerres K: Klassifikation und Genetik spinaler Muskelatrophien. Stuttgart, Thieme, 1989, pp 15–21. Dubowitz V: Chaos in the classification of SMA: A posible resolution. Neuromuscul Disord 1995; 5:3–5. Dubowitz V: Very severe spinal muscular atrophy (SMA type 0): An expanding clinical phenotype. Eur J Pediatr Neurol 1999;3:49–51. Munsat TL, Davies KE: Meeting report: International SMA Consortium meeting. Neuromuscul Disord 1992;2:424–428. Pearn J: Segregation analysis of chronic childhood spinal muscular atrophy. J Med Genet 1978;15: 414–417. Fried K, Mundel G: High incidence of spinal muscular atrophy type I (Werdnig-Hoffmann disease) in the Karaite community in Israel. Clin Genet 1977;12:250–251. Buchtal F, Olsen PZ: Electromyography and muscle biopsy in infantile spinal muscular atrophy. Brain 1970;93:15–30. Hausmanowa-Petrusewicz I, Karwan´ska A: Electromyography findings in different forms of infantile and juvenile proximal spinal muscular atrophy. Muscle Nerve 1986;9:36–46. Renault F, Chartier JP, Harpey JP: Apport de l’e´lectromyogramme au diagnostic d’amyotrophie spinale infantile en pe´riode neónatale. Arch Pediatr 1996;3:319–323. Dubowitz V: Muscle Biopsy – A Practical Approach, ed 2. London, Baillie`re, Tindall, 1985. Fidzian´ska A: Spinal muscular atrophy in childhood. Semin Pediatr Neurol 1996;32:53–58.


173

17 18 19 20 21 22

23

24 25

26

27

28

29

30

31

32 33 33a

34 35

Fidzian´ska A, Goebel HH, Warlo J: Acute infantile spinal muscular atrophy. Muscle apoptosis as a proposed pathogenetic mechanism. Brain 1990;113:433–445. Drac H: Morphological changes in peripheral nerves in Werdnig-Hoffmann disease. Neuropathol Pol 1977;15:1–10. Chou SM: Glial bundles of nerve roots in Werdnig-Hoffmann disease. Ann Neurol 1980;8:79–81. Morrison KE: Advances in SMA research: Review of gene deletions. Neuromuscul Disord 1996; 6:397–408. Hausmanowa-Petrusewicz I, Fidzian´ska A, Niebro´j-Dobosz I, Strugalska-Cynowska MH: Is Kugelberg-Welander spinal muscular atrophy a fetal defect? Muscle Nerve 1980;3:389–402. Hausmanowa-Petrusewicz I, Fidzian´ska-Dolot A: Clinical features of infantile and juvenile spinal muscular atrophy; in Gamstorp I, Sarnat HB (eds): Progressive Spinal Muscular Atrophies. New York, Raven Press, 1984, pp 31–42. Vrbova G, Lowrie MB: The role of nerve-muscle interactions in the pathogenesis of spinal muscular atrophy; in Merlini L, Granata C, Dubowitz V (eds): Current Concepts in Childhood Spinal Muscular Atrophy. Vienna, Springer, 1989, pp 33–40. Gendron NH, MacKenzie AE: Spinal muscular atrophy: Molecular pathophysiology. Curr Opin Neurol 1999;12:137–142. Brzustowicz LM, Lehner T, Castilla LH, Penchaszadeh GK, Wilhelmsen KC, Daniels R, Davies KE, Leppert M, Ziter F, Wood D, Dubowitz V, Zerres K, Hausmanowa-Petrusewicz I, Ott J, Munsat LM, Gilliam TC: Genetic mapping of chronic childhood-onset spinal muscular atrophy to chromosome 5q11.2-13.3. Nature 1990;344:540–541. Lefebvre S, Burglen L, Reboullet S, Clermont O, Burlet P, Viollet L, Benichou B, Cruanud C, Millasseau P, Zeviani M, Le Paslier S, Frezal J, Cohen D, Weissenbach J, Munnich A, Melki J: Identification and characterization of a spinal muscular atrophy-determining gene. Cell 1995;80: 155–165. Roy N, Mahadevan MS, McLean M, Shutler G, Yaraghi Z, Farahani R, Baird S, Besner-Johnston A, Lefebvre C, Kang X, Salih M, Aubry H, Tamai K, Guan X, Ioannou P, Crawford TO, de Jong PJ, Surh L, Ikeda J-E, Korneluk RG, MacKenzie A: The gene for neuronal apoptosis inhibitory protein is partially deleted in individuals with spinal muscular atrophy. Cell 1995;80:167– 178. Bu¨rglen L, Seroz T, Miniou P, Lefebvre S, Burlet P, Munnich A, Viegas-Pequignot E, Egly JM, Melki J: The gene encoding p44, a subunit of the transcription factorTFIIH, is involved in largescale deletions associated with Werdnig-Hoffmann disease. Am J Hum Genet 1997;60:72–79. Scharf JM, Endrizzi MG, Wetter A, Huang S, Thompson TG, Zerres K, Dietrich WF, Wirth B, Kunkel LM: Identification of a candidate modifying gene for spinal muscular atrophy by comparative genomics. Nat Genet 1998;20:83–86. McAndrew PE, Parsons DW, Simard LD, Rochette C, Ray PN, Mendell JR, Prior TW, Burghes AHM: Identification of proximal spinal muscular atrophy carriers and patients by analysis of SMNt and SMNc gene copy number. Am J Hum Genet 1997;60:1411–1422. Campbell L, Potter A, Ignatius J, Dubowitz V, Davies K: Genomic variation and gene conversion in spinal muscular atrophy: Implications for disease process and clinical phenotype. Am J Hum Genet 1997;61:40–50. Burghes AHM: When is a deletion not a deletion? When it is converted. Am J Hum Genet 1997; 61:9–15. Lefebvre S, Bu¨rglen L, Frezal, Munnich A, Melki J: The role of SMN gene in proximal spinal muscular atrophy. Hum Mol Genet 1998;7:1531–1536. Talbot K, Rodrigues NR, Ignatius J, Muntoni F, Davies KE: Gene conversion at the SMA locus in autosomal recessive spinal muscular atrophy does not predict a mild phenotype. Neuromuscul Disord 1997;7:198–201. Liu Q, Dreyfuss G: A novel nuclear structure containing the survival of motor neurons protein. EMBO J 1996;15:3555–3565. Iwahashi H, Eguchi Y, Yasuhara N, Hanafusa T, Matzuzawa Y, Tsujimoto Y: Synergistic antiapoptotic activity between Bcl-2 and SMN implicated in spinal muscular atrophy. Nature 1997; 390:413–417.


