myotonic dystrophy type 2 (dm2) : diagnostic methods and
TRANSCRIPT
Myotonic dystrophy type 2 (DM2): Diagnostic methods and molecular pathology
Anna Naukkarinen
Folkhälsan Institute of Genetics Helsinki, Finland
Department of Medical Genetics University of Helsinki Helsinki, Finland
Vaasa Central Hospital
Department of Pathology Vaasa, Finland
ACADEMIC DISSERTATION
To be presented, with the permission of the Faculty of Medicine of the University of Helsinki, for public examination in
lecture hall 3, Biomedicum, Haartmaninkatu 8, Helsinki on April 8th 2011, at 1 PM.
Helsinki 2011
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Supervisor: Professor Bjarne Udd, MD, PhD
Folkhälsan Institute of Genetics Department of Medical Genetics Biomedicum, C319a University of Helsinki PB 63, 00014 Helsinki Neuromuscular Research Unit University of Tampere Department of Neurology Tampere University Hospital 33520 Tampere, Finland Vasa Central Hospital Department of Neurology 65130 Vasa, Finland
Reviewers: Professor Caroline Sewry, PhD
Dubowitz Neuromuscular Centre Institute of Child Health Great Ormond Street Hospital London UK
Associate Professor Katarina Pelin, PhD Division of Genetics Department of Biosciences University of Helsinki Finland
Opponent: Professor Jaakko Ignatius, MD, PhD
Department of Clinical Genetics Turku University Hospital Finland
ISBN 978‐952‐92‐8732‐1 (paperback) ISBN 978‐952‐10‐6894‐2 (PDF) http://ethesis.helsinki.fi Unigrafia Oy Helsinki 2011
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To my family
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TABLE OF CONTENTS
1. LIST OF ORIGINAL PUBLICATIONS ................................................................................... 7
2. ABBREVIATIONS ............................................................................................................ 9
3. ABSTRACT .................................................................................................................... 10
4. REVIEW OF THE LITERATURE ........................................................................................ 12
4.1. Introduction to cell and molecular biology .................................................................. 12
4.1.1. Microsatellite‐DNA ................................................................................................ 13
4.1.2. Mutations .............................................................................................................. 13
4.1.3. Proteins .................................................................................................................. 14
4.2. Molecular methods for research and diagnostics ......................................................... 15
4.2.1. Detecting nucleic acids with hybridization techniques ......................................... 15
4.2.1.1. Southern blotting ........................................................................................ 15
4.2.1.2. In situ –hybridization ................................................................................... 16
4.2.1.3. Microarrays ................................................................................................. 16
4.2.2. Detecting proteins using antibody recognition ..................................................... 17
4.2.2.1. Antibodies ................................................................................................... 17
4.2.2.2. Immunohistochemistry ............................................................................... 19
4.2.2.3. Western blotting ......................................................................................... 21
4.3. The muscle ..................................................................................................................... 21
4.3.1. The muscle structure ............................................................................................. 23
4.3.2. Skeletal myogenesis .............................................................................................. 25
4.3.3. The satellite cells ................................................................................................... 26
4.3.4. Fiber types ............................................................................................................. 27
4.3.4.1. Fiber type distributions in disease .............................................................. 28
4.3.5. The skeletal muscle fiber ....................................................................................... 29
4.3.6. The sarcomere ....................................................................................................... 32
4.3.7. Muscle contraction ................................................................................................ 34
4.3.8. Role of Ca2+ in muscle contraction ........................................................................ 35
4.3.8.1. The ryanodine receptor ............................................................................... 36
4.3.8.2. The SERCA family of Ca2+ re‐uptake channels ............................................. 36
4.3.8.3. The triad and calsequestrin ......................................................................... 37
4.3.9. Role of Ca2+ in muscle signaling ............................................................................. 37
4.3.10. The muscle biopsy ............................................................................................... 39
4.4. The muscular dystrophies ............................................................................................. 40
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4.4.1. Repeat expansions and disease ............................................................................. 41
4.4.2. The myotonic dystrophies DM1 and DM2 ............................................................. 43
4.4.2.1. History and prevalence ............................................................................... 43
4.4.2.2. DM1 and DM2 mutations ............................................................................ 44
4.4.2.3. Symptoms .................................................................................................... 47
4.4.2.4. Muscle histopathology ................................................................................ 48
4.4.2.5. Diagnostics .................................................................................................. 50
4.4.2.6. Pathomechanisms ....................................................................................... 51
4.4.2.6.1. The muscleblind proteins .................................................................. 53
4.4.2.6.2. CUGBP1 .............................................................................................. 55
4.4.2.6.3. Role of DMPK in DM1 ........................................................................ 56
4.4.2.6.4. Role of ZNF9 in DM2 .......................................................................... 57
4.4.2.6.5. The nuclear foci ................................................................................. 58
4.4.2.6.6. Neighboring genes ............................................................................ 58
4.4.2.6.7. Other pathomechanisms .................................................................. 58
4.4.2.7. Animal models ............................................................................................. 59
4.4.2.8. Management and gene therapy .................................................................. 61
5. AIMS OF THE STUDY ..................................................................................................... 63
6. MATERIALS AND METHODS .......................................................................................... 64
6.1. Patients and samples ..................................................................................................... 64
6.1.1. Consent and ethics committee permissions .......................................................... 64
6.1.2. Patients included in the histopathological study (I) ............................................... 65
6.1.3. Patients included in the study of new diagnostic methods (II) .............................. 65
6.1.4. Patients included in the studies of sarcomeric and Ca2+ handling proteins and mRNA expression (III, IV) .................................................................................................. 65
6.1.5. Patients with neuromuscular diseases included in the study of protein expression in the highly atrophic fibers (U) ........................................................................................ 65
6.2. Methods ........................................................................................................................ 66
6.2.1. Immunohistochemistry (I, III, IV), immunofluorescence (III, IV) and microscopy (I‐IV) .................................................................................................................................... 68
6.2.2. Enzyme histochemistry (I) ..................................................................................... 68
6.2.3. Muscle morphometry (I) ........................................................................................ 68
6.2.4. Chromogenic in situ –hybridization (II) .................................................................. 69
6.2.5. Fiber‐FISH (II) .......................................................................................................... 69
6.2.6. PCR‐based allele sizing (II) ...................................................................................... 70
6.2.7. Repeat‐primed PCR (RP‐PCR) (II) ............................................................................ 70
6.2.8. Field‐inversion gel electrophoresis (FIGE) (II) ........................................................ 71
6.2.9. Western blotting (III, IV) ......................................................................................... 71
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6.2.10. Microarray expression profiling (III, IV) ................................................................ 71
6.2.11. Splice variant analysis (III, IV) ............................................................................... 72
7. RESULTS AND DISCUSSION ........................................................................................... 73
7.1. Histopathology and protein expression in DM2 (I, III, IV, U) ......................................... 73
7.1.1. The atrophy of type II fibers in DM2 (I) .................................................................. 76
7.1.2. Protein expression in highly atrophic fibers (III, U) ................................................ 78
7.1.3. Using fiber type II atrophy as a predictive factor for DM2 diagnosis .................... 84
7.1.4. Satellite cells (U) ..................................................................................................... 84
7.1.5. Expression of Ca2+ ‐handling genes in DM (IV) ....................................................... 86
7.2. Molecular genetic tools for DM2 mutation identification and size determination (II) . 88
7.2.1. PCR‐based allele sizing (II) ...................................................................................... 88
7.2.2. RP‐PCR and FIGE (II) ................................................................................................ 88
7.2.3. In situ –hybridization (II) ......................................................................................... 88
7.2.3.1. CISH (II) ......................................................................................................... 89
7.2.3.2. Fiber‐FISH (II) ................................................................................................ 92
7.3. Splice variant analysis (III, IV) ........................................................................................ 93
7.4. Expression profiling (III, IV) ............................................................................................ 96
7.4.1. Muscle‐specific gene array (III) .............................................................................. 97
7.4.2. Ca2+ gene array (IV) ................................................................................................. 99
8. CONCLUDING REMARKS AND FUTURE PROSPECTS ..................................................... 101
9. ACKNOWLEDGEMENTS .............................................................................................. 104
10. REFERENCES ............................................................................................................. 107
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1. LIST OF ORIGINAL PUBLICATIONS This thesis is based on the following original publications, which are referred to in the text by their roman numerals. In addition, some unpublished results (U) are presented. I Vihola A*, Bassez G*, Meola G, Zhang S, Haapasalo H, Paetau A, Mancinelli E,
Rouche A, Hogrel J Y, Laforêt P, Maisonobe T, Pellissier J F, Krahe R, Eymard B, Udd B. Histopathological differences of myotonic dystrophy type 1 (DM1) and PROMM/DM2. Neurology. 2003;60(11):1854‐7.
II Sallinen R*, Vihola A*, Bachinski LL, Huoponen K, Haapasalo H, Hackman P,
Zhang S, Sirito M, Kalimo H, Meola G, Horelli‐Kuitunen N, Wessman M, Krahe R, Udd B. New methods for molecular diagnosis and demonstration of the (CCTG)n mutation in myotonic dystrophy type 2 (DM2). Neuromuscul Disord. 2004;14(4):274‐83.
III Vihola A, Bachinski LL, Sirito M, Olufemi S‐E, Hajibashi S, Baggerly KA, Raheem
O, Haapasalo H, Suominen T , Holmlund‐Hampf J, Paetau A, Cardani R, Meola
G, Kalimo H, Edström L, Krahe R, Udd B. Differences in aberrant expression and splicing of sarcomeric proteins in the myotonic dystrophies DM1 and DM2. Acta Neuropathol. 2010;119(4):465‐79.
IV Vihola A, Sirito M, Bachinski L, Raheem O, Suominen T, Krahe R, Udd B.
Aberrant expression and splicing of Ca2+ metabolism genes in myotonic dystrophies DM1 and DM2. Manuscript submitted
Anna Naukkarinen née Vihola
*These authors contributed equally to this work
Publication II also appears in the thesis of Riitta Sallinen (2004)
The articles are reprinted with the permission of their copyright holders
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2. ABBREVIATIONS
aa amino acid Ab antibody Ach acetylcholine ATP adenosine‐5´‐triphosphate ATPase adenosine‐5´‐triphosphatase BSA bovine serum albumin cDNA complementary DNA CISH chromogenic in situ –hybridization CK creatine kinase CNS central nervous system DAB 3,3´‐diaminobentzidine DAG dystrophin‐associated glycoprotein DHPR dihydropyridine receptor DM1 myotonic dystrophy type 1 DM2 myotonic dystrophy type 2 DNA deoxyribonucleic acid EP expression profiling FIGE field‐inversion gel electrophoresis FISH fluorescent in situ –hybridization IF immunofluorescence Ig immunoglobulin IHC immunohistochemistry ISH in situ –hybridization LGMD limb‐girdle muscular dystrophy nt nucleotide mAb monoclonal antibody MD muscular dystrophy MRF myogenic regulatory factor mRNA messenger‐RNA MyHC myosin heavy chain NMD neuromuscular disease nNOS neuronal nitric oxide synthase pAb polyclonal antibody PCR polymerase chain reaction PDM proximal myotonic dystrophy PFA paraformaldehyde PROMM proximal myotonic myopathy RT‐PCR reverse transcription –PCR RNA ribonucleic acid RYR ryanodine receptor SERCA sarcoplasmic/endoplasmic reticulum Ca2+ ATPase
SNP single nucleotide polymorphism SR sarcoplasmic reticulum SVA splice variant analysis TSF transcription factor
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3. ABSTRACT
Myotonic dystrophies type 1 (DM1) and type 2 (DM2) are autosomal dominant
multisystemic disorders caused by microsatellite tri‐ or tetranucleotide‐repeat
expansion mutations in transcribed but not translated gene regions. Since the
primary tissue affected is muscle, they are classified as muscular dystrophies. DM1
and DM2 share some features in general clinical presentation and molecular
pathology, yet they show distinctive differences, including disease severity and
differential muscle and fiber type involvement. Thus far more than 300 DM2 patients
have been identified in Finland, which is in stark contrast with the previously
reported prevalence of 1/105 (corresponding to 50 patients), and the true number is
believed to be much higher.