174

36

37 38

39

40 41 42 43

44

45

46 47 48 49 50

51 52

53 54 55

56

57 58

Liu Q, Fischer U, Wang F, Dreyfuss G: The spinal muscular atrophy disease gene product, SMN, and its associated protein SIP1 are in a complex with spliceosomal snRNP proteins. Cell 1997;90: 1013–1021. Mattaj IW: Ribonucleoprotein assembly: Clues from spinal muscular atrophy. Curr Biol 1998;8: R93–R95. Lefebvre S, Burlet P, Liu Q, Bertrandy S, Clermont O, Munnich A, Dreyfuss G, Melki J: Correlation between severity and SMN protein level in spinal muscular atrophy. Nat Genet 1997;16:265– 269. Lorson CL, Strasswimmer J, Yao J-M, Baleja JD, Hahnen E, Wirth B, Le T, Burghes AHM, Androphy EJ: SMN oligomerization defect correlates with spinal muscular atrophy severity. Nat Genet 1998;19:63–66. Lorson CL, Hahnen E, Androphy EJ, Wirth B: A single nucleotide in the SMN gene regulates splicing and is responsible for spinal muscular atrophy. Proc Natl Acad Sci USA 1999;96:6307–6311. Hausmanowa-Petrusewicz I, Zaremba J, Borkowska J, Prot J: Genetic investigations on chronic form of infantile and juvenile spinal muscular atrophy. J Neurol 1976;213:335–346. Hausmanowa-Petrusewicz I, Zaremba J, Borkowska J, Szirkowiec W: Chronic proximal spinal muscular atrophy of childhood and adolescence: Sex influence. J Med Genet 1984;21:447–450. Zerres K, Rudnik-Schoneborn S, Forrest E, Łusakowska A, Borkowska J, Hausmanowa-Petrusewicz I: A collaborative study on the natural history of childhood and juvenile onset proximal spinal muscular atrophy (type II and III SMA): 569 patients. J Neurol Sci 1997;146:67–72. Cobben JM, van der Steege G, Grootscholten P, de Visser M, Scheffer H, Buys CHCM: Deletions of the survival motor neuron gene in unaffected siblings of patients with spinal muscular atrophy. Am J Hum Genet 1995;57:805–808. Hahnen E, Forket R, Marke C, Rudnik-Schoneborn S, Schonling J, Zerres K, Wirth B: Molecular analysis of candidate genes on chromosome 5q13 in autosomal recessive spinal muscular atrophy: Evidence of homozygous deletions of the SMN gene in unaffected individuals. Hum Mol Genet 1995;4:1927–1933. Harding AE, Thomas PK: Hereditary distal spinal muscular atrophy. J Neurol Sci 1980;45:337– 345. Mackey RP: Course and prognosis in amyotrophic lateral sclerosis. Arch Neurol 1963;8:117–127. Rosen AD: Amyotrophic lateral sclerosis. Arch Neurol 1978;35:638–642. Kondo K, Tsubaki T, Sakarmoto F: The Ryukyuan muscular atrophy. An obscure heritable neuromuscular disease found in the island of southern Japan. J Neurol Sci 1970;11:359–364. Hirayama K, Tomanoga M, Kitano K, Yamada T, Kojima S, Arai K: Focal cervical poliopathy causing juvenile muscular atrophy of distal upper extremity: A pathological study. J Neurol Neurosurg Psychiatry 1987;50:285–290. Pearn J, Hudgson P: Distal spinal muscular atrophy: A clinical and genetic study of eight kindreds. J Neurol Sci 1979;43:183–191. Bertini E, Gadisseux JL, Palmieri G, Ricci E, DiCapua M, Ferriere G, Lyon G: Distal infantile spinal muscular atrophy associated with paralysis of the diaphragm: A variant of infantile spinal muscular atrophy. Am J Med Genet 1989;33:328–335. Fazio M: Eridetarieta della paralise bulbare progressive. Riforme Med 1892;8:327–331. Londe P: Paralysie bulbaire progressive infantile et familiale. Rev Med 1894;14:212–215. Chou SM, Gilbert RF, Chun RWM, Laxowa R, Tuffi GA, Sufit RL, Krassikot N: Infantile olivopontocerebellar atrophy with spinal muscular atrophy (infantile OPCA+SMA). Clin Neuropathol 1990; 9:21–32. Rudnik-Schoneborn S, Wirth B, Rohrig D, Saule H, Zerres K: Exclusion of the gene locus for spinal muscular atrophy on chromosome 5q in a family with olivopontocerebellar atrophy (OPCA) and anterior horn cell degeneration. Neuromuscul Disord 1995;5:19–23. Aberfeld DC, Numba T: Progressive ophthalmoplegia in Kugelberg-Welander disease. Arch Neurol 1969;20:253–262. Korinthenberg R, Sauer M, Ketelsen UP, Hanemann CO, Stoll G, Graf M, Barborie A, Volk B, Wirth B, Rudnik-Schoneborn S, Zerres K: Congenital axonal neuropathy caused by deletions in the spinal muscular atrophy region. Ann Neurol 1997;42:364–368.


175

59

60

61 62 63

64 65

66

Burglen L, Amiel J, Viollet L, Lefebvre S, Burlet P, Clermont O, Raclin V, Landrieu P, Verloes A, Munnich A, Melki J: Survival motor neuron gene deletion in the arthrogryposis multiplex congenitaspinal muscular atrophy association. J Clin Invest 1996;98:1130–1132. Bingham P, Shen N, Rennert H, Rorke LB, Black AW, Marin-Padilla M, Nordgren RE: Arthrogryposis due to infantile neuronal degeneration associated with deletion of the SMNt gene. Neurology 1997;49:848–851. Greenberg F, Fenolio KR, Hejtmancik JF, Armstrong D, Willis JK, Shapira E, Huntington HW, Haun RL: X-linked infantile spinal muscular atrophy. Am J Dis Child 1988;142:217–219. Burglen L, Spiegel R, Ignatius J, Cobben JM, Landrien P, Lefebvre S, Munnich A, Melki J: SMN gene deletion in variant of infantile spinal muscular atrophy. Lancet 1995;346:306–317. Wirth B, Schmidt T, Hahnen E, Rudnik-Schoneborn S, Krawczak M, Muller-Myhsok B, Schonling J, Zerres K: De novo rearrangements found in 2 percent of index patients with spinal muscular atrophy: Mutation mechanisms, parental origin, mutation rate, and implications for genetic counselling. Am J Hum Genet 1997;61:1102–1111. Matthijs G, Devriendt K, Fryns JP: The prenatal diagnosis of spinal muscular atrophy. Prenat Diagn 1998;18:607–610. Somerville MJ, Hunter AGW, Aubry HL, Korneluk RG, MacKenzie AE, Surh LC: Clinical application of the molecular diagnosis of spinal muscular atrophy: Deletions of neuronal apoptosis inhibitor protein and survival motor neuron genes. Am J Med Genet 1997;69:159–165. Parsons DW, McAndrew PE, Allinson PS, Parker WD Jr, Burghes AHM, Prior TW: Diagnosis of spinal muscular atrophy in an SMN non-deletion patient using a quantitative PCR screen and mutation analysis. J Med Genet 1998;35:674–676.

Irena Hausmanowa-Petrusewicz, Neuromuscular Unit, Polish Academy of Sciences, Banacha 1a, bl.D, Room 718, PL–02097 Warsaw (Poland) E-Mail: [email protected]


176


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

Familial Amyotrophic Lateral Sclerosis: A Review Mitsuya Morita, Robert H. Brown, Jr. Day Neuromuscular Research Center, Charlestown, Mass., USA

Amyotrophic lateral sclerosis (ALS), a degenerative disorder of cortical and spinal motor neurons, causes a progressive paralysis that is usually fatal because of respiratory weakness within 5 years of onset. The cause of the progressive degeneration of motor neurons in this disease remains unknown and these is no definitive treatment. The prevalence of ALS is about 5:100,000 and its incidence is approximately 1:100,000. In a subset of cases (approximately 10% of ALS cases), it is apparent that the disease is inherited, often as a dominant trait [1]. Since the clinical and pathological characteristics of familial ALS (FALS) and sporadic ALS (SALS) are almost identical, we and others have presumed that understanding of the molecular defects that cause FALS will provide insights into the pathogenesis of both FALS and SALS. Although several loci have been linked to FALS, the only well-established cause of FALS are defects in the gene encoding Cu/Zn-superoxide dismutase (CuZn-SOD; SOD1). In this chapter, we summarize recent developments in FALS research, consider how mutant SOD1 causes motor neuron cell death and evaluate possible therapies for ALS.

Genetic Linkage Analysis in FALS In 1991, we and collaborators [2] established that the gene defect in some FALS pedigrees maps to chromosome 21q22.1-22.2. Because some families were not linked to this locus, this investigation indicated that there is genetic heterogeneity in FALS [2]. In 1993, Rosen et al. [3] in the same collaborative group reported that some cases of FALS are associated with mutations in the gene at this locus encoding SOD1. The first report described 11 different

missense mutations in the SOD1 gene in 13 FALS pedigrees. We now recognize that approximately one quarter of FALS pedigrees arises from mutations in the SOD1 gene [4, 5]. As of this writing, more than 70 mutations have been reported among FALS and (to a considerably lesser extent) SALS cases [6]; these are detected exclusively in individuals with motor neuron disease. That these mutations can cause ALS is strikingly underscored by the observation that several lines of mice with transgenes for mutant SOD1 develop a progressive, lethal paralysis that is clinically and pathologically highly homologous to human ALS [7–10]. An unusual, juvenile-onset form of ALS was described in seventeen Tunisian families by Ben Hamida et al. [11] in 1990. This was characterized by recessive inheritance, early onset (typically at about 12 years), and very slow disease progression. To date, the neuropathology of this ALS variant has not been defined. Ben Hamida et al. recognized three forms of juvenile ALS. Type 1, the most common form, is characterized by distinctive lower motor neuron involvement early in the disease, with denervational atrophy and weakness in distal limb muscles and ultimately the tongue as well. This form was genetically associated with a locus at 15q15-q22 [12]. Clinically, more prominent upper and less evident lower motor neuron features mark type 2 juvenile ALS. A locus for type 2 juvenile ALS has not been mapped. In type 3 juvenile ALS, the distinctive feature is prominent spasticity of both face and limb muscles; hand and peroneal atrophy are mild. Type 3 ALS has been mapped to 2q33q35 [13, 14] in a region spanning 1.7 cM between markers D2S116 and D2S2237 [15]. A clinically distinct ALS-like disorder with juvenile onset (late teen years) and very slow progression has been described in a large American pedigree of English descent. Affected individuals develop distal limb denervation and wasting associated with corticospinal signs, but do not develop respiratory or bulbar paralysis. This disease is transmitted as an autosomal dominant trait that has recently been linked to 9q34 [16, 17]. X-linked dominant ALS with clinical and neuropathological features indistinguishable from other types of ALS has been found in a single large American pedigree. The causative gene defect is not known [18].

Pathophysiology of FALS Linked to the Mutant SOD1 SOD1 is an abundant, cytoplasmic enzyme consisting of 153 amino acids encoded by five small exons. Each SOD1 molecule binds one atom of copper and one of zinc. Functional SOD1 is homodimeric. The enzyme is extremely stable, perhaps because the two SOD1 molecules are tightly packed together via powerful hydrophobic interactions [19]. The biological importance of

Morita/Brown

178

SOD1 may be gauged by its wide expression pattern (it is expressed in all eukaryotic cells), its abundance (about 0.5–2% of the soluble proteins in the human brain [20, 21]) and its high degree of conservation during evolution. Through cyclic reduction and oxidation of Cu, SOD1 detoxifies or dismutates intracellular superoxide anion to form hydrogen peroxide (H2O2). H2O2 is subsequently converted to H2O by catalase or glutathione peroxidase. Although the mutant forms of SOD1 have been investigated in several laboratories, the molecular basis for the neurotoxicity of mutant SOD1 protein is not well understood. However, several lines of evidence suggest that the mutant SOD1 molecule has acquired adverse, cytotoxic properties. First, some of the SOD1 mutations (e.g. G37R, D90A) do not substantially reduce dismutation activity [22, 23]. Second, mice that are totally devoid of SOD1 (SOD1 knockout mice) do not show developmental defects and, in contrast with mice with mutant SOD1 transgenes, do not subsequently develop motor neuron cell death [24]. Finally, for many of the mutant mouse strains, findings of motor neuron cell death develop in the face of above-normal total dismutation activity [7–10]. In contrast, control mice expressing equivalent levels of wild-type SOD1 do not develop such paralysis. Despite the fact that the age of onset in FALS pedigrees with high penetrance may be earlier than in SALS [25, 26], FALS patients with SOD1 defects are clinically indistinguishable from SALS or FALS patients without a known SOD1 mutation [15]. In pedigrees with diminished disease penetrance, there may be intra- or interfamilial phenotypic differences [27]. For some mutations, the phenotype tends to be rather stereotypical. The A4V (alanine changed to valine at position 4) mutation, detected in about half of all North American families with SOD1 mutations [15, 28], triggers a disease with a survival time of about 12–18 months [15]. Also, on clinical and pathological study, A4V patients reveal almost exclusively lower motor neuron involvement [29]. In A4V cases, there is more widespread nonmotor pathology than SALS [29, 30]. In contrast with the A4V cases, others (including E21G [31], G37R [15, 28], G41D [15], H46R [28, 32], D90A [33], G93C [15], L144S [34], L144F [28], I151T [35]) may occasionally survive for 10–15 years. Particularly striking is the D90A mutation, most commonly encountered in northern Scandinavia. In that setting, the resulting motor neuron disease is inherited as a recessive trait; individuals heterozygous for D90A are asymptomatic. Moreover, the disorder that results when D90A is homozygous is highly atypical for ALS, with a very slow onset over several years characterized by pain in the lower body and cramps in the muscles of the lower extremities [23, 33]. Those patients typically survive 20 years or more. In contrast, individuals in Europe or the United States who are heterozygous for D90A survive no longer than other ALS patients in those regions. These observations raise

Familial Amyotrophic Lateral Sclerosis

179

the exciting possibility that there is a genetic modifying factor within the Scandinavian population that in some manner mitigates the neurotoxic influences of the mutant SOD1 allele. Given the phenotypic diversity that can be encountered in families with different SOD1 mutations, one might anticipate that subtle but important differences in the biophysical properties of the different mutant proteins could explain phenotypic differences. However, studies to date have failed to support this possibility. Thus, Ratovitski et al. [36] examined the relationship between several biochemical properties (e.g. dismutation activity, polypeptide half-time, resistance to proteolytic degradation, solubility, etc.) of mutant SOD1 proteins and disease phenotype; no specific biophysical parameter correlated with clinical phenotype [36].

Neurotoxicity of Mutant SOD1 Protein Several hypotheses have been proposed to explain the toxicity of mutant SOD1 protein. Each proposed mechanism is based on the observation that the mutant SOD1 protein can become unstable and misfold. Two hypotheses are predicated on aberrant copper catalysis within the mutant protein. One argument is that the SOD1 mutations render the copper more accessible to peroxynitrite, allowing the formation of reactive nitronium-like intermediates that can nitrate tyrosine residues [37]. Some studies documented the elevation of nitrotyrosine in G37R transgenic mice [38] and SOD1-related FALS patients [39]. Another catalytic mechanism suggests that the mutations increase the peroxidase activity of SOD1, leading to the formation of more hydroxyl radicals from hydrogen peroxide [40, 41]. A second, broad hypothesis to explain the injurious properties of the mutant SOD1 molecule is that it is sufficiently unstable that, under some circumstances, it misfolds and forms aggregates that are in some manner toxic. Aggregated protein containing SOD1 has been detected in different transgenic ALS mice; it is possible that the mutant protein coaggregates with unidentified, essential component(s). These aggregates are often ubiquitinated. In a recent report, Bruijn et al. [42] genetically manipulated the levels of wild-type human SOD1 in ALS mice expressing the mutant SOD1 protein (G85R). These investigators found that changing the level of wild-type SOD1 (none to 6 times of the normal level of SOD1) did not affect the pathology, onset, or progression of disease and concluded that the aggregation of mutant SOD1 and its neurotoxicity are independent of the activity of the wild-type molecule. This finding has not been repeated in other strains of mice, and thus the generality of the conclusions from these experiments is not established as yet.

Morita/Brown

180

Autopsy studies have revealed aggregation and abnormal assembly of neurofilaments in the perikarya and axons of motor neurons in both SALS [43–45] and FALS with posterior column and spinocerebellar tract involvement [46]. Similar findings have been reproduced in transgenic mice with wild-type mouse NF-L subunits [47], wild-type human NF-H subunits [48], or mutant NF-L subunits [49]. Moreover, mutations in NF-H gene have been reported among sporadic ALS patients [50–52]. These findings raise the possibility that abnormalities in neurofilament organization may be involved in the pathogenesis of ALS. Marked neurofilamentous pathology is evident in FALS patients with the A4V [53], I113T [54] and H48Q [55] mutations in SOD1 as well as in transgenic mice expressing the G93A [56] protein. The evidence that disruption of NF-L or overexpression of human NF-H in mutant SOD1 transgenic mice delay onset of disease and/or extend life span suggests the possibility that neurofilaments modulate SOD1-mediated toxicity [57, 58]. However, it is uncertain whether accumulation of neurofilaments itself directly causes motor neuron cell death; in contrast with the above studies of mouse NH-L and human NF-H, forced expression of mouse NF-H does not result in any overt phenotype or enhanced motor neurodegeneration, despite severe perikaryal accumulation of neurofilaments and proximal axonal swellings in motor neuron [59]. Furthermore, recent reports that impairment of axonal transport was observed at an early, asymptomatic stage before any pathological changes in SOD1 transgenic mice [60–63] also suggest that accumulation of neurofilaments might be caused as a second phenomenon. Another important pathogenetic mechanism in SOD1-mediated ALS implicates the excitatory neurotransmitter glutamate. The extracellular concentration of glutamate in the central nervous system must be kept low to prevent neuronal damage from excessive activation of glutamate receptors [64]. One mechanism for this involves reuptake of glutamate by one or more astroglial transporters. EAAT2/GLT-1 in astrocytes is believed to be the predominant glutamate transporter involved in this regulation of extracellular glutamate levels in the nervous system [65]. Diminished EAAT2 function has been reported in ALS patients [66] and SOD1 transgenic mice [10]. An increase of glutamate in extracellular space was also reported in some patients [67]. The basis for this putative loss of glutamate transport activity is not clear. However, in one detailed study, variant mRNA transcripts for the EAAT2 glutamate transporter were detected in affected central nervous system tissue of 60% of autopsied SALS patients [68]. In vitro analyses indicate that the variant transcripts reduce expression of the EAAT2 protein, and thereby augment glutamate activity at synapses. As yet, these findings have not been duplicated in other studies [69, 70].


181

A recent report by Trotti et al. [71] demonstrated that FALS-linked SOD1 mutants can induce loss of function of EAAT2, particularly in the face of superimposed oxidative stress (e.g. addition of iron). These investigators argued that the mechanism for this inhibition of EAAT2 involves peroxidase activity of the mutant SOD1, because the chelation of the copper ion prevented EAAT2 inhibition, and because a similar pattern of EAAT2 inactivation was obtained by injecting Fe2+. Fe2+ catalyzes the production of highly reactive hydroxyl radicals via the Fenton reaction. In both human and mouse ALS, riluzole, which blocks glutamate release from presynaptic terminals, slows the disease course, albeit moderately. This also supports the possibility that glutamatemediated toxicity contributes to the pathogenesis of ALS. Other studies of ALS pathogenesis point to a role for mitochondrial dysfunction in this disease. Thus, abnormal mitochondria are evident early in the course of motor neuron degeneration in the G37R ALS mice [8]. Other studies show that the G93A SOD1 mutation alters electron transport enzymes in mitochondria and induces elevation of cytosolic calcium concentration [72]. Moreover, both in humans and mice with ALS, calcium is observed to accumulate in mitochondria within distal motor neuron terminals present in motor point biopsies in muscle [73, 74]. On these grounds, it is conceivable that mitochondrial dysfunction contributes to the pathogenesis of SOD1 transgenic mice. Two closely related concepts are that the mutant SOD1 protein alters the oxidative milieu within the motor neuron cytoplasm, setting the stage for oxidative injury to critical cellular constituents [75–77], and thereby triggering such adverse events as the aggregation of proteins and the activation of programmed cell death. Indeed, there are now several reports that mutant SOD1 protein can activate cell death genes [78–80]. It is clear, for example, that caspase-1 is activated both in the spinal cords of ALS mice and in serumdeprived, oxidatively stressed neuroblastoma cells that express mutant SOD1 protein [30].