DM2 is particularly intriguing, as it shows an incredibly wide spectrum
of clinical manifestations. For this reason, it constitutes a real diagnostic challenge. It
is important to improve the diagnostic accuracy in order to efficiently identify the
symptomatic DM2 patients for appropriate medical attention and management.
Myalgic pains may be the most disabling symptom for decades, sometimes leading
to incapacity for work. DM2 may cause major socio‐economical consequences for
the patient, if not diagnosed, due to misunderstanding and false stigmatization.
In this thesis work, DM2 patient skeletal muscle has been compared to
muscle from DM1 patients and healthy individuals. This is the first description of
histopathological differences between DM1 and DM2, which can be used in
differential diagnostics. Two novel high‐resolution applications of in situ ‐
hybridization, which can be used for direct visualization of the DM2 mutation in
patient muscle biopsy sections or on extended DNA‐fibers have been described. By
measuring protein and mRNA expression in the samples, differential changes in
expression patterns affecting contractile proteins, other structural proteins and
calcium handling proteins in DM2 compared to DM1 were found. The dysregulation
at mRNA level was caused by altered transciption and abnormal splicing. The
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findings reported here indicated that the extent of aberrant splicing is higher in DM2
compared to DM1. These abnormalities to some extent correlate to the differences
in fiber type involvement in the two disorders.
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4. REVIEW OF THE LITERATURE
4.1. Introduction to cell and molecular biology
The eukaryotic cell contains a plasma membrane, delineating the cell, which is filled
with cytoplasm and organelles: at least a nucleus, membranous endoplasmic
reticulum system, mitochondria and ribosomes. Even though genetically identical,
the cells of an individual organism develop into very distinctive forms depending on
their environment and functional requirements. Groups of differentiated cells are
organized to form tissues and organs with specialized functions. Complex networks
of molecular pathways regulate the multitude of biochemical events taking place in
individual cells, enabling the full spectrum of cellular functions: division, maturation,
synthesis, degradation, respiration, secretion, movement, and senescence [1].
In humans, the genetic code, composed of deoxyribonucleic acid
(DNA), is packed inside the nucleus in 46 chromosomes (23 pairs, one set from both
parents) [293]. The whole human genome contains approximately 20,000 ‐25,000
different protein‐coding genes, as was revealed by the Human Genome Project in
2004 [116]. Four different nucleotides (nt) (Adenine, Thymine, Cytosine, Guanine)
serve as building blocks of the DNA, connected to each other via phosphate
backbones, to form long filamentous molecules. The DNA molecule is made up of
two anti‐parallel and complementary strands, which pair to form A‐T and C‐G pairs,
and fold into a secondary structure known as the double helix [314].
Genes carry the information needed to build proteins, the large
biomolecules serving as structural components, enzymes and regulatory molecules
of the cell. First, the gene is transcribed, to create a pre‐messenger ribonucleic acid
(mRNA) molecule complementary to the gene [155]. However, the eukaryotic gene
also contains segments which do not code for a protein: these intron sequences are
spliced out to form the final mRNA molecule, containing only exons, which then
serves as a template for ribosomal translation to a protein. The nucleotide sequence
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of the mRNA molecule determines the order in which the 20 amino acids are
inserted into a growing polypeptide chain to create a protein. However, the number
of different proteins is much larger than the number of genes. Differential splicing
[72] is one of the means the cells employ to create more variability in the protein
structure and function: in addition to introns, some of the exons may sometimes be
excised. In fact, it is estimated that approximately 80 percent of human genes go
through such alternative splicing, giving rise to multiple isoforms [176]. Typically, the
splicing pattern is regulated spatially and temporally: exon usage is specific in
different tissues and cell types, and at different points of development [155].
4.1.1. Microsatellite‐DNA
In addition to the paradigmatic protein‐coding function, both DNA and RNA
molecules have other, regulatory functions. For instance, the human genome
contains vast amounts of short tandem repeat (STR) sequences, most often 1 ‐ 6 nt
in length, called microsatellite‐DNA [28]. The repetitive sequences, which can be
perfect or interrupted in nature, occur widely in both eukaryotes and prokaryotes.
The functions of the microsatellite DNA are not fully elucidated. However, the
repeat‐containing mRNA transcripts have been implicated in regulation of
transcription, translation and cell signaling [120]. These highly repetitive tandem
sequences typically show a high degree of polymorphism, enabling their use as tools
for several applications including linkage analyses, in the search for disease‐causing
genes, and DNA fingerprinting [278, 284]. In addition, microsatellites have been
implicated in disease: especially, they play a significant role in neurodegenerative
diseases [42].
4.1.2. Mutations
Due to errors in translation, radiation and chemical stress, genomes are constantly
subject to modifications, or mutations: nucleotides are deleted, inserted, or replaced
by others. All organisms therefore accumulate mistakes in their genomes, in both
somatic cells and gametes [155]. Although most changes are silent, causing no
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effect, mutations in somatic cells for example are the cause of cancer. When a
mutation occurs in a gamete, it is transmitted to every cell of the offspring. It should
be noted that mutations in gametes make evolution possible: sometimes the
transformed gene encodes a protein which gives more benefit to the organism than
the original form, which may lead to its establishment in the population.
Unfortunately, such beneficial mutations are very rare. It is more common that a
mutation leads to malfunctioning of the protein or a regulatory element, which is
lethal or may lead to illness. [278]
Monogenic disorders follow Mendelian modes of inheritance:
autosomal recessive (AR), autosomal dominant (AD), X‐linked and Y‐chromosomal
[278]. The mutation is called dominant if just one mutated allele is sufficient to cause
the disease. The pathomechanisms, through which the mutations cause diseases,
have classically been divided into protein loss‐of‐function (mostly recessive
mutations) and protein gain‐of‐function (mostly dominant mutations). However,
increasing numbers of mutations evoking their effect via an RNA gain‐of‐function
mechanism have been described; many of them are repeat expansion mutations
[238]. The myotonic dystrophies, which are the subject of this work, are autosomal
dominant diseases, caused by microsatellite repeat expansions in non‐coding but
transcribed gene regions, and the main pathomechanism underlying them is an RNA
gain‐of‐function [158, 173].
4.1.3. Proteins
Proteins are chains of polypeptides, synthesized of 20 different amino acids,
according to the genetic blueprint. The linear nascent polypeptides first form
secondary structures (α‐helices and β‐sheets), and then fold independently, or with
assistance from the molecular chaperones, to adapt the final functional
conformation, the tertiary structure. Proteins often form permanent or transient
complexes, quaternary structures, when multiple cooperative proteins are linked
together via non‐covalent interactions [38]. Posttranslational modifications, such as
glycosylation or phosphorylation, increase the versatility of the protein molecules
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[1]. Proteins are the main building blocks of the living cell, serving for example as
structural and contractile components, enzymes, antibodies, and also as regulatory
and signaling proteins. They show huge variation in size and shape, from small
oligopeptide signaling molecules to huge filamentous structural proteins, such as
titin (TTN), the largest polypeptide in nature with a molecular mass of more than
3,000 kD [147].
4.2. Molecular methods for research and diagnostics
4.2.1. Detecting nucleic acids with hybridization techniques
Hybridization methods for detecting DNA or RNA are based on the double helical
nature of these molecules, or the tendency of the nucleic acids to bind specifically to
a complementary and anti‐parallel sequence [155]. The target sequence is a mixture
of nucleic acids in the specimen examined. The target is incubated with a known
probe sequence, under high‐stringency conditions which causes the denaturation of
any double helical structures. Next, the target and probe are allowed to hybridize,
after which their binding is visualized and analyzed. DNA and RNA may be detected
interchangeably, as all kinds of hybrids between them are possible [278].
4.2.1.1. Southern blotting
A basic molecular biology technique for detecting specific DNA sequences is called
Southern blotting, according to its inventor [276]. The target sequence, most often
genomic DNA, is cleaved to small fragments using restriction endonucleases. The
yielded DNA fragments are separated electrophoretically on agarose gels, according
to their size. In order to expose the fragments by hybridization, the DNA has to be
transferred (blotted) from the gel onto a membrane, often nitrocellulose or nylon,
simply using capillary action, or electrically, taking advantage of the negative charge
of the DNA molecule. After immobilizing the target DNA, it can be hybridized with a
labeled probe. Fluorescent and chemiluminescence labels are most commonly used
today, whereas radioactive tags have been popular in the past. The binding of the
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probe is visualized using a laser reader or autoradiography. The Southern blot
method may be used, for example, for detecting gene copy numbers, or for
diagnostics. [41, 278]
4.2.1.2. In situ –hybridization
The in situ ‐hybridization (ISH) technique dates back to the 1960´s [92]. The principle
of in situ hybridization (ISH) is that the nucleic acid target sequence in the preparate,
which is often a tissue section, cell or chromosome on a microscope slide, is
detected by a specific labeled probe, which hybridizes with a region in the target
complementary to the probe. The binding site is then visualized and localized, hence
the name in situ. Fluorescent labels are most often used for detection today, due to
their high sensitivity and possibility to visualize multiple targets simultaneously
[162]. However, chromogenic labels may be more convenient in some cases as the
procedure is more robust, the end product is more resistant, the specimens may be
stored for later documentation, and their visualization does not require highly
specialized equipment. In addition, high sensitivity with chromogenic methods is
possible to reach nowadays, due to improved detection systems, including the use of
polymers [221].
4.2.1.3. Microarrays
Microarray technology represents a multiplex approach, where probes are
immobilized on a solid support or microscopic beads, and they are detected with
labeled target sequence mixture in solution, such as cDNA. The probe‐target
hybridization is detected and quantified in order to determine the presence and
relative abundance of a specific target sequence in the sample solution. One array
may contain tens of thousands of probes, which makes it a highly efficient high‐
throughput method for research and diagnostic purposes. [108]
Microarrays can be used in a multitude of applications: for example
using the cDNA (DNA complementary to mRNA) library as a target, expression
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profiling can be used to investigate the relative mRNA expression in a given tissue or
cell type; similarly, with an exon‐specific array, different splice variants can be
detected [48]. When genomic DNA is used as a target, it is possible to perform
comparative genomic hybridization to detect copy number changes in closely related
specimens, or to identify single nucleotide polymorphisms (SNPs), or mutations. An
increasingly used application utilizing microarrays is genetic testing [323]. Microarray
experiments produce a vast amount of data, handling of which requires
bioinformatics: elaborate statistical analysis and normalization in order to filter out
background noise and reach biologically and statistically significant conclusions [45].