Therapeutic Trials in ALS Transgenic mice with mutant SOD1 provide an excellent opportunity to dissect the etiology of this disease and to explore new avenues for ALS therapy. Although the exact mechanism how mutant SOD1 induces motor neuron cell death is unclear, several of the foregoing hypotheses lead directly to drug trials in the ALS mice. Thus, the hypothesis that oxidative metabolism is aberrant led to a trial of vitamin E in the ALS mice and then in ALS patients. As reported by Gurney et al. [81], dietary supplementation with vitamin E and

Morita/Brown

182

selenium causes a subtle delay in disease onset in the G93A mice but does not prolong survival once the disease begins. More recently, a French trial of vitamin E in ALS patients also showed a modest but significant benefit [82]. The hypothesis that glutamate toxicity is a factor in this disease also led Gurney et al. to explore the effect of two putative modulators of the glutamatergic system, riluzole and gabapentin. Both showed a modest benefit [81]. Based on the hypothesis that the mutant SOD1 molecule is toxic because of increased exposure to copper, the copper-chelating compound d-penicillamine was administered orally to G93A SOD1 transgenic mice; a modest delay in onset was seen [83]. The data that mitochondrial function, and hence ATP generation, are abnormal in ALS prompted Beal and colleagues [84] to try oral administration of creatine. At either 1 or 2% of the mouse diet, this compound improves the survival of G93A mice and prevents oxidative damage. Creatine is thought to help to buffer intracellular energy stores and inhibit mitochondrial transition pore opening. Since mutant SOD1 has been thought to be proapoptotic in motor neurons, two antiapoptotic genetic manipulations were tried in mice. These entailed (1) breeding the G93A animals to mice that overexpressed a dominant negative inhibitor of caspase-1 and (2) overexpressing the antiapoptotic protein, bcl-2, in the G93A mice. Bcl-2 is a well-described inhibitor of several cell death genes, including the interleukin-1b-converting enzyme (ICE). Friedlander et al. [85] crossed transgenic mice expressing a dominant negative inhibitor of ICE with a G93A mouse. The resulting F1 animals showed no difference in disease onset relative to the G93A mice without the ICE inhibitor. However, once the disease started, the doubly transgenic animals survive significantly longer from the onset of the disease (27 days) than the mutant SOD1 mice (11.7 days). Kostic et al. [86] crossed G93A mice with transgenic bcl-2 mice to determine whether the overexpression of human bcl-2 protects these ALS mice. Disease onset was significantly later (203 days) in the G93A/bcl-2 mice as compared with the G93A mice (170 days). On the other hand, the duration of the disease did not differ between these two groups. In this context, it is of considerable interest that some compounds that have not worked in human trials in ALS appear to have failed in the ALS mice as well. These prominently include the growth factors BDNF (brainderived neurotrophic factor), GDNF (glial-derived neurotrophic factor), CNTF (ciliary neurotrophic factor) and IGF-1 (insulin-like growth factor-1). As a final point, in considering the targets for therapy in ALS, it must be recalled that the death process may involve cell types other than the motor neuron itself. As the above discussion of EAAT2 indicates, one suspects that dysfunction within astrocytes may be important in this process. Several other lines of argument incriminate astrocytic and microglial dysfunction in ALS


183

[87–89]. Ultimately, strategies for slowing or reversing this disease may target both these nonneural cells and motor neurons. An extrapolation from this consideration is the more general point that in the long term, one also anticipates that effective therapy for ALS will require multiple agents, perhaps administered at different times in the illness. That is, if the evolution of the disease involves multiple stages (initiation, propagation, spreading or dissemination) then different agents may act differentially on these phases. Of course, this possibility – that there will be polydrug therapy for this illness – has precedence in cancer chemotherapy. Two final points should be noted regarding ALS therapeutics. First, for those cases that arise from inherited, abnormal proteins that are cytotoxic, the ultimate therapy will be to devise methods such as ribozyme or somatic cell gene mutation therapies to inactivate the toxic allele. It is conceivable that this might obviate multiple therapies that target events downstream from synthesis of the offending protein. Second, there is a growing interest in the use of neural stem cells in neurodegenerative diseases. These cells have a remarkable capacity to migrate within the brain, and an extraordinary ability to differentiate into diverse types of neural tissue cells. It is conceivable that such cells might migrate to sites of dying motor neurons and provide trophic support by becoming new populations of normal astrocytes, avidly taking up glutamate or secreting neurotrophic substances. It is also possible that these cells might even differentiate into motor neurons and thereby have at least the potential of reinnervating denervated muscle. At all events, it is now clear that a variety of potential therapeutic strategies exist to treat both inherited and sporadic ALS. The existence of a mouse model for this disease will undoubtedly assist greatly in testing these therapeutic options.

Acknowledgement This work was supported by the National Institutes of Health, the ALS Association, the Muscular Dystrophy Association, the Pierre L. de Bourgknecht ALS Research Association, and Project ALS. Dr. Brown’s laboratory receives support from the C.B. Day Foundation.

References 1 2

Mulder DW, Kurland LT, Offord KP, Beard CM: Familial adult motor neuron disease: Amyotrophic lateral sclerosis. Neurology 1986;36:511–517. Siddique T, Figlewicz DA, Pericak-Vance MA, Haines JL, Rouleau G, Jeffers AJ, Sapp P, Hung W-Y, Bebout J, McKenna-Yasek D, Deng G, Robert Horvitz H, Gusella JF, Brown RH Jr, Roses AD: Linkage of a gene causing familial amyotrophic lateral sclerosis to chromosome 21 and evidence of genetic-locus heterogeneity. N Engl J Med 1991;324:1381–1384.