Standardization methods are used to enable comparison of separate experimental
data, even though it is often very difficult due to the numerous variables involved in
the experimental procedure. Microarray results are commonly visualized using heat
maps.
4.2.2. Detecting proteins using antibody recognition
4.2.2.1. Antibodies
Antibodies (Ab) are Y‐shaped immunoglobulin (Ig) molecules composed of two
identical light chains and heavy chains. Hypervariable N‐termini form the antigen
binding site, whereas the C‐termini form the stabilizing Fc‐tail, which also serves as a
binding site for the other Ab molecules [244]. Most antibodies used for IHC are of
IgG‐type, with two identical Ab‐binding sites (paratopes).
A good Ab shows high specificity and sensitivity. Specificity describes
the ability of a given Ab to recognize only its specific antigen, whilst sensitivity refers
to the affinity of an Ab to its antigen. Ab‐antigen binding is based on weak, non‐
covalent bonds, mainly hydrophobic and electrostatic by nature [236]. Two main
classes of antibodies are the polyclonal and monoclonal antibodies.
Polyclonal antibodies (pAb) are produced by immunizing an animal, for
example a mouse, rabbit or goat, by injecting the purified antigen, often linked to a
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carrier protein to enhance its stability and introduction to the immune system. After
a successful immunization, usually several cell clones produce immunoglobulin
molecules raised against different epitopes in the antigen, and secrete them into the
blood serum. The serum may be used as such, however it is common to affinity
purify the immunoglobulin molecules using the antigen as bait, in order to reduce
cross‐reactions caused by the numerous Ig species in the serum. The polyclonal
antibodies provide a powerful tool in many cases. However, there are some
disadvantages: while the polyclonal antibodies generally show more sensitivity,
because of several different Ig‐molecules binding to several different epitopes in the
protein, their specificity is often compromised when compared to monoclonal
antibodies due to higher risk for cross‐reactions. In addition, the exact epitope(s) is
often not known, especially when an intact protein has been used as an immunogen.
[105]
Monoclonal antibodies (mAb) are most often produced in mice. After
successful immunization, the animal is sacrificed and the spleen cells are collected to
start a cell culture. The cells are immortalized by fusing them with mouse myeloma
cells, after which they are separated and cultured individually, so that every sub‐
culture is a clone derived from a single B‐cell. The clones are then screened for Ig
molecules which specifically recognize the original immunogen [138]. The procedure
results in stable hybridoma cell lines, which produce and secrete a single Ig species.
In general, monoclonal antibodies provide a more specific tool compared to
polyclonals. New technologies have been developed in the recent years to meet the
restrictions of the traditional mouse hybridoma technique, including phage libraries
[169].
Regardless of the method used for Ab production, each Ab is unique,
and should be carefully tested and characterized before research or diagnostic use,
to avoid misinterpretation of results.
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4.2.2.2. Immunohistochemistry
The advent of the immunofluorescence (IF) technology in the 1940s [57, 58] initiated
the development of techniques based on proteins being detected in situ with
antibodies, methods collectively called “immunohistochemistry” (or
immunocytochemistry, if cells rather than tissues are used). The molecular basis of
immunohistochemistry (IHC) is the specific interaction of an antibody (Ab) protein
molecule with its antigen, most often a protein, residing in the tissue section. The
antibody molecule recognizes a defined part of the protein, which is called epitope
or antigenic determinant. The epitope is a tertiary structure on the surface of the
protein, and it may be composed of linear amino acid sequence (linear epitope), or
non‐linear amino‐acids, residing in distinct parts of a protein, which come into
proximity when the protein folds into its native conformation (conformational
epitope). The epitope may also involve a covalent modification, such as a phosphate
group or a linked oligosaccharide (glycoprotein). By IHC staining methods, the
subcellular localization of specific proteins and to a limited extent, the detection of
quantitative changes of protein expression may be studied. However, the method
does not provide information relating to protein interactions. [236]
It is common to fix tissue sections in order to preserve good‐quality
morphology and prevent autolysis. Fixation is absolutely required when soluble
molecules are being detected, to prevent diffusion and displacement of components.
Fixation using formaldehydes is a conventional method, resulting in formation of
cross‐links between proteins via amino groups. However, such fixation often
modifies the tertiary (and quaternary) structures of proteins and destroys the
immunogenic regions, rendering the proteins unrecognized by Abs. Short fixation in
paraformaldehyde (PFA) solution is commonly used on frozen tissues, as it creates
only limited amount of cross‐linking, and is partially reversible. An alternative means
is to use coagulating fixatives, such as ethanol, methanol or acetone. They do not
form cross‐linking, but rather precipitate the proteins, leading to partial
denaturation. There is no fixation method which suits all epitopes, but it always
depends on the properties of the antigen and the Ab, and the optimal fixation
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approach needs to be tested on each Ab individually. Some antigens are extremely
vulnerable to any fixation methods, leaving no choice but to detect the tissue
sections in their native state. [105, 236, 237]
To make the Ab‐antigen reaction visible in tissue sections, a selection of
immunohistochemical detection methods are available. The primary antibody may
be directly visualized if covalently labeled with a reporter molecule, such as
fluorescein. However, the indirect detection method, which utilizes a labeled
secondary Ab raised against the host animal used to produce the primary Ab, offers
several advantages over the direct method, including higher sensitivity, specificity,
and more versatile use of the primary Ab. Indirect immunofluorescence allows high
sensitivity, and use of double staining methods to study co‐localization of distinct
proteins, when labels with different absorption and emission wavelengths
maximums are used. High resolution is achieved through confocal microscopy. [105,
237]
A more robust method, which is often advantageous in diagnostics,
involves the use of enzyme‐linked secondary Abs. The enzyme, most often a
peroxidase or alkaline phosphatase, catalyses the reaction between a specific
substrate and a chromogen to produce a coloured precipitate at the Ab binding site.
The most widely used peroxidase substrate is 3,3´diaminobenzidine (DAB), which
results in brown deposit, insoluble in organic solvents. The sensitivity of enzyme‐
based detection methods can be increased with several techniques, including
branching polymer molecules carrying multiple enzymes (summarized in [236]).
Double staining, with distinct colored end products, may also be applied when the
antigens reside in distinct cell types or structures.
IHC may be used to evaluate muscle pathology in various cases. The
classical method is the sarcolemmal staining with dystrophin antibodies: in
Duchenne muscular dystrophy the sarcolemmal labeling is nearly or completely lost,
whereas in Becker type the finding is often intermediate [6]. Also accumulation of
proteins in specific localizations, such as rimmed vacuoles or ribonuclear inclusions,
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may be studied. When assessing quantitative changes, using an internal control,
which shows the overall preservation of the sample, is mandatory. For example, for
the detection of muscle sarcolemmal integrity, beta‐spectrin staining is often used
[71]. The IHC results should always be interpreted with caution; in addition to
adequate controls, one should be aware of the handling and storage history of the
sample. It should also be noted that there are differences between stainings
performed in different laboratories, even though the same protocols may be used.
4.2.2.3. Western blotting
Proteins in a given sample, such as tissue or cells, can be solubilized, denatured and
separated electrophoretically. The most common method used is sodium dodecyl
sulfate ‐ polyacrylamide gel electrophoresis (SDS‐PAGE), described by Laemmli in
1970 [148], which allows the separation of proteins according to their size. The
proteins may be transferred from gel to a membrane for detection with antibodies,
with the Western blotting (WB) method [46]. When assessing total protein
expression, Western blotting is considered a more quatitative method compared to
IHC. However, the ability to identify protein localization in the sample is lost, which
may in some cases be detrimental to interpretation. When possible, IHC and
Western blotting should be used in parallel, because they provide information
complementary to each other.
4.3. The muscle
Muscle tissue is required for generating mechanical force, which enables a vast
amount of bodily functions, including locomotion, maintaining posture, breathing,
heartbeat and digestion, among others. There are three main classes of muscle: (1)
skeletal and (2) heart muscle, both of which are striated, in addition to (3) smooth,
non‐striated muscle (Figure 1). Only skeletal muscles are voluntary, and they are
used for locomotion. The extrafusal (skeletal muscle) fibers are innervated by axons
of the alpha motor neurons, large lower motoneurons with their cell bodies in the
22 |
brainstem and spinal cord. Heart muscle contraction is under autonomous nervous
control, as is smooth muscle. [71, 131]
The cardiac muscle cells, termed cardiomyocytes, are branched
cylinder‐shaped, mononuclear cells. They are joined end‐to‐end by intercalated
discs, specialized structures which stabilize cell adhesion, anchor actin filaments, and
allow signal transduction to spread from cell to cell via gap junctions, leading to
synchronous muscle contraction. The cardiomyocytes have a similar sarcomeric
organization of contractile proteins to skeletal muscle, with some differences in
cellular architecture to fulfill the special functional needs of the cardiac muscle. [131]
The spindle‐shaped smooth muscle cells are found as sheets or bundles
in the walls of internal organs, such as blood vessels, ureters, uterus, and the
respiratory and gastrointestinal tracts. In smooth muscle the contractile proteins are
not organized in sarcomeres. [1]
Figure 1. Schematic pictures of (A) skeletal muscle, with syncytial and multinucleated fibers, (B) cardiac muscle, showing the intercalated discs, and (C) smooth muscle. Both cardiac and smooth muscle fibers have only one, centrally located nucleus. (Adapted from Human Anatomy, 2nd edition by Dee Silverthorn. Used by permission of the copyright holder Pearson Education, Inc.)
| 23
The human body contains more than 300 different skeletal muscles,
which differ in size and shape and biochemical properties, depending on their
function. Those close to the trunk are called proximal muscles (e.g. quadriceps,
biceps brachii), and those distant from the trunk are called distal muscles (e.g. small
muscles in hands and feet but also lower leg and forearm such as tibialis anterior,
brachioradialis). [191]
Skeletal muscle energy metabolism relies on phosphorylated
metabolites, glycogen and fat: the myocytes store glucose in the form of glycogen
which can be metabolized by glycogenolysis and glycolysis during exercise, under
anaerobic conditions, resulting in lactic acid formation. In aerobic exercise, oxidative
phosphorylation generates the phosphorylated metabolites phosphocreatine and
ATP needed, and in long duration (over 30 minutes) lipid droplets via lipolytic
cascades are used to generate ATP also through oxidative phosphorylation (beta‐
oxidation). [101]
4.3.1. The muscle structure
The skeletal muscle fiber is a single cell, which can be up to several centimeters in
length, depending on the muscle. Each of the muscle fibers is surrounded by a basal
lamina, which acts as the interface to the connective tissue surrounding them,
termed the endomysium (Figure 2). Bundles of muscle fibers form fascicles
enveloped by the perimysium, which is rich in blood vessels. The muscle is composed
of bundles of fascicles, which are ultimately enclosed in a strong connective tissue
sheath epimysium, the muscle fascia. Most skeletal muscles are attached to tendons
in a specialized structure called the myotendinous junction, which in turn are
connected to the bones. [71]
24 |
Figure 2. The structure of the muscle. (The figure was produced using Servier Medical Art at http://www.servier.com.)
During development, axons of myelinated motor nerves, which are of
two types, fast and slow, invade the skeletal muscle where they branch. The axons
lose their myelin sheaths before forming nerve terminals in grooves of the muscle
fiber, an invagination of sarcolemma, a synapse called the neuromuscular junction.
Each adult muscle fiber is innervated by a single motor nerve. A motor unit is defined
as a single parent axon and all the fibers it innervates with its branches. Depending
on the need for fine control, a motor unit consists of just a few fibres or several
hundred fibres. [76]
The muscle spindles, present in all skeletal muscles except for facial
muscles, are specialized structures with intrafusal fibers innervated by gamma motor
neurons. They serve to sense muscle stretch and tone, conveying the information to
the central nervous system (CNS) via sensory nerves, hence coordinating muscle
activity [71].
| 25
4.3.2. Skeletal myogenesis
All striated muscle is derived from the mesoderm, the primary embryonic tissue
layer between ectoderm and endoderm [75]. The mesodermal tissue gives rise to
specific lineages of progenitor cells, which develop into specialized skeletal muscles
and other tissues such as heart, skin, bones, blood vessels and blood [75]. The group
of embryonic tissues which develops into skeletal muscles is called the myotome. It
divides into epaxial and hypaxial myotome, developing into axial and limb muscles,
respectively [68]. Cell fate specification is a genetically regulated, hierarchical
process which relies on spatial and temporal expression of specific regulatory
proteins, complex regulatory networks and molecular cascades.
The myogenic regulatory factors (MRFs) Myf5, MyoD, myogenin and
MRF4 form a conserved group of basic helix‐loop‐helix (bHLH) transcription factors
highly specific for skeletal muscle [3]. The early markers Myf5 and MyoD are
expressed in the paraxial mesoderm, where they control (partially redundantly)
muscle progenitor specification. Their transcriptional activation is controlled by
developmental signals [36].
To enable differentiation into myocytes, the cells need to undergo G1‐
specific cell‐cycle withdrawal [208]. During the process of differentiation, myogenin
and MRF4 are activated, with MEF2C and together these transcription factors induce
the expression of muscle genes encoding the contractile and membrane proteins
[74]. Myf5 expression is ceased in differentiating myocytes, while MyoD, the “master
regulator”, maintains expression in newly formed myocytes [286].
The MADS domain family of transcription factors, (myocyte enhancer
factor‐2, MEF2), are encoded by four genes, MEF2A‐D. They function in cooperation
with MRFs to modify their actions in driving muscle and fiber type specific genes,
including those encoding contractile proteins [179, 189, 229].
26 |
Both MRFs and the MEF2 family are phosphoproteins, and their activity
and association with each other and several co‐factors are regulated by protein
kinase driven phosphorylation state [75, 179]. Combinatorial associations of MRFs,
the MEF2‐family, and several co‐factors (growth factors) drive the temporally
controlled and coordinated transcription of muscle genes [202].
In skeletal myogenesis, the myoblasts proliferate and fuse to form
primary myotubes. Post‐mitotic myoblasts then continue to fuse with the primary
myotubes, giving rise to secondary myotubes. These elongated, multinucleated
cyncytia then mature into muscle fibers, also called myocytes. The process involves
dramatic changes in the organization of the cell membrane and the filament system,
as well as development of the highly specified contractile apparatus,
myofibrillogenesis. [259]
4.3.3. The satellite cells
Once fused, the myonuclei lose their ability to proliferate. Much of the
adult muscle regeneration capacity relies on satellite cells, which are quiescent
progenitor cells located between the sarcolemma and the basal lamina of a muscle
fiber [100, 178]. A second pool of interstitial stem cells, displaying broader
differentiation capacity also exists in muscle [31]. When regeneration is required, for
example after. injury or overuse, the satellite cells are activated to proliferate. A
subset of the newly formed myoblasts fuse into the existing myocytes, while some
remain in the satellite cell position for later use. Despite this, the number of the
satellite cells usually declines with age [252]. Quiescent satellite cells can be revealed
by their morphology and position, especially with electron microscopy. In addition,
they express a number of protein markers, which can be used for their identification
using immunohistochemistry; however, few have proven to be specific to satellite
cells, such as the paired‐box transcription factor Pax7 [241, 269, 327].
| 27
4.3.4. Fiber types
Skeletal muscle fibers display different biochemical and physiological profiles,
depending on the intrinsic properties of the specific muscle, developmental stage,
age, exercise, and disease. See Table 1 for a summary of the skeletal muscle fiber
types. Slow and fast fibers differ in performance, reflecting differential expression of
enzymes involved in energy metabolism. Slow‐twitch type 1 fibers, and fast‐twitch
type 2A fibers use oxidative phosphorylation for generating ATP, and are suited for
endurance, whereas the ultra‐fast type 2B fibers rely mostly on glycolytic
metabolism, displaying fast maximal performance, but are subject to fatigue. Type I
fibers are rich in myoglobin, which gives them the typical red color, whereas fast
type fibers are referred to as white muscle fibers (in the absence of myoglobin). [71,
249]
Table 1. Adult muscle fiber types in human.
Fiber type (by ATPase method)
1 (slow)
2A (fast)
2B (ultra fast)
2C (mixed)
Not separated
Energy metabolism aerobic aerobic glycolytic aerobic mixed
MyHC isoform I (beta/slow) IIa IIx hybrids of I/IIa hybrids of
IIa/IIx
MYH gene MYH7 MYH2 MYH1 MYH7 / MYH2 MYH2/MYH1
The classification of fiber type is most often based on the properties of
the ATPase domain of the myosin heavy chain (MyHC) molecule. Enzyme
histochemical ATPase assay, based on the differential pH sensitivity of the MyHC
ATPases, has been used for decades as the routine method for fiber type
differentiation [71, 248]. However, the immunohistochemical detection of different
MyHC isoforms has proved to be as qualified, or, in some cases, even more efficient
tool in fiber type differentiation (Figure 3) [233]. The classical division into fast
glycolytic and slow oxidative fibers can be further subdivided into several categories.
The three most common fiber types in normal human skeletal muscles are type 1
fibers expressing MyHC‐beta/slow (encoded by MYH7 gene), type 2A fibers
expressing MyHC‐IIa (MYH2) and type 2B fibers expressing MyHC‐IIx (MYH1). It may
28 |
be confusing that the human skeletal muscle fibers do not express MyHC isoform IIb,
despite the fiber type is designated as 2B. The isoform IIb, encoded by MYH4, is
expressed in 2B fibers of small mammals, such as mouse, rat and rabbit. In heart
muscle, MyHC‐beta/slow, MyHC‐alpha (MYH6) and MyHC‐7B (MYH7B) are
expressed, the latter two being absent in skeletal muscles. Hybrid fibers expressing
two or more MyHC isoforms are common in myopathies, sometimes presenting
typical patterns of fiber type transitions. Hybrid fibers co‐expressing MyHC‐
beta/slow and IIa are designated as type 2C fibers by ATPase staining [71].
Expression of atypical MyHC isoforms is sometimes encountered in injured or
dystrophic muscle. Reprogrammed atrophic or regenerating fibers frequently
express perinatal/neonatal MyHC‐pn (MYH8), whilst regenerating, severely injured
fibers in dystrophic muscles typically show expression of MyHC‐emb (MYH3).
Additional MyHC isoforms exist, showing very restricted muscle‐specific expression
patterns [249] (summarized in Table 2. of original publication III). Most human
muscles are composed of a variably mixed population of slow and fast fibers. The
proportions correlate with the function of a given muscle.
4.3.4.1. Fiber type distributions in disease
Basophilic fibers, which appear bluish in H&E stain due to high sarcoplasmic RNA
content, are usually considered regenerating fibers. They often show swollen or
“vesicular” nuclei, and may coexpress beta/slow and IIa myosins, in addition to
perinatal/neonatal MyHC‐pn [71].
Figure 3. Immunohistochemical fiber typing using double staining. 2A fibers appear red; 2B fibers blue; 1 fibers brown; and hybrid fibers expressing myosins IIa and IIx appear pink. (Picture kindly provided by Olayinka Raheem, University of Tampere.)
| 29
Nuclear clumps are characterized as atrophic fibers with clustered
nuclei and with a very scarce amount of sarcoplasm left. Although present in chronic
dystrophies, including limb‐girdle muscular dystrophies (LGMD), they are considered
a hallmark of prolonged neurogenic atrophy [71].
Variation in fiber size is one of the most common myopathic findings. In
most cases, a random pattern of atrophy and hypertrophy is seen, affecting both
fiber types equally. Grouping of atrophic fibers is typical of neurogenic pathology, as
a result of reinnervation of the denervated fibers with sprouts of a fast or slow
motor nerve. Selective fiber type 1 atrophy is encountered in DM1 [106] and
congenital myopathies, termed congenital fiber type disportion [71]. On the other
hand, fiber type 2 atrophy is usually considered a non‐specific finding, occurring in
variable myopathic conditions. Known causes of fast fiber atrophy are muscle disuse
and corticosteroid therapy [71]. In most cases, either both fast fiber subtypes 2A and
2B are affected, or predominantly 2B fibers. There is one exception, a recently
described human disease with mild generalized muscle weakness, where a total loss
of type 2A fibers was discovered, due to compound heterozygous mutations in the
MYH2 gene [281]. Fiber type 1 predominance may be a common feature in
congenital myopathies, whereas fiber type 2 predominance is sometimes seen in
motor neuron diseases [71].
4.3.5. The skeletal muscle fiber
Skeletal muscle cells, or myocytes, are enveloped by the sarcolemma, composed of
the plasma membrane and the basal lamina [83]. The multiple, elongated nuclei are
scattered along the fiber length, located just beneath the sarcolemma. The muscle
fiber has a highly specialized inner structure, dominated by the contractile units, the
sarcomeres, forming the myofibrils. The cytoplasm, or sarcoplasm, contains an
elaborate membrane system, including the extensive sarcoplamic reticulum (SR), and
invaginations of the (electrically excitable) plasma membrane, the transverse (T) ‐
tubular system (Figure 4). At the junction site between the sarcomeric thick filament
A‐band and thin filament I‐band (I/Ajunction) the SR forms membranous sacks, or
30 |
terminal cisternae, which juxtapose to a T‐tubule [82]. This triad structure, formed
by two terminal cisternae flanking one T‐tubule, allows the rapid progression of
depolarization in the SR, initiating Ca2+ release needed for the muscle contraction
(see below). Mitochondria are present in large numbers between the myofibrils,
near the I‐bands, together with granules of glycogen. The sarcoplasm also contains
ribosomes, both subsarcolemmal/perinuclear and between myofibrils [111]; lipid
droplets; the Golgi apparatus; and a cytoskeleton of microtubules, intermediate
filaments (such as desmin, syncoilin, desmuslin and actin) [32, 131].
Figure 4. Schematic picture of the excitable membrane system. The triad structure is constituted of a T‐tubule surrounded by terminal cisternae of the sarcoplasmic reticulum, at the level of sarcomeric A/I junction. M = M‐line. (Drawing taken from Unit 2, Medical Curriculum, McGIll University, CA, with permission of C. R. Morales. The drawing was originally conceived by Y. Clermont and subsequently modified by Molson Medical.)
The dystrophin‐associated glycoprotein (DAG) complex on the
sarcolemma provides a structural link between the cytoskeleton and the
extracellular matrix, and stabilizes the sarcolemma against mechanical stress. In
| 31
addition, the complex organizes receptors and ion channels on the sarcolemma.