Morita/Brown

184

3

4

5 6

7

8

9

10

11

12

13

14

15

16

17 18 19

Rosen DR, Siddique T, Patterson D, Figlewicz DA, Sapp P, Hentati A, Donaldson D, Goto J, O’Regan JP, Deng HX, Rahmani Z, Krizus A, McKenna-Yasek D, Cayabyab A, Gaston SM, Berger R, Tanzi RE, Halperin JJ, Herzfeldt B, Van den Bergh R, Hung W-Y, Bird T, Deng G, Mulder DW, Smyth C, Laing NG, Soriano E, Pericak-Vance MA, Haines J, Rouleau GA, Gusella JS, Horvitz HR, Brown RH Jr: Mutations in Cu/Zn superoxide dismutase gene are associated with familial amyotrophic lateral sclerosis. Nature 1993;362:59–62. Deng H-X, Hentati A, Tainer JA, Iqbal Z, Cayabyab A, Hung WY, Getzoff ED, Hu P, Herzfeldt B, Roos RP, Warner C, Deng G, Soriano E, Smyth C, Parge HE, Ahmed A, Roses AD, Hallewell RA, Pericak-Vance MA, Siddique T: Amyotrophic lateral sclerosis and structural defects in Cu,Zn superoxide dismutase. Science 1993;261:1047–1051. Brown RH Jr: Amyotrophic lateral sclerosis: Recent insights from genetics and transgenic mice. Cell 1995;80:687–692. Andersen PM, Morita M, Brown RH Jr: Genetics of amyotrophic lateral sclerosis: An overview; in Brown RH Jr, Meininger V, Swash M (eds): Amyotrophic Lateral Sclerosis. London, Dunitz, 1999, pp 223–250. Gurney ME, Pu H, Chiu AY, Dal Canto MC, Polchow CY, Alexander DD, Caliendo J, Hentati A, Kwon YW, Deng H-X, Chen W, Zhai P, Sufit RL, Siddique T: Motor neuron degeneration in mice that express a human Cu,Zn superoxide dismutase mutation. Science 1994;264:1772–1775. Wong PC, Pardo CA, Borchelt DR, Lee MK, Copeland NG, Jenkins NA, Sisodia SS, Cleveland DW, Price DL: An adverse property of a familial ALS-linked SOD1 mutation causes motor neuron disease characterized by vacuolar degeneration of mitochondria. Neuron 1995;14:1105–1116. Ripps ME, Huntley GW, Hof PR, Morrison JH, Gordon JW. Transgenic mice expressing an altered murine superoxide dismutase gene provide an animal model of amyotrophic lateral sclerosis. Proc Natl Acad Sci USA 1995;92:689–693. Bruijn LI, Becher MW, Lee MK, Anderson KL, Jenkins NA, Copeland NG, Sisodia SS, Rothstein JD, Borchelt DR, Price DL, Cleveland DW: ALS-linked SOD1 mutant G85R mediates damage to astrocytes and promotes rapidly progressive disease with SOD1-containing inclusions. Neuron 1997; 18:327–338. Ben Hamida M, Hentati F, Ben Hamida C: Hereditary motor system diseases (chronic juvenile amyotrophic lateral sclerosis). Conditions combining a bilateral pyramidal syndrome with limb and bulbar amyotrophy. Brain 1990;113:347–363. Hentati A, Ouahchi K, Pericak-Vance MA, Nijhawan D, Ahmad A, Yang Y, Rimmler J, Hung W-Y, Schlotter B, Ahmed A, Ben Hamida M, Hentati F, Siddique T: Linkage of a commoner form of recessive amyotrophic lateral sclerosis to chromosome 15q15-q22 markers. Neurogenetics 1998;2: 55–60. Hentati A, Bejaoui K, Pericak-Vance MA, Hentati F, Speer MC, Hung W-Y, Figlewicz DA, Haines J, Rimmler J, Ben Hamida C, Ben Hamida M, Brown RH Jr, Siddique T: Linkage of recessive familial amyotrophic lateral sclerosis to chromosome 2q33-q35. Nat Genet 1994;7:425–428. Hosler BA, Sapp PC, Berger R, O’Neill G, Bejaoui K, Hamida MB, Hentati F, Chin W, McKennaYasek D, Haines JL, Patterson D, Horvitz HR, Brown RH Jr, Day CB: Refined mapping and characterization of the recessive familial amyotrophic lateral sclerosis locus (ALS2) on chromosome 2q33. Neurogenetics 1998;2:34–42. Cudkowicz ME, McKenna-Yasek D, Sapp PE, Chin W, Geller B, Hayden DL, Schoenfeld DA, Hosler BA, Horvitz HR, Brown RH Jr: Epidemiology of mutations in superoxide dismutase in amyotrophic lateral sclerosis. Ann Neurol 1997;41:210–221. Chance PF, Rabin BA, Ryan SG, Ding Y, Scavina M, Crain B, Griffin JW, Cornblath DR: Linkage of the gene for an autosomal dominant form of juvenile amyotrophic lateral sclerosis to chromosome 9q34. Am J Hum Genet 1998;62:633–640. Rabin BA, Griffin JW, Crain BJ, Scavina M, Chance PF, Cornblath DR: Autosomal dominant juvenile amyotrophic lateral sclerosis. Brain 1999;122:1539–1550. Siddique T, Hong S, Brooks BR, Hung W-Y, Siddique NA, Rimmler J, Kaplan JP, Haines JL, Brown RH Jr, Pericak-Vance MA: X-linked dominant ALS (abstract). Neurology 1998;51:310. Forman HJ, Fridovich I: On the stability of bovine superoxide dismutase. The effects of metals. J Biol Chem 1973;248:2645–2649.


185

20

21

22

23

24

25 26 27

28

29

30

31

32 33

34

35 36

37

Pardo CA, Xu Z, Borchelt DR, Price DL, Sisodia SS, Cleveland DW: Superoxide dismutase is an abundant component in cell bodies, dendrites, and axons of motor neurons and in a subset of other neurons. Proc Natl Acad Sci USA 1995;92:954–958. Bowling AC, Barkowski EE, McKenna-Yasek D, Sapp P, Horvitz HR, Beal MF, Brown RH Jr: Superoxide dismutase concentration and activity in familial amyotrophic lateral sclerosis. J Neurochem 1995;64:2366–2369. Borchelt DR, Lee MK, Slunt HS, Guarnieri M, Xu ZS, Wong PC, Brown RH Jr, Price DL, Sisodia SS, Cleveland DW: Superoxide dismutase 1 with mutations linked to familial amyotrophic lateral sclerosis possesses significant activity. Proc Natl Acad Sci USA 1994;91:8292–8296. Andersen PM, Nilsson P, Ala-Hurula V, Kera¨nen M-L, Tarvainen I, Haltia T, Nilsson L, Binzer M, Forsgren L, Marklund SL: Amyotrophic lateral sclerosis associated with homozygosity for an Asp90Ala mutation in CuZn-superoxide dismutase. Nat Genet 1995;10:61–66. Reaume AG, Elliott JL, Hoffman EK, Kowall NW, Ferrante RJ, Siwek DF, Wilcox HM, Flood DG, Beal MF, Brown RH Jr, Scott RW, Snider WD: Motor neurons in Cu/Zn superoxide dismutasedeficient mice develop normally but exhibit enhanced cell death after axonal injury. Nat Genet 1996;13:43–47. Li TM, Alberman E, Swash M: Comparison of sporadic and familial disease amongst 580 cases of motor neuron disease. J Neurol Neurosurg Psychiatry 1988;51:778–784. Strong MJ, Hudson AJ, Alvord WG: Familial amyotrophic lateral sclerosis, 1850–1989: A statistical analysis of the world literature. Can J Neurol Sci 1991;18:45–58. Radunovic A, Leigh PN: Cu/Zn superoxide dismutase gene mutations in amyotrophic lateral sclerosis: Correlation between genotype and clinical features. J Neurol Neurosurg Psychiatry 1996;61: 565–572. Juneja T, Pericak-Vance MA, Laing NG, Dave S, Siddique T: Prognosis in familial amyotrophic lateral sclerosis: Progression and survival in patients with glu100gly and ala4val mutations in Cu,Zn superoxide dismutase. Neurology 1997;48:55–57. Cudkowicz ME, McKenna-Yasek D, Chen C, Hedley WE, Brown RH Jr: Limited corticospinal tract involvement in amyotrophic lateral sclerosis subjects with the A4V mutation in the copper/ zinc superoxide dismutase gene. Ann Neurol 1998;43:703–710. Shibata N, Hirano A, Kobayashi M, Siddique T, Deng HX, Hung WY, Kato T, Asayama K: Intense superoxide dismutase-1 immunoreactivity in intracytoplasmic hyaline inclusions of familial amyotrophic lateral sclerosis with posterior column involvement. J Neuropathol Exp Neurol 1996; 55:481–490. Boukaftane Y, Khoris J, Moulard B, Salachas F, Meininger V, Malafosse A, Camu W, Rouleau GA: Identification of six novel SOD1 gene mutations in familial amyotrophic lateral sclerosis. Can J Neurol Sci 1998;25:192–196. Aoki M, Ogasawara M, Matsubara Y, Narisawa K, Nakamura S, Itoyama Y, Abe K: Mild ALS in Japan associated with novel SOD mutation. Nat Genet 1993;5:323–324. Andersen PM, Forsgren L, Binzer M, Nilsson P, Ala HV, Kera¨nen M-L, Bergmark L, Saarinen A, Haltia T, Tarvainen I, Kinnunen E, Udd B, Marklund SL: Autosomal recessive adult-onset amyotrophic lateral sclerosis associated with homozygosity for Asp90A1a CuZn-superoxide dismutase mutation. A clinical and genealogical study of 36 patients. Brain 1996;119: 1153–1172. Sapp PC, Rosen DR, Hosler BA, Esteban J, McKenna-Yasek D, O’Regan JP, Horvitz HR, Brown RH Jr: Identification of three novel mutations in the gene for Cu/Zn superoxide dismutase in patients with familial amyotrophic lateral sclerosis. Neuromusc Disord 1995;5:353–357. Kostrzewa M, Damian MS, Mu¨ller U: Superoxide dismutase 1: Identification of a novel mutation in a case of familial amyotrophic lateral sclerosis. Hum Genet 1996;98:48–50. Ratovitski T, Corson LB, Strain J, Wong P, Cleveland DW, Culotta VC, Borchelt DR: Variation in the biochemical/biophysical properties of mutant superoxide dismutase 1 enzymes and the rate of disease progression in familial amyotrophic lateral sclerosis kindreds. Hum Mol Genet 1999;8: 1451–1460. Beckman JS, Carson M, Smith CD, Koppenol WH: ALS, SOD and peroxynitrite. Nature 1993; 364:584.

Morita/Brown

186

38

39 40

41

42

43 44

45

46

47

48 49

50

51

52

53

54

55

Bruijn LI, Beal MF, Becher MW, Schulz JB, Wong PC, Price DL, Cleveland DW: Elevated free nitrotyrosine levels, but not protein-bound nitrotyrosine or hydroxyl radicals, throughout amyotrophic lateral sclerosis (ALS)-like disease implicate tyrosine nitration as an aberrant in vivo property of one familial ALS-linked superoxide dismutase 1 mutant. Proc Natl Acad Sci USA 1997;94: 7606–7611. Beal MF, Ferrante RJ, Browne SE, Matthews RT, Kowall NW, Brown RH Jr: Increased 3-nitrotyrosine in both sporadic and familial amyotrophic lateral sclerosis. Ann Neurol 1997;42:644–654. Wiedau-Pazos M, Goto JJ, Rabizadeh S, Gralla EB, Roe JA, Lee MK, Valentine JS, Bredesen DE: Altered reactivity of superoxide dismutase in familial amyotrophic lateral sclerosis. Science 1996; 271:515–518. Yim MB, Kang JH, Yim HS, Kwak HS, Chock PB, Stadtman ER: A gain-of-function of an amyotrophic lateral sclerosis-associated Cu,Zn-superoxide dismutase mutant: An enhancement of free radical formation due to a decrease in Km for hydrogen peroxide. Proc Natl Acad Sci USA 1996;93:5709–5714. Bruijn LI, Houseweart MK, Kato S, Anderson KL, Anderson SD, Ohama E, Reaume AG, Scott RW, Cleveland DW: Aggregation and motor neuron toxicity of an ALS-linked SOD1 mutant independent from wild-type SOD1. Science 1998;281:1851–1854. Carpenter S: Proximal axonal enlargement in motor neuron disease. Neurology 1968;18:841–851. Munoz DG, Greene C, Perl DP, Selkoe DJ: Accumulation of phosphorylated neurofilaments in anterior horn motoneurons of amyotrophic lateral sclerosis patients. J Neuropathol Exp Neurol 1988;47:9–18. Manetto V, Sternberger NH, Perry G, Sternberger LA, Gambetti P: Phosphorylation of neurofilaments is altered in amyotrophic lateral sclerosis. J Neuropathol Exp Neurol 1988;47:642– 653. Hirano A, Nakano I, Kurland LT, Mulder DW, Holley PW, Saccomanno G: Fine structural study of neurofibrillary changes in a family with amyotrophic lateral sclerosis. J Neuropathol Exp Neurol 1984;43:471–480. Xu Z, Cork LC, Griffin JW, Cleveland DW: Increased expression of neurofilament subunit NF-L produces morphological alterations that resemble the pathology of human motor neuron disease. Cell 1993;73:23–33. Coˆte´ F, Collard J-F, Julien J-P: Progressive neuronopathy in transgenic mice expressing the human neurofilament heavy gene: A mouse model of amyotrophic lateral sclerosis. Cell 1993;73:35–46. Lee MK, Marszalek JR, Cleveland DW: A mutant neurofilament subunit causes massive, selective motor neuron death: Implications for the pathogenesis of human motor neuron disease. Neuron 1994;13:975–988. Figlewicz DA, Krizus A, Martinoli MG, Meininger V, Dib M, Rouleau GA, Julien J-P: Variants of the heavy neurofilament subunit are associated with the development of amyotrophic lateral sclerosis. Hum Mol Genet 1994;3:1757–1761. Tomkins J, Usher P, Slade JY, Ince PG, Curtis A, Bushby K, Shaw PJ: Novel insertion in the KSP region of the neurofilament heavy gene in amyotrophic lateral sclerosis (ALS). Neuroreport 1998; 9:3967–3970. Al-Chalabi A, Andersen PM, Nilsson P, Chioza B, Andersson JL, Russ C, Shaw CE, Powell JF, Leigh PN: Deletions of the heavy neurofilament subunit tail in amyotrophic lateral sclerosis. Hum Mol Genet 1999;8:157–164. Rosen DR, Bowling AC, Patterson D, Usdin TB, Sapp P, Mezey E, McKenna-Yasek D, O’Regan J, Rahmani Z, Ferrante RJ, Brownstein MJ, Kowall NW, Beal MF, Horvitz HR, Brown RH Jr: A frequent ala 4 to val superoxide dismutase-1 mutation is associated with a rapidly progressive familial amyotrophic lateral sclerosis. Hum Mol Genet 1994;3:981–987. Rouleau GA, Clark AW, Rooke K, Pramatarova A, Krizus A, Suchowersky O, Julien J-P, Figlewicz D: SOD1 mutation is associated with accumulation of neurofilaments in amyotrophic lateral sclerosis. Ann Neurol 1996;39:128–131. Shaw CE, Enayat ZE, Powell JF, Anderson VER, Radunovic A, Al-Sarraj S, Leigh PN: Familial amyotrophic lateral sclerosis. Molecular pathology of a patient with a SOD1 mutation. Neurology 1997;49:1612–1616.


187

56

57

58

59

60

61

62 63

64 65

66 67

68

69 70

71 72

73 74

Tu P-H, Raju P, Robinson KA, Gurney ME, Trojanowski JQ, Lee VM-Y: Transgenic mice carrying a human mutant superoxide dismutase transgene develop neuronal cytoskeletal pathology resembling human amyotrophic lateral sclerosis lesions. Proc Natl Acad Sci USA 1996;93:3155–3160. Couillard-Despre´s S, Zhu Q, Wong PC, Price DL, Cleveland DW, Julien J-P: Protective effect of neurofilament heavy gene overexpression in motor neuron disease induced by mutant superoxide dismutase. Proc Natl Acad Sci USA 1998;95:9626–9630. Williamson TL, Bruijn LI, Zhu Q, Anderson KL, Anderson SD, Julien J-P, Cleveland DW: Absence of neurofilaments reduces the selective vulnerability of motor neurons and slows disease caused by a familial amyotrophic lateral sclerosis-linked superoxide dismutase 1 mutant. Proc Natl Acad Sci USA 1998;95:9631–9636. Marszalek JR, Williamson TL, Lee MK, Xu Z, Hoffman PN, Becher MW, Crawford TO, Cleveland DW: Neurofilament subunit NF-H modulates axonal diameter by selectively slowing neurofilament transport. J Cell Biol 1996;135:711–724. Zhang B, Tu PN, Abtahian F, Trojanowski JQ, Lee VM-Y: Neurofilaments and orthograde transport are reduced in ventral root axons of transgenic mice that express human SOD1 with a G93A mutation. J Cell Biol 1997;139:1307–1315. Borchelt DR, Wong PC, Becher MW, Pardo CA, Lee MK, Xu Z-S, Thinakaran G, Jenkins NA, Copeland NG, Sisodia SS, Cleveland DW, Price DL, Hoffman PN: Axonal transport of mutant superoxide dismutase 1 and focal axonal abnormalities in the proximal axons of transgenic mice. Neurobiol Dis 1998;5:27–35. Williamson TL, Cleveland DW: Slowing of axonal transport is a very early event in the toxicity of ALS-linked SOD1 mutants to motor neurons. Nat Neurosci 1999;2:50–56. Warita H, Itoyama Y, Abe K: Selective impairment of fast anterograde axonal transport in the peripheral nerves of asymptomatic transgenic mice with a G93A mutant SOD1 gene. Brain Res 1999;819:120–131. Choi DW: Glutamate neurotoxicity and diseases of the nervous system. Neuron 1988;1:623–634. Tanaka K, Watase K, Manabe T, Yamada K, Watanabe M, Takahashi K, Iwama H, Nishikawa T, Ichihara N, Kikuchi T, Okuyama S, Kawashima N, Hori S, Takimoto M, Wada K: Epilepsy and exacerbation of brain injury in mice lacking the glutamate transporter GLT-1. Science 1997; 276:1699–1702. Rothstein JD, Van Kammen M, Levey AI, Martin LJ, Kuncl RW: Selective loss of glial glutamate transporter GLT-1 in amyotrophic lateral sclerosis. Ann Neurol 1995;38:73–84. Shaw PJ, Forrest V, Ince PG, Richardson JP, Wastell HJ: CSF and plasma amino acid levels in motor neuron disease: Elevation of CSF glutamate in a subset of patients. Neurodegeneration 1995; 4:209–216. Lin C-LG, Bristol LA, Jin L, Dykes-Hoberg M, Crawford T, Clawson L, Rothstein JD: Aberrant RNA processing in a neurodegenerative disease: The cause for absent EAAT2, a glutamate transporter, in amyotrophic lateral sclerosis. Neuron 1998;20:589–602. Nagai M, Abe K, Okamoto K, Itoyama Y: Identification of alternative splicing forms of GLT-1 mRNA in the spinal cord of amyotrophic lateral sclerosis patients. Neurosci Lett 1998;244:165–168. Meyer T, Mu¨nch C, Liebau S, Fromm A, Schwalensto¨cker B, Vo¨lkel H, Ludolph AC: Splicing of the glutamate transporter EAAT2: A candidate gene of amyotrophic lateral sclerosis. J Neurol Neurosurg Psychiatry 1998;65:954. Trotti D, Rolfs A, Danbolt NC, Brown RH Jr, Hediger MA: SOD1 mutants linked to amyotrophic lateral sclerosis selectively inactivate a glial glutamate transporter. Nat Neurosci 1999;2:427–433. Carrı` MT, Ferri A, Battistoni A, Famhy L, Gabbianelli R, Poccia F, Rotilio G: Expression of a Cu,Zn superoxide dismutase typical of familial amyotrophic lateral sclerosis induces mitochondrial alteration and increase of cytosolic Ca2+ concentration in transfected neuroblastoma SH-SY5Y cells. FEBS Lett 1997;414:365–368. Siklos L, Engelhardt J, Harati Y, Smith RG, Joo F, Appel SH: Ultrastructural evidence for altered calcium in motor nerve terminals in amyotrophic lateral sclerosis. Ann Neurol 1996;39:203–216. Siklos L, Engelhardt JI, Alexianu ME, Gurney ME, Siddique T, Appel SH: Intracellular calcium parallels motoneuron degeneration in SOD-1 mutant mice. J Neuropathol Exp Neurol 1998;57: 571–587.