Dystrophin is located on the sarcoplasmic side, where it anchors the DAG complex
via β‐dystroglycan to F‐actin and sarcomeres (Figure 5). The other DAG complex
constituents include α‐ and β‐dystroglycans (DG), the membrane‐embedded
sarcoglycans (α, β, γ, δ, and ε), sarcospan and the dystrobrevin‐syntrophins complex
(Figure 5), as well as the neuronal nitric oxide synthase (nNOS), an enzyme
synthesizing the signaling molecule nitric oxide (NO) (not shown).
Figure 5. The DAG‐complex on the sarcolemma, and its linkage to the myofibrils. (Adapted with permission from “Histology and Cell Biology: an introduction to pathology” by A. Kierszenbaum, p. 208, Elsevier 2007.)
The costameres of the sarcolemma serve as anchoring points to the
sarcomeres in the peripheral myofibrils, which is crucial to enable coordinated
contraction of adjacent myofibrils, and to avoid mechanical ruptures during muscle
32 |
contractions. Desmin intermediate filaments, cross‐linked to each other by plectin,
extend from costameres to myofibrils, surrounding them at the level of Z‐discs
(Figure 5). Other costameric constituents include at least vinculin, talin, spectrin,
ankyrin, synemin, and β1‐integrin [49, 97]. The heat shock protein αB‐crystallin is
associated with desmin, protecting it from stress‐induced damage [131, 305].
4.3.6. The sarcomere
The sarcomere forms the structural basis of muscle contraction. Its striated
appearance is created by regularly aligned sarcomeres in a row, the myofibrils, with
specific regions showing differential electron densities under microscopic
examination (sarcomere structure is outlined in Figure 6). The sarcomere is
composed of the thick filament A‐band, with dense M‐band in the middle, and thin
filament I‐band, with a very electron dense Z‐disc in the middle [71]. The M‐band
shows a fine structure with three to five thin lines, composed of non‐myosin
proteins. Fast fibers shows three lines (M1, M4 and M4´), slow fibers show four lines
(M4, M4´, M6 and M6´), and intermediate fibers show all five lines [61]. One human
skeletal muscle sarcomere extends from Z‐disc to Z‐disc, being 2.0 ‐ 3.0 µm in length
at rest depending on the muscle. The A band has a fixed length of 1.5 – 1.6 µm, while
the flexible I band region extends during muscle contraction. Large muscles
generating much force have longer sarcomeres and a greater ability to shorten
during contraction. The A‐band is composed of a hexagonal lattice of myosin
filaments, tertiary aggregates of a polar molecule consisting of two heavy chains
(MyHC) and two light chains (MYL). The myosin molecule has two globular heads
with neck regions continuing into one coiled‐coil rod and tail region. The MyHC
globular head has an ATPase domain and it binds actin, the major constituent of the
thin filament. Myosin molecules are aligned in the thick filament in the way that the
rod and tail domains interact tail‐to‐tail in the M‐band, and the globular heads point
to opposite directions, forming the dense A‐band [61]. The H‐zone, para‐central of
the M‐band, is devoid of myosin heavy chain heads, and is rich in creatine kinase
(CK) enzyme, necessary to maintain steady levels of ATP during contraction: it
hydrolyses creatine phosphate to generate ATP where it is needed near the
| 33
myofibrillar ATPase. Elevated serum CK levels can be used as marker for muscle
damage and various myopathies, caused by leaky sarcolemma of the damaged
myocytes [71].
The thin filament is mainly composed of filamentous actin, formed by
globular G‐actin subunits, which attach head‐to‐tail forming a polar double helical
structure, with pointed (minus) ends and barbed (plus) ends, the latter inserting into
the Z‐disc. The grooves of the helix accommodate the troponin‐tropomyosin
complex, troponins T, I and C, the latter of which binds Ca2+. [61]
Figure 6. (Top) schematic presentation of the major structural components of the sarcomere. (Bottom) electron micrograph of the corresponding ultrastructure, showing mitochondria between the myofibrils. (Reproduced from Ottenheijm et al. Respiratory Research 2008 [214] with permission of BioMed Central.)
34 |
In addition to serving as an anchoring point to actin filaments, the Z‐
disc binds several proteins via another major constituent, alpha‐actinin (ACTN).
ACTN binds, among others, telethonin/T‐cap, myotilin, myozenin, γ‐filamin, vinculin,
the Z‐disc alternatively spliced PDZ‐containing protein (ZASP) and obscurin [14, 61,
282].
Other important sarcomeric proteins include the third filament system
constituent titin, and nebulin being part of the thin filament in skeletal muscles
[218]. A single titin molecule extends from the Z‐disc to M‐line, and serves as a
molecular ruler for sarcomeric assembly, contributes to passive tension during
stretching, and functions as a signaling molecule through its multiple ligand binding
sites, including calpain3, and a kinase domain near the C‐terminus in the M‐line [147,
149].
4.3.7. Muscle contraction
The excitation‐contraction cycle refers to the chain of events where the sarcolemma
depolarization leads to mechanical activation of the myofibrils to contract [22]. It is
initiated when a motor neuron action potential reaches the neuromuscular junction.
The presynaptic button contains vesicles filled with the neurotransmitter
acetylcholine (ACh), which is released to the primary synaptic cleft in response to an
action potential. ACh binds to specific receptors in the postsynaptic membrane,
which in turn undergo a conformational change forming a channel for net Na+ influx.
As a result of this local depolarization, the major sodium channels open, leading to
rapid depolarization in the T‐tubule system, which activates the L‐type voltage
sensitive Ca2+ channel dihydropyridine receptor (DHPR) once the depolarization
reaches the terminal cisternae of the T‐tubules [22]. Consequently, via mechanical
coupling, activated DHPR induces a conformational change in the ryanodine receptor
(RYR) located in the adjacent SR membrane (of the triad) [231]. In turn, the activated
RYR rapidly releases Ca2+ from the intra‐SR stores to the sarcoplasm, where it
diffuses to the sarcomeres and binds troponin C, and starts the final part of EC‐
coupling, the myofibrillar contraction [247].
| 35
Figure 7. Muscle contraction. (A) The sarcomeric I band shortens as the thin (actin) filaments slide past the thick (myosin) filaments during muscle contraction. (B) ATP‐driven conformational change in the myosin head forces the actin filament to move towards the “center” of the sarcomere. (The figure was produced using Servier Medical Art at http://www.servier.com.)
Binding of Ca2+ induces a conformational change in troponin C, which in
turn allosterically regulates the troponin‐tropomyosin complex, resulting in exposure
of the myosin‐binding site on the actin filament. When the transient cross bridge
between actin and myosin is formed, the myosin ATPase hydrolyses ATP to induce a
conformational change of the globular head, which mechanistically pushes the actin
filament forward approximately 10 nm at each stroke. As a result of several cycles,
the myosin and actin filaments slide past each other, resulting in shortening of the
sarcomeric I‐band and hence, myofibrillar contraction (Figure 7). Withdrawal of Ca2+
from the sarcoplasm ceases the muscle contraction. [247]
4.3.8. Role of Ca2+ in muscle contraction
The calcium ion is the main mediator of muscle contraction. Regulation of its
concentration and distribution is crucial to proper physiological functioning of the
muscle. The major Ca2+ regulating proteins in skeletal muscle include RYR1 and
DHPR, forming the voltage‐sensitive Ca2+ release complex, the Ca2+ reuptake SERCA
ATPases in the SR membrane and the Ca2+ storage protein calsequestrin in the SR
lumen [247].
36 |
4.3.8.1. The ryanodine receptors
Three genes encode the sarcoplamic reticulum (SR) Ca2+ release channel ryanodine
receptor, showing tissue‐specific pattern of expression. RYR1 is the major isoform in
skeletal muscle, whereas RYR2 is mainly expressed in cardiac muscle and brain, and
RYR3 is a minor isoform in brain [91]. Moreover, RYR1 expression undergoes a
developmental switch from a fetal isoform, lacking exons 70 and 83 [ASI(‐) and ASII(‐
)], to an adult isoform containing both exons postnatally [91, 225]. In DM1, RYR1
exon 70 is aberrantly spliced, resulting in the ASI(‐) isoform [134]. The ASI domain is
part of the RYR1 inhibitory domain [132]. Functional studies using myotubes have
indicated that the adult ASI(+) isoform shows reduced RYR1 channel activity in
comparison to the fetal isoform [134]. In parallel with this, the expression of the
fetal ASI(‐) isoform increased open probability, and resulted in enhanced E‐C
coupling [133]. Mutations in RYR1 cause malignant hyperthermia and central core
disease [163].
4.3.8.2. The SERCA family of Ca2+ re‐uptake channels
The sarcoplasmic/endoplasmic reticulum Ca2+ reuptake pump SERCA family
members are also encoded by three homologous genes, which go through tissue‐
dependent and developmentally regulated alternative splicing in their 3´ termini [44,
166]. SERCA1 isoform SERCA1a (ATP2A1A), which includes the differentially spliced
exon 22, is mainly expressed in adult fast fibers. In immature muscle and myotubes,
the embryonic SERCA1b isoform predominates, lacking exon 22 [39, 40]. The
functional difference between the developmental and adult isoforms is not currently
known. SERCA2a (ATP2A2A) predominates in adult slow muscle fibers and cardiac
muscle [330].
SERCA1 activity is inhibited by a small regulatory protein, sarcolipin,
presumably by decreasing its affinity for Ca2+ [205]. Similarly, phospholamban
modulates SERCA2 activity [275].
| 37
Mutations in SERCA1 are the cause of Brody disease, which presents impaired
muscle relaxation [128, 206].
4.3.8.3. The triad and calsequestrin
Efficient E‐C coupling requires stable and functional triad structures. Triadin (TRDN)
is an integral junctional SR membrane protein and a central structural protein in
triads, as is junctin [23]. Associated junctophilin (JPH1) facilitates the interaction
between DHPR and RYR [93].
Calsequestrin (CASQ) is the major Ca2+ binding protein in the muscle
sarcoplasmic reticulum (SR), with high capacity and low affinity for the Ca2+ ion [24].
CASQ in striated muscle is encoded by two different genes. CASQ1 encodes the
skeletal/fast isoform expressed in both fast and slow fibers, whereas CASQ2 encodes
the cardiac/slow form, which is the minor isoform in skeletal slow fibers [29, 62].
CASQ has binding sites for TRDN, junctin and RYR, and it recruits Ca2+ to
the junctional membrane, in the vicinity of RYR, keeping the free [Ca2+] low in SR.
Also TRDN binds to calsequestrin with its highly charged lumenal domain [137].
Instead of serving as a passive Ca2+ storage protein, CASQ modifies the function of
RYR in concert with TRDN and junctin, in a [Ca2+] dependent manner [23, 318].
CASQ1 has an inhibitory effect on RYR1 under physiological conditions ([Ca2+] 1mM).
When luminal Ca2+ increases, CASQ1 forms a supercompacted polymer, losing its
interaction with triadin and junctin. There is evidence indicating that the fast and
slow isoforms of CASQ regulate RYR isoforms in different ways in heart and skeletal
muscles. Similarly, CASQ1 and CASQ2 may well have distinct effects on RYR1 in fast‐
twitch and slow‐twitch muscles [216, 317].
4.3.9. Role of Ca2+ in muscle signaling
In addition to regulating muscle contraction, Ca2+ is also an important signaling
molecule [26, 222]. Many myogenic transcription factors, including the members of
38 |
the MEF2 and nuclear factor of activated T‐cell (NFAT) families, respond to multiple
Ca2+‐regulated signals [47, 179].
In addition to regulating muscle development, the MEF2 family of
transcription factors has important roles in adult muscle. MEF2C promotes a slow
fiber phenotype, and also has a crucial role in maintaining sarcomeric integrity, as
the expression of several muscle structural genes is dependent on it [229]. MEF2A
has been shown to promote a fast fiber phenotype, by increasing the activity of the
MYH2 (MyHC‐IIa) promoter [2]; in addition, MEF2A may increase the expression of
neonatal myosin (MyHC‐pn) [240].
Fiber type plasticity in adult muscle is mainly affected by contractile
activity and nerve stimuli. The amplitude and duration of the [Ca2+] transients
determine which fiber type specific transcriptional programs are activated. Several
mechanisms mediate this activation, including the calcineurin (Ca2+/calmodulin‐
dependent protein Ser/Thr phosphatase) and CaMK (Ca2+/calmodulin‐dependent
kinases) signaling pathways [196].
The major downstream effectors of calcineurin are members of the
NFAT family of transcription factors [47]. Dephosphorylation of NFATs by calcineurin
induces their activation and nuclear translocation. Calcineurin has been strongly
implicated in establishing and maintaining the slow fiber phenotype [47, 196, 272],
but may also mediate fast fiber‐specific transformations [2]. Different combinations
of activated NFAT family members are capable of inducing distinct MYH genes [47].
In addition, calcineurin is a modulator of skeletal muscle hypertrophy of both slow
and fast fibers [183].
The resting Ca2+ gradient between the extracellular space and
sarcoplasm is huge, 1000:1. Free Ca2+ is toxic to cells, and its strict regulation is very
important. Elevated cytoplasmic Ca2+ concentrations have been shown in cultured
DM1 myotubes [117], suggested to be due to abnormal Ca2+ influx through
(nifedipine‐sensitive) L‐type Ca2+ channels under resting conditions. Ca2+ is also
| 39
believed to play a significant role in dystrophinopathies and other dystrophies
caused by deficient DAG‐complex. In these disorders loss of sarcolemmal integrity
leads to leaking of Ca2+ inside the cells, causing damage and eventually muscle cell
necrosis [71].
4.3.10. The muscle biopsy
In order to prevent degradation, muscle biopsies are immediately snap frozen, for
example in liquid nitrogen‐cooled isopentane, which results in rapid freezing of the
specimen without considerable formation of ice crystals. For histochemical staining,
the tissue is embedded in supporting material (“tissue glue”) prior to freezing, and
sectioned on slides using a cryostat microtome [71]. Frozen material is suitable for a
variety of analyses, including enzyme histochemistry, immunohistochemistry (IHC);
in situ ‐hybridization (ISH); as well as RNA and protein extraction.
For diagnostic purposes, a multitude of histochemical staining can be
performed. Only the most common staining approaches are described here, which
are usually the most informative in any given myopathy. Haematoxylin & eosin (HE)
is the most widely used staining to reveal general tissue morphology. Haematoxylin
stains the acidic (basofilic) molecules, such as the nucleic acids blue, also cytoplasm
of regenerating fibers (“basofilic fibers”) appear blue due to large amount of RNA
molecules. Eosin stains muscle fibers and connective tissue red‐pink. The modified
Gomori trichrome method [77] stains the mitochondria red, resulting in darker
staining of the slow oxidative fibers, and reveals the presence of nemaline rods and
rimmed vacuoles, both red (Figure 8). The enzyme histochemical stainings for the
mitochondrial oxidative enzymes Cytochrome C‐oxidase (COX) and succinate
dehydrogenase (SDH) are often combined, where COX‐negative fibers appear blue,
and the COX‐positive fibers brown. The reduced nicotinamide adenine dinucleotide
dehydrogenase –tetrazolium reductase (NADH‐TR) staining also resolves the
localization of mitochondria, the sarcoplasmic reticulum (SR) is revealed, and
distortions in the myofibrillar structure are visible [71]. The ATPase method, which is
based on the different pH optimums of the ATPase domains in the distinct myosin
40 |
heavy chain isoforms [248], has served as the standard method for fiber type
differentiation [71]. However, immunohistochemical methods offer a more rapid
and reliable means for fiber type differentiation, and the role of MyHC IHC is
increasing in many laboratories [71, 233].
4.4. The muscular dystrophies
The muscular dystrophies (MDs) are a heterogeneous group of genetic disorders
caused by progressive loss of muscle cells, resulting in muscle weakness and wasting.
They show considerable variation in clinical manifestation and severity [127].
Table 2 shows a simplified classification of the neuromuscular diseases.
They can be divided into 1) myopathies, which are the primary diseases of the
muscle fiber, 2) myasthenias, caused by defects of the the neuromuscular junction
and 3) neurogenic disorders, caused by nerve disease or injury. The disease
etiologies in different MDs are divergent, although they all result in variable extents
of death of the muscle cells. An interesting general feature particular to the MDs is
that usually the involvement of different skeletal muscles is selective, and the
pattern is rather typical of the specific disease. This is important because it indicates
that some muscles are able to escape the effects of the mutation, reflecting the
biochemical and molecular diversity of the different muscles. [127]
Figure 8. Gomori trichrome staining showing granular degenerative changes corresponding to minirods (arrows) in atrophic muscle fibers of a patient with progressive muscular dystrophy of unknown cause.
| 41
Cardiomyopathies affect primarily the heart muscle. However, certain
skeletal muscular dystrophies, such as dystrophinopathies, are sometimes
accompanied with cardiomyopathy. In myotonic dystrophies cardiac arrhythmias
and conduction defects are encountered to variable extent, being more severe in
DM1.
The myotonic dystrophies, the object of this research, are major forms
of muscular dystrophy. Myotonia means delayed muscle relaxation after voluntary
contraction, which typically manifests as painless muscle stiffness. The major
primary myotonic disorders include the nondystrophic skeletal muscle
channelopathies and the myotonic dystrophies DM1 and DM2. The channelopathies
are caused in most cases by mutations in the chloride channel (CLCN1; e.g. myotonia
congenita) or the sodium channel (SCN4A; paramyotonia congenita, periodic
paralysis) encoding genes [250]. The myotonic dystrophies are classified as muscular
dystrophies, even though they differ from the other myotonic disorders by showing
unique variability in clinical presentation, and multisystemic involvement.
The prevalence of the neuromuscular disorders altogether in Finland is
estimated to be 200/105, comprising more than 10,000 patients [299]. The most
common genetic myopathies in Finland are the tibial muscular dystrophy (TMD), a
distal myopathy which belongs to the Finnish disease heritage (approximately 500
confirmed, symptomatic patients, plus an equal number of still undiagnosed patients
are estimated), and the myotonic dystrophies DM1 and DM2 (see chapter 4.4.2.1.)
[299].
4.4.1. Repeat expansions and disease
The myotonic dystrophies belong to the group of microsatellite repeat
expansions, which are the cause of almost 20 neurological diseases, including the
first identified Fragile X syndrome [86] and Kennedy´s diasease (or spinal and bulbar
muscular atrophy, SBMA) [146], Huntington´s disease, oculopharyngeal myotonic
dystrophy, and a variety of spinocerebellar ataxias (SCA) [42, 211].
42 |
Table 2. Classification of neuromuscular diseases.
Myopathies Gene(s) Muscular dystrophies Myotonic dystrophy type 1 DMPK Myotonic dystrophy type 2 ZNF9 Duchenne muscular dystrophy (DMD) DMD Becker muscular dystrophy (BMD) DMD Limb-girdle muscular dystrophies (LGMD) > 20 genes Facioscapulohumeral muscular dystrophy (FSHD) D4Z4
deletion in 4q35
Emery-Dreifuss muscular dystrophies (EDMDs) EMD, LMNA, SYNE1/2
Distal myopathies several E.g. Tibial muscular dystrophy (TMD) TTN Congenital muscular dystrophies (CMD) several Oculopharyngeal muscular dystrophies (OPM,
OPDM) PABPN1
Ion channelopathies Chloride channelopathies CLCN1 Sodium channelopathies SCN4A Inflammatory myopathies Mitochondrial myopathies Congenital myopathies several Metabolic myopathies Secondary myopathies (endocrinologic, toxic and amyloid myopathies) Myasthenias Myasthenia gravis Autoimmune diseases of the NMJ Congenital myasthenic syndromes Genes encoding synaptic proteins several Neurogenic disorders
Motoneuron diseases Amyotrophic lateral sclerosis (ALS) FALS genes Spinal muscular atrophy (SMA), recessive SMN1 SMA with severe respiratory distress (SMARD) IGHMBP2 Spinal and bulbar muscular atrophy (SBMA) AR Charcot-Marie-Tooth-disease > 35 genes Amyloid neuropathies TTR, GNS Inflammatory and toxic neuropathies
| 43
The majority of the mutations consist of repetitive trinucleotide sequences, the only
exceptions being DM2 with a (CCTG)n tetranucleotide, and SCA10 with a (ATTCT)n
pentanucleotide repeat expansion. Different pathomechanisms underlie these
diaseases; depending on several cis‐ and trans‐acting factors, they may cause protein
loss‐of‐function (e.g. Fragile X syndrome), protein gain‐of‐function (e.g. Kennedy´s
disease), or an RNA‐gain‐of‐function, which is the case in the myotonic dystrophies
DM1 and DM2, Fragile X‐associated tremor ataxia syndrome (FXTAS), and possibly
also in SCA8, SCA10 and SCA12. In these latter conditions, the mutation resides in
noncoding areas, which are transcribed into pre‐mRNA but not translated.
4.4.2. The myotonic dystrophies DM1 and DM2
4.4.2.1. History and prevalence
Myotonic dystrophies type 1 (DM1) and type 2 (DM2) comprise the most common
form of muscular dystrophy affecting adults. They are dominantly inherited diseases
caused by microsatellite repeat expansions, with clinical and molecular parallels.
DM1 was first described more than 100 years ago and its prevalence (12/105) is fairly
stable in European and North American populations [10, 106], even though
considerable regional variation occurs, probably due to a founder effect [329]. In
accordance with this, there are approximately 600 patients with symptomatic DM1
in Finland.
Before the DM2 mutation was identified, families with DM but lacking
the DM1 mutation were reported under different denominations: proximal myotonic
myopathy (PROMM) was first used by Ricker and colleagues [242, 243], followed by
several reports on European and North American families with similar clinical
features [115, 180, 195, 257, 260, 277]. A family in Finland was reported with
proximal myotonic dystrophy (PDM), presenting with more severe proximal muscle
wasting, but less myotonic features [300]. In 1998, a large Minnesota family was
described as having myotonic dystrophy type 2 (DM2), with proximal and distal
44 |
(hand) muscle weakness, when the disease was mapped to chromosome 3q21 [65,
239]. The DM2 mutation was subsequently found in 2001 [158].