Morita/Brown

188

75 76 77

78

79 80

81

82 83

84

85 86 87 88

89

Fitzmaurice PS, Shaw IC, Kleiner HE, Miller RT, Monks TJ, Lau SS, Mitchell JD, Lynch PG: Evidence for DNA damage in amyotrophic lateral sclerosis. Muscle Nerve 1996;19:797–798. Shaw PJ, Ince PG, Falkous G, Mantle D: Oxidative damage to protein in sporadic motor neuron disease spinal cord. Ann Neurol 1995;38:691–695. Bowling AC, Schulz JB, Brown RH Jr, Beal MF: Superoxide dismutase activity, oxidative damage, and mitochondrial energy metabolism in familial and sporadic amyotrophic lateral sclerosis. J Neurochem 1993;61:2322–2325. Ghadge GD, Lee JP, Bindokas VP, Jordan J, Ma L, Miller RJ, Roos RP: Mutant superoxide dismutase-1-linked familial amyotrophic lateral sclerosis: Molecular mechanisms of neuronal death and protection. J Neurosci 1997;17:8756–8766. Durham HD, Roy J, Dong L, Figlewicz DA: Aggregation of mutant Cu/Zn superoxide dismutase proteins in a culture model of ALS. J Neuropathol Exp Neurol 1997;56:523–530. Pasinelli P, Borchelt DR, Houseweart MK, Cleveland DW, Brown RH Jr: Caspase-1 is activated in neural cells and tissue with amyotrophic lateral sclerosis-associated mutations in copper-zinc superoxide dismutase. Proc Natl Acad Sci USA 1998;95:15763–15768. Gurney ME, Cutting FB, Zhai P, Doble A, Taylor CP, Andrus PK, Hall ED: Benefit of vitamin E, riluzole, and gabapentin in a transgenic model of familial amyotrophic lateral sclerosis. Ann Neurol 1996;39:147–157. Desnuelles C: International Motor Neuron Disease Meeting, Munich, 1998. Hottinger AF, Fine EG, Gurney ME, Zurn AD, Aebischer P: The copper chelator d-penicillamine delays onset of disease and extends survival in a transgenic mouse model of familial amyotrophic lateral sclerosis. Eur J Neurosci 1997;9:1548–1551. Klivenyi P, Ferrante RJ, Matthews RT, Bogdanov MB, Klein AM, Andreassen OA, Mueller G, Wermer M, Kaddurah-Daouk R, Beal MF: Neuroprotective effects of creatine in a transgenic animal model of amyotrophic lateral sclerosis. Nat Med 1999;5:347–350. Friedlander RM, Brown RH Jr, Gagliardini V, Wang J, Yuan J: Inhibition of ICE slows ALS in mice. Nature 1997;388:31. Kostic V, Jackson-Lewis V, de Bilbao F, Dubois-Dauphin M, Przedborski S: Bcl-2: Prolonging life in a transgenic mouse model of familial amyotrophic lateral sclerosis. Science 1997;277:559–562. Nagy D, Kato T, Kushner PD: Reactive astrocytes are widespread in the cortical gray matter of amyotrophic lateral sclerosis. J Neurosci Res 1994;38:336–347. Migheli A, Piva R, Atzori C, Troost D, Schiffer D: c-Jun, JNK/SAPK kinases and transcription factor NF-kappa B are selectively activated in astrocytes, but not motor neurons, in amyotrophic lateral sclerosis. J Neuropathol Exp Neurol 1997;56:1314–1322. Kato S, Saito M, Hirano A, Ohama E: Recent advances in research on neuropathological aspects of familial amyotrophic lateral sclerosis with superoxide dismutase 1 gene mutations: Neuronal Lewy body-like hyaline inclusions and astrocytic hyaline inclusions. Histol Histopathol 1999;14: 973–989.

Robert H. Brown, Jr., MD, Day Neuromuscular Research Center, MGH, East, Room 6627, Building 149, 13th Street, Navy Yard, Charlestown, MA 02129 (USA) Tel. +1 617 726 5750, Fax +1 617 726 8543, E-Mail [email protected]


189

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

Author Index

Bo¨nnemann, C.G. 26 Bella, I.R. 147 Brown, R.H. Jr 177 Buyse, G.M. 1

Meola, G. 61 Morita, M. 177 Moxley, R.T. 61 Nelis, E. 128

De Jonghe, P. 128 Deymeer, F. 113

Ohno, K. 96 ¨ zdemir, C. 113 O

Engel, A.G. 96 Ru¨del, R. 79 Hausmanowa-Petrusewicz, I. 163 Hoffman, E.P. 1

Serdarog˘lu, P. 113 Skuk, D. 12 Stans, A.A. 96

Jurkat-Rott, K. 79 Lehmann-Horn, F. 79 Lunt, P.W. 44

Timmerman, V. 128 Tremblay, J.P. 12 Zaremba, J. 163

190

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

Subject Index

Acetylcholine receptor antibodies in myasthenia gravis juvenile disease 118 late-onset disease 122, 123 mutations, see Congenital myasthenic syndromes Acetylcholinesterase deficiency, see Congenital myasthenic syndromes Acute inflammatory demyelinating polyradiculoneuropathy clinical features 147–149 differential diagnosis 155, 156 epidemiology 147 infection role in onset 149, 150 laboratory findings antiglycoconjugate autoantibodies 151, 152 blood analysis 150, 151 cerebrospinal fluid analysis 151 electrodiagnostic studies 152, 159 management complications 155 intravenous immunoglobulin 158 overview 155, 157 plasmapheresis 155, 157, 158 molecular mimicry 154 nomenclature 147, 149 outcomes 158, 159 pathogenesis 152–154 pathology 154, 155 Acute motor axonal neuropathy antiglycoconjugate autoantibodies 152

classification 147, 149 clinical features 149 electrodiagnostic studies 152, 159 infection role in onset 149, 150 pathogenesis 153, 154 Acute motor sensory axonal neuropathy classification 147, 149 clinical features 149 electrodiagnostic studies 152, 159 infection role in onset 149, 150 pathogenesis 153, 154 Adeno-associated virus, gene therapy vector 4–8 Adenovirus, gene therapy vector 4–6 Amyotrophic lateral sclerosis epidemiology 177 hereditary form, see Familial amyotrophic lateral sclerosis Anticipation facioscapulohumeral muscular dystrophy 48 myotonic dystrophy 64, 65 Azathioprine, myasthenia gravis treatment 124 Basal lamina onion bulbs, Charcot-Marie-Tooth disease 135 Campylobacter jejuni, infection role in Guillain-Barre´ syndrome 149, 150, 154 Carbamazepine, myotonic dystrophy trials 74

191

Central core disease clinical features 89, 90 genetics 90 heredity 8 Charcot-Marie-Tooth disease classification 128–131 clinical phenotypes 129 CMT1 autosomal dominant CMT1A 132, 133 CMT1B 133, 134 EGR2 mutations 134 overview 131, 132 autosomal recessive basal lamina onion bulbs 135 CMT4A 135 CMT4B 135 HMSN-L 136 overview 135 X-linked 134, 135 CMT2 autosomal dominant CMT2A 137 CMT2B 137 CMT2D 137 loci 136, 137 autosomal recessive 137 overview 136 X-linked 137 gene mutations, overview 128, 130–132 CLC-1 channelopathy 90, 91, 93, 94 structure and function 94 ColQ, structure and mutations 101, 102 Congenital hypomyelination clinical phenotype 129 gene mutations 130 Congenital myasthenic syndromes candidate genes 99 classification 96, 97 diagnosis clinical history 97 differential diagnosis 98 electromyography 98 molecular analysis 99 physical examination 97, 98 investigation 98, 99, 108