In contrast to DM1, DM2 prevalence is not well established, due to its
later identification, late age of onset, milder clinical picture, and huge variation in
severity and symptoms [15, 301, 302]. Similar to DM1, DM2 occurs mostly in
populations of European descent [18, 157, 302], although a single Japanese patient
has been described with a distinct haplotype [251]. The minimum prevalence of DM2
was estimated to be 1/105 in European population [301], but it has since been shown
to be much higher. In central Finland, a prevalence of 16/105 has been established by
clinical ascertainment (Udd et al. unpublished results), and a recent study in the
Finnish population determines the frequency of the DM2 mutation at a surprisingly
high level of 53/105 in the general population (Suominen et al. unpublished results).
To date, DM2 has not been reported in sub‐Saharan populations.
Despite vast clinical variability, the myotonic dystrophies show less
genetic heterogeneity than was earlier considered. Most families first reported to
lack DM2 locus linkage were subsequently shown to have the DM2 mutation [142,
322] and one candidate family with frontotemporal dementia reported for a
potential DM3 locus on chromosome 15 was later found with a VCP mutation [152].
4.4.2.2. DM1 and DM2 mutations
The DM1 mutation was identified in 1992 as an expanded trinucleotide (CTG)n
repeat track in the 3´ untranslated (UTR) region of the dystrophia myotonica protein
kinase coding gene (DMPK; OMIM *605377) [41, 85, 167]. In DM1 patients the
repeat sizes range from 50 – 4,000 (150 – 12,000 bp), whereas normal individuals
have 5 – 37 repeats. Repeat lengths of 38 – 49 are considered a premutation allele
pool, showing decreased stability [9, 266]. The DM1 mutation length predicts the
clinical outcome to some extent: classical DM1 100 – 1,000 repeats; congenital
>2,000 repeats [9, 266].
| 45
The mutation causing DM2 was identified in 2001 in the first intron of
the zinc finger 9 (ZNF9) protein coding gene (ZNF9; OMIM *116955) [158]. The
mutation turned out to be an uninterrupted repeat expansion of the tetranucleotide
(CCTG)n, which is part of the complex polymorphic repeat motif
(TG)n(TCTG)n(CCTG)n(NCTG)n(CCTG)n [18, 158]. The normal alleles contain less than
30 copies of the (CCTG) repeat [158], while the range of expansion sizes in DM2
patients is huge; the smallest reported mutations vary between (CCTG)55‐75 [17, 158],
and the largest expansions have been measured to be approximately (CCTG)11,000
[158]. The mean repeat length is approximately 5,000, however, 10,000 repeats are
commonly encountered [17, 157]. The uninterrupted (CCTG)n repeat tract is less
stable than the normal interrupted form, and losing the interrupting sequences is
thought to predispose to the mutation. Because of the extreme somatic instability,
the threshold size of the disease‐causing mutation is under debate, however, it is
likely to be between repeat sizes (CCTG)30‐75. Bachinski et al. have reported large
uninterrupted alleles (CCTG)22‐33 in European population, which show elevated
instability, strongly suggesting that they represent the DM2 premutation allele pool,
with crude estimation of premutation allele frequency between 0.1 – 0.6 [17].
Interestingly, large uninterrupted (premutation) alleles, showing increased
instability, are more frequent in African Americans of sub‐Saharan origin [17].
Both DM1 and DM2 mutations consist of long uninterrupted repeat
tracks, which are thermodynamically less stable than normal alleles, showing a more
complex pattern with interrupting sequences. The tendency of a “dynamic” repeat
tract to contract or expand during replication depends on the primary sequence,
which determines its capacity to form secondary hairpin structures, the distance
from the replication origin, the flanking sequences, and the genomic location [42,
273]. Only the TG‐element contributes to hypervariability in interrupted alleles [17].
In other words, only uninterrupted alleles showed expansive variability in the (CCTG)
tract.
Both DM1 and DM2 expansion mutations show mitotic instability, with
variation in different tissues and cell types, causing somatic mosaicism [151, 190,
46 |
303, 325]. The DM1 and DM2 mutations show a tendency to grow longer during the
patient´s lifetime [158, 326]. However, there is a difference between the two DMs in
intergenerational passage of the mutation: the DM1 mutation tends to get larger in
successive generations, resulting in increased severity and earlier onset, a
phenomenon known as anticipation [11, 12, 13, 104, 151], whereas in DM2, the
mutation usually contracts when passed to the next generation, being shorter in
children [66]. This may explain some of the distinct features of DM2 phenotype, such
as the missing congenital form of the disease, the later onset of symptoms, and the
lack of anticipation [301], even though some implications of anticipation have been
postulated in some families [143, 264]. There is a gender bias in the tendency of a
repeat tract to expand or contract between generations in DM1, as the (CTG)DM1
repeat more often contracts after paternal transmission, and very large expansions
are usually encountered following maternal transmission. Interestingly, the mutation
length does not seem to correlate with the age of onset and phenotypic severity for
DM2 [18, 66, 158], in contrast to DM1 [102, 160]. However, it should be noted that
somatic variation is extensive, and the measured mutation repeat lengths are usually
derived from peripheral blood leucocytes. The repeat lengths in target tissues such
as muscle are shown to be much larger in DM1 [5, 288].
Fine mapping using microsatellite and SNP markers showed
conservation of a single common haplotype in a widespread DM2 patient cohort of
European descent. This data indicates a single founder DM2 mutation dating back
4,000 – 12,000 years ago (approximately 200 ‐ 540 generations) [18].
Primates (gorilla and chimpanzee) have slightly different repeat motifs
in intron 1 in ZNF9 still including the CCTG motif, whereas rodents (mouse and rat)
do not contain the CCTG repeat element at all [157, 158]. The normal function(s) of
the evolutionarily conserved noncoding repeat tract in the DM2 locus is not
currently known.
| 47
4.4.2.3. Symptoms
Both DM1 and DM2 are progressive multisystemic disorders with both shared and
distinct clinical features and pathomechanisms. The core symptoms that express to
varying degrees in DM2 include proximal muscle weakness, muscle pain, myotonia,
posterior subcapsular iridescent cataracts, tremors, cardiac conduction defects, and
endocrinological disturbances such as increased insulin resistance and male
hypogonadism. Laboratory findings include elevated creatine kinase (CK), hypo IgG,
positive autoimmune serology and elevated liver enzyme γ‐glutamyl transferase [66,
290, 301]. Many of the clinical manifestations are similar in classical adult onset
DM1, although with clearly distinct features including more severe weakness of
distal and facial muscles (including ptosis), leading to dysphagia, and respiratory
deficiency. The congenital DM1 form and the early childhood onset form with
mental retardation are not observed in DM2 [106, 298] (see Table 1. in publication
III, p. 467, for a summary of clinical features in DM1 and DM2.) However, the clinical
phenotype is widely variable in both forms, and even more so in the case of DM2.
DM2 may manifest with only a few symptoms, very late in the 7th or 8th decade and
there are no mandatory features common to all patients. There is also wide variation
in symptoms between different DM2 families and among individuals in the same
family [194, 195, 243]. DM2 patients have normal life expectancy and they do not
usually need help with daily activities, until after age 60, unless they suffer from
extensive pain. Musculoskeletal/myalgic pain may be the most disabling, or even the
only symptom in DM2 [15, 95]. In addition, several less frequent symptoms, or those
typical of a specific family, include excessive sweating and frontal balding [65], male
hypogonadism [300], and myotonia induced by pregnancy [203]. Sudden cardiac
deaths are rare but may account for DM2‐associated mortality [267]. Recently, it was
reported that DM2 patients suffer much more frequently from autoimmune diseases
than normal population, but such bias was not associated with DM1 [290]. In
contrast to DM1, no significant increase in complications associated with the use of
anesthesia has been encountered in DM2 [135, 319].
48 |
In magnetic resonance studies, brain white and grey matter changes, as
well as cerebral atrophy have been reported to be similar but less pronounced in
DM2 compared to DM1. However, these structural abnormalities have no clear
functional correlation, as severe cognitive impairment or extensive sleepiness is not
encountered with DM2 patients, in contrast to DM1 [129, 140, 188, 306]. However,
some CNS changes including avoidant personality trait [181] and impairment of
nonverbal episodic memory, which was linked to specific cerebral structural changes
[315] have been described also in DM2.
4.4.2.4. Muscle histopathology
Muscle histopathologies for DM1 and DM2 share some general common chronic
myopathic features, such as the high number of central nuclei and fiber size
variation, which were initially reported to be quite similar (Figure 9) [242, 287]. The
muscle pathology is progressive, with end‐stage pathology showing severe
reductions in fiber number and size accompanied with fatty and connective tissue
replacement in the most severely affected muscles [65, 106]. In DM1, ring fibers and
sarcoplasmic masses are more frequent, and type 1 fibers are generally more
affected [106].
In general, muscle wasting in DM2 is less prominent, and in accordance
with this, severe muscle fiber loss is not usually encountered. Muscle biopsies
typically demonstrate mild to moderate myopathic changes, in addition to more
severe fatty replacement and fibrosis starting in the interfascicular space; observed
only in proximal muscles of older patients with considerable muscle weakness
(Figure 9D). Small angulated fibers and atrophic nuclear clump fibers, similar to
those in neurogenic atrophy, are frequently found [66].
| 49
No clear correlation has been observed between the pattern of muscle
histopathological findings and the clinical muscle phenotype; including myotonia,
cramps, weakness and myalgia, or the order of onset of symptoms [268]. Electron
microscopic studies have not revealed any specific changes [243], even the
ribonuclear inclusions showed no distinct structure in EM [301].
Figure 9. Panel showing muscle histopathological findings in DM1 (A, B) and DM2 (C, D). Cryosections were stained with Hematoxylin & Eosin. Fiber size variation is present in all specimens, as are nuclear clump fibers (black arrows), and internalized nuclei (white arrows). (A) DM1 VL muscle showing relatively mild pathology; (B) DM1 TA muscle showing severe end stage pathology, with prominent fibrosis (Fib) and fatty degeneration (F). (C) DM2 VL muscle with early changes, including marked fiber size variation and abundant nuclear clumps. (B), Blood vessels. (D) DM2 VL muscle with late pathology and muscle weakness, some fatty replacement is seen (F), and remnants of a necrotic fiber (N). Note marked hypertrophy of the spared fibers.
50 |
4.4.2.5. Diagnostics
The gold standard for DM1 and DM2 diagnostics is mutation verification by genetic
testing as is seen for all genetic diseases with identified mutations. In the case of
DM1, the symptoms and family history are often clear and distinctive enough to
make a clinical diagnosis, and the mutation can be confirmed with direct PCR across
the mutation locus (< 200 CTG‐repeats), or with Southern analysis for repeats larger
than 100 repeats [9, 41, 167]. However, for DM2, the symptoms show such great
variability that a clinical diagnosis is most often difficult to achieve. Also the
molecular diagnosis is more challenging in DM2 due to the extremely large size
(mean approximately 5,000 CCTG repeats) and extensive somatic instability [18, 66].