Subject Index

overview 96 postsynaptic syndromes fast-channel syndrome gating abnormality 105 low-affinity fast-channel syndrome 105 mode-switching kinetics 105, 106 overview 105 mutations causing acetylcholine receptor deficiency with or without minor kinetic abnormalities 106–109 slow-channel syndrome gene mutations 102, 104 pathology 102 quinidine treatment 104 presynaptic syndromes episodic apnea 99, 100 heredity 108 paucity of synaptic vesicles and reduced quantal release 99 resembling Lambert-Eaton myasthenic syndrome 100 synaptic acetylcholinesterase deficiency pathology 100, 101 structure and mutations 101, 102, 108 Creatine, amyotrophic lateral sclerosis treatment 183 Creatinine, muscle strength enhancement 4 Cytomegalovirus, infection role in Guillain-Barre´ syndrome 150 Dehydroepiandrosterone sulfate, myotonic dystrophy trials 73 De´jerine-Sottas syndrome classification 136 clinical phenotype 129 gene mutations 130, 136 Dihydropyridine receptor channelopathies 85, 86, 89 functions 85 Disopyramide, myotonic dystrophy trials 74 Distal hereditary motor neuropathies clinical presentation 137, 138 congenital nonprogressive disease 138

192

recessive disease 139 type II 138 type IV 138 Drug screening, animal models 3, 4 Duchenne muscular dystrophy animal models 1–3 myoblast transplantation, see Myoblast transplantation Dystroglycan dystrophin binding 27, 28 gene 27 sarcoglycan complex 30 Dystrophin deficiency animal models 2, 3 expression in sarcoglycanopathy 33 neuronal nitric oxide synthase localization role 2 Dystrophin-associated proteins dystrophin complexes 26, 27 sarcoglycan complex 29, 30 types 27, 28 EAAT2, mutant SOD1 effects 181, 182 EGR2, mutations 134 Facioscapulohumeral muscular dystrophy anticipation 48 childhood testing 56 course 44, 47 diagnosis index case 46–48 linkage analysis 52, 53, 56 molecular testing distribution of fragment size and type 49, 50 false-positives 51 sensitivity and specificity 51, 52 family studies 54, 55 probes 48, 49, 51 pulsed-field gel electrophoresis 49–51 epidemiology 44 family history 48 gene mutation 44, 45 management 53, 54 molecular pathogenesis position effects 45, 56, 57

Subject Index

regulator role of repeats 57, 58 repeat length effects 46, 52 penetrance and risk to offspring 55 prenatal testing 56 prognosis 53 prospects for research 58 Familial amyotrophic lateral sclerosis clinical features 177 juvenile forms 178 linkage analysis 177, 178 SOD1 mutations glutamate transporter effects 181, 182 mitochondrial dysfunction 182 neurotoxicity of mutant proteins 180–182 phenotypes 179, 180 structure and function 178, 179 types 177, 178 therapeutic trials combination therapy 184 copper chelators 183 creatine 183 growth factors 183 prospects 184 vitamin E 182, 183 X-linked dominant form 178 FK506, immunosuppression for myoblast transplantation 15, 18, 19 Gene therapy myoblast modification 15, 16 sarcoglycanopathy 37 vectors adeno-associated virus 4–8 adenovirus 4–6 Glutamine, muscle strength enhancement 4 Growth hormone, myotonic dystrophy trials 72 Guillain-Barre´ syndrome axonal variants, see Acute motor axonal neuropathy, Acute motor sensory axonal neuropathy demyelinating variant, see Acute inflammatory demyelinating polyradiculoneuropathy

193

Hereditary neuralgic amyotrophy clinical features 140, 141 linkage studies 141 Hereditary neuropathy with liability to pressure palsies clinical features 139, 140 gene defects 140 Hereditary peripheral neuropathies, see also Charcot-Marie-Tooth disease, Congenital hypomyelination, De´jerine-Sottas syndrome, Distal hereditary motor neuropathies classification 128–131 clinical phenotypes 129 gene mutations 128, 130, 131 hereditary recurrent neuropathies 139–141 hereditary sensory neuropathies 139 Hyperkalemic periodic paralysis clinical features 80, 81 electrophysiological basis 81, 82 gene mutation 79 pathogenesis of symptoms 84, 85 Hypokalemic periodic paralysis electrophysiology 86 gene defect 86 pathogenesis 86, 87 Influenza vaccine, role in Guillain-Barre´ syndrome 150 Insulin, myotonic dystrophy trials 73 Insulin-like growth factor-1 muscle strength enhancement 4 myotonic dystrophy trials 73 Intravenous immunoglobulin, Guillain-Barre´ syndrome management 158 Limb-girdle muscular dystrophy, see Sarcoglycanopathy Malignant hyperthermia susceptibility gene mutations 88, 89 management 87, 88 overview 87 Mexiletine, myotonic dystrophy trials 74

Subject Index

Myasthenia gravis age of onset 113 juvenile disease anti-acetylcholine receptor antibodies 118 autoimmune disease association 118 definition 113 demographic features 115, 116 onset symptoms ocular symptoms 116, 117 prepuberty 117, 120 puberty and postpuberty 117, 118, 120 racial differences 116, 120 severity and outcome prepuberty 119 puberty and postpuberty 120 study design 118, 119 studies for analysis 114, 115 thymoma 118 late-onset disease anti-acetylcholine receptor antibodies 122, 123 autoimmune disease association 123 azathioprine treatment 124 definition 120, 121 demographic features 121 onset symptoms 121, 124 severity and outcome 123, 124 thymoma 123 Myoblast transplantation animal studies dog 18 monkey 19 mouse 13, 19 efficacy congenital dystrophy 20 Duchenne muscular dystrophy benefits 19, 20 early clinical trials 13, 14 metabolic myopathy 20 genetically modified myoblasts 15, 16 immune response 14, 15 immunosuppression therapy 15 migration of cells 17, 18 rationale 12, 13 survival factors 17

194

MyoD, deficiency animal models 2, 3 Myotonia congenita molecular pathology 91, 93, 94 transfected cell studies of channel mutants 91, 93 types 90, 91 Myotonic dystrophy animal models 74, 75 anticipation 64, 65 cardiac manifestations 67 clinical features 61, 62, 66, 68 diagnosis 61–63 DMPK deficiency 61, 68 genetic counseling 63 management and trials antimyotonia therapy 73, 74 dehydroepiandrosterone 73 growth hormone 72 infancy and childhood 68, 69 insulin 73 insulin-like growth factor-1 73 maternal-obstetric complications 68, 71 skeletal muscle problems 68, 70 surgical management of patients 68, 71 testosterone 68, 72 natural history 68, 72 sex differences 65 trinucleotide expansion developmental changes 67, 68 enlargement 65 locus 61 molecular testing 61–63 mosaicism and clinical presentation 66–68 Neuronal nitric oxide synthase, localization role of dystrophin 2 Oxatomide, muscle strength enhancement 4 Paramyotonia congenita clinical features 80, 81 electrophysiological basis 81, 82 gene mutation 79 pathogenesis of symptoms 84, 85 temperature sensitivity 85

Subject Index

Pentoxifylline, muscle strength enhancement 4 Phenytoin, myotonic dystrophy trials 74 Plasmapheresis, Guillain-Barre´ syndrome management 155, 157, 158 Potassium-aggravated myotonia clinical features 80, 81 electrophysiological basis 81, 82 gene mutation 79 pathogenesis of symptoms 84, 85 Procainamide, myotonic dystrophy trials 74 Quinidine, slow-channel congenital myasthenic syndrome treatment 104 Ryanodine receptor channelopathies 85, 86, 89 functions 85 Sarcoglycan dystrophin complex 29, 30, 37 functions 30 genes 29 structures 29 Sarcoglycanopathy animal models 35, 36 clinical features 31, 32 diagnosis 32, 33 limb-girdle muscular dystrophy classification 26, 27 mutations a-sarcoglycan 33, 34 b-sarcoglycan 34 c-sarcoglycan 34, 35 d-sarcoglycan 35 protein expression patterns 33 treatment 37 types and distribution 30, 31, 33 SCN4A, mutation 79 SMN protein, function and mutations 169, 170 Spinal muscular atrophy, proximal disease of childhood differential diagnosis 171 epidemiology 165 forms and clinical features 164 gene mutations 168–170

195

Spinal muscular atrophy (continued) genetic counseling 172 history of study 163 laboratory findings electromyography 165 histology 166, 167 muscle morphology 165–167 linkage analysis 168 pathophysiology 167, 168 prenatal diagnosis 172, 173 sex differences 170 SMN protein function 169, 170 treatment 171, 172 Superoxide dismutase, mutations in familial amyotrophic lateral sclerosis glutamate transporter effects 181, 182 mitochondrial dysfunction 182

Subject Index

neurotoxicity of mutant proteins 180–182 phenotypes 179, 180 structure and function 178, 179 types 177, 178 Testosterone, myotonic dystrophy trials 68, 72 Thymoma, myasthenia gravis association juvenile disease 118 late-onset disease 123 Tocainide, myotonic dystrophy trials 74 Utrophin, deficiency animal models 2, 3 Vitamin E, amyotrophic lateral sclerosis treatment 182, 183

196

Neuromuscular Diseases

Recommend Documents