Standard molecular genetic methods, such as Southern analysis and PCR, are not
sufficient for definite mutation verification. Hence, developing reliable and
standardized genetic tests for the mutation confirmation was challenging. A
recommended three‐step method for DM2 mutation testing [301] can achieve
unequivocal verification of the mutation: (I) A conventional PCR assay across the
mutation locus using probes binding to mutation flanking sequences can be used for
mutation exclusion. In all DM2 patients, a single PCR product representing the
normal allele can be identified because the DNA polymerase fails to amplify the
mutant allele due to its length and stable secondary structures. Two differently sized
normal alleles exclude DM2 [158]. Due to the fact that approximately 10 % of normal
population carry, by chance, alleles of exactly similar lengths, this method produces
a large number of false positives. (II) The sensitivity of a genomic Southern analysis is
70‐80%, resulting in high rate of false negatives [66]. (III) Repeat‐primed PCR‐
methods (RP‐PCR), using an internal probe with hanging tail, offer a powerful tool for
DM2 mutation verification, with 99% accuracy [66]. Several alternative, highly
specific and sensitive methods have been developed in addition to the RP‐PCR. A
modified Southern method using field‐inversion electrophoresis (FIGE) is particularly
efficient in determining the mutation length, in addition to qualitative mutation
verification [18]; likewise high sensitivity (reported 98%) for mutation identification
can be achieved using pulsed‐field gel electrophoresis [119]. Other novel Southern‐
based methods have been developed, for example utilizing HCl‐treatment of gels for
| 51
more efficient transfer of mutant DNA onto a blotting membrane [51], or repeat
expansion‐specific locked nucleic acid probes, in combination with rolling circle
amplification of short expansions, for increased detection sensitivity [199, 232].
Additional methods developed for the DM2 mutation verification include long‐range
PCR, [34] and a tetraplet‐primed PCR [52].
The combination of Southern analysis and one of the “third step”
methods should verify the DM2 mutation unequivocally [232]. Currently, several
European laboratories use either modified Southern‐ or RP‐PCR‐based methods for
DM2 mutation verification, and exchange DNA samples when needed for quality
control purposes [302].
Interestingly, it has been reported that currently diagnosed DM2
patients co‐segregate a heterozygous recessive CLCN1 mutation more often than
control population (5% versus 1.6%), indicating a selection bias to DM2 molecular
diagnostics with these patients [280]. No such bias was observed in DM1 patients.
In the reported family studies, the penetration of DM2 mutation was
100% [15, 300]. However, these families were selected for inclusion in the linkage
studies and represent the most prominent clinical manifestations of DM2. Later
experience has shown that in many families with very late onset symptoms in the
probands, the very old parents have not necessarily been identified to have
neuromuscular or other symptoms related to DM2. When asked for symptoms in the
family the response will thus frequently be negative, even if later work‐up may
identify some proximal muscle weakness or cataracts difficult to separate from aging
(Udd et al., unpublished results).
4.4.2.6. Pathomechanisms
The main molecular pathomechanism underlying the myotonic dystrophies is
dominant RNA gain‐of‐function [64, 153, 158, 173]. The expansion mutation, (CCTG)n
in DM2 and (CTG)n in DM1, is transcribed but not translated, and the mutant RNA
52 |
itself is necessary and sufficient to induce the molecular events leading to clinical
symptoms. The mutant repeat RNA species are retained in the nucleus, where they
aggregate and sequester RNA‐binding proteins, forming the ribonuclear inclusions
typical of DM1 and DM2 [158, 283]. According to the present body of knowledge,
the mutant RNA exerts its toxic effect by disturbing the distribution and function,
and hence the equilibrium between the two antagonistic splicing factors, MBNL1
(muscleblind‐like 1) and CUGBP1 (CUG‐binding protein 1), resulting in aberrant
mRNA splicing and translation [153]. According to recent reports, there are
additional, MBNL/CUGBP1‐independent pathomechanisms, involving dysregulation
of splicing, transcription, translation, and protein turnover [114, 124, 254, 296, 308].
The key pathomechanisms in DM are summarized in Figure 10.
Figure 10. Key pathomechanisms in myotonic dystrophies.
| 53
MBNL and CUGBP1 both act as splicing regulators, often promoting
opposite splicing events in their mutual target genes, although their binding sites are
distinct [110, 124]. Both regulate multiple alternative splicing events. The important
downstream effect of the dysregulation of these two antagonistic splicing factors is
the aberrant splicing in myotonic dystrophies, resulting in inappropriate embryonic
splicing pattern, which is considered one of the main causes of direct symptoms. The
aberrant splicing events identified in DMs are summarized in Table 3. Of these, at
least chloride channel 1 (CLC1) has been linked to myotonia [53, 172, 320]; insulin
receptor (INSR) to insulin resistance [262]; and cardiac troponin T (TNNT2) to cardiac
problems [224].
4.4.2.6.1. The muscleblind proteins
The Drosophila muscleblind gene encodes a zinc finger motif‐containing homeobox
protein required for terminal differentiation of muscle and eye [8, 25]. In human,
three genes (MBNL1, MBNL2, and MBNL3) encode the muscleblind‐like proteins, of
which MBNL1 is the major isoform expressed in skeletal and heart muscle [78].
MBNL2 is ubiquitously expressed, while MBNL3 is only found in
placenta. MBNL1 binds to dsRNA hairpin structure, and its binding is proportional to
the size of the expanded repeat [187]. All MBNL isoforms are capable of binding the
nuclear foci [78], and are able to cause alternative splicing of TNNT2 and INSR
minigenes [110]. However, only MBNL1 has been consistently implicated in DM
pathophysiology [156], whilst MBNL2 may play some role [103]. In fact, a
comparative study of HSALR and Mbnl1 –knockout mice indicated that majority (up
to 80%) of the abnormal splicing could be due to decreased Mbnl1 levels [70]. In
normal postnatal development, MBNL1 is predominantly translocated from
cytoplasm to nucleus [156], where it controls the temporally regulated
transcriptional switch from embryonic to adult isoforms, through alternative splicing
of certain exons [156].
.
54 |
Table 3. Abnormally spliced genes in myotonic dystrophies. Protein Gene Splice event Tissue DM1 DM2 Ref Cardiac troponin T (TNNT2) TNNT2 Exon 5 retention Heart muscle x [224]
Insulin receptor (IR) INSR Exon 11 exclusion Sk muscle x [262]
Insulin receptor (IR) INSR Exon 11 exclusion Sk muscle x [263]
Tau (MAPT) MAPT Exon 2 exclusion Brain x [271]
Tau (MAPT) MAPT Exons 2/10 exclusion Brain x [121]
Tau (MAPT) MAPT Exon 6c exclusion/exon 6d
retention Brain x [154]
Tau (MAPT) MAPT Exons 2/3 exclusion Brain x x [177]
Chloride channel 1 CLCN1 Exon 7a retention Sk muscle x x [53, 172]
Chloride channel 1 CLCN1 Intron 2 retention Sk muscle x x [53, 172]
Myotubularin-related protein 1 (MTMR1)
MTMR1 Exons 2.1/2.3
exclusion Sk/heart muscle x [43]
Fast sk troponin T (fTnT) TNNT3 Fetal (F) exon inclusion Sk muscle x [126]
Fast sk troponin T (fTnT) TNNT3 Fetal (F) exon inclusion Sk muscle x x [255]
N-methyl-D-Asp receptor (NMDAR1)
NMDAR1 Exon 5 retention Brain x [121]
Amyloid precurson protein (APP)
APP Exon 7 exclusion Brain x [121]
Ryanodine receptor 1 (RYR1) RYR1 Exon 70 exclusion Sk muscle x [134]
SERCA1 Ca2+ ATPase ATP2A1 Exon 22 exclusion Sk muscle x [134]
SERCA2 Ca2+ ATPase ATP2A2 Intron 19 retention Sk muscle x [134]
ZASP LDB3 Exon 7 inclusion Sk muscle x x [156]
ZASP LDB3 Exon 7 inclusion Heart muscle x [170]
Titin (TTN) TTN Exons Zr4/Zr5/Mex5 retention Sk muscle x x [156]
Calpain3 (CAPN3) CAPN3 Exon 16 exclusion Sk muscle x x [156]
GFAT1 GFAT1 Exon 10 exclusion Sk muscle x [156]
MBNL1 MBNL1 Exon 7 retention Sk muscle x x [156]
Dystrophin (DYS) DMD Exons 71/78 exclusion Sk/heart muscle x [200]
Alpha-dystrobrevin (DTNA) DTNA Exons 11A/12 retention Sk/heart muscle x [201]
Ankyrin-2 ANK2 Exon 21 exclusion Sk muscle x [210]
F-actin capping protein beta CAPZB Exon 8 exclusion Sk muscle x [210]
Zinc finger protein 9 (ZNF9) ZNF9 Intron 1/3 partial retention Sk muscle x [234]
| 55
The binding sites for MBNL1 or CUGBP1 in some mRNA species are known, and the
molecular effect on splicing already discovered. For example, MBNL1 binds to TNNT2
intron upstream of exon 5 and blocks access of the splicing factor U2AF65 promoting
skipping of TNNT2 exon 5 [110]. In DM1 and DM2, the concentration of soluble
nuclear MBNL1 is reduced due to its sequestration in ribonuclear foci, resulting in
loss‐of‐function and incomplete switch to adult isoforms [156]. Interestingly, most
aberrant splicing events described in DMs involve the expression of fetal or
embryonic isoforms.
4.4.2.6.2. CUGBP1
Specifically in DM1, an additional feature of CUGBP1 protein up‐regulation is
observed [262, 297]. CUGBP1 belongs to the family of CELF (CUGBP and ETR‐3 like
factors) proteins. Under normal conditions, CUGBP1 is upregulated during muscle
differentiation, where it has regulatory functions on both pre‐mRNA splicing [295]
and translation [296]. In DM1, the steady‐state CUGBP1 protein levels are also
elevated in adult muscle, due to protein kinase C (PKC)‐mediated
hyperphosphorylation, which increases the half‐life of CUGBP1 [145], resulting in
increased activity. However, the exact mechanism of this activation is not known.
Also other kinases are known to phosphorylate CUGBP1, such as Akt kinase at Ser28
[253], and cyclin D3/cdk4/6 at Ser302 [294]. Developmentally regulated
phosphorylation of the different sites in CUGBP1 directs its affinity to different target
mRNA species.
Unlike MBNL1, CUGBP1 specifically binds ssRNA, and it does not co‐
localize with the ribonuclear foci, even though there is evidence that it binds to the
stem of RNA hairpin structures in DM1 [182]. However, it is increased in both
nucleus and cytoplasm in DM1, where it associates predominantly with the soluble
fraction of the mutant (CUG)n transcript [297]. Interestingly, only mouse models
expressing the expanded (CTG)n repeat within its native genetic context, the DMPK
3´‐UTR, show CUGBP1 upregulation, suggesting that the mutation flanking
sequences have a critical role in CUGBP1 activation [210]. In agreement with this,
56 |
CUGBP1 shows no dysregulation in animal models, which express expanded (CTG)n
from an unrelated sequence [173], or in DM2, with one exception: CUGBP1‐eIF2‐
complex was found to be increased in DM2 patient myoblasts, and to interact with
(CCUG) repeats [254].
CUGBP1 was first implicated in splicing dysregulation of TNNT2 in DM1
[224]. It has been linked to numerous splicing events since. In addition to splicing,
CUGBP1 has other functions involved in translation regulation and mRNA stability
[296, 297]. CUGBP1 is required for proper myocyte differentiation [295]: MEF2A
(myogenic enhancer factor 2A) and p21 are translational targets of CUGBP1 [295,
296]. By increasing their translation, and subsequent elevation of other proteins,
such as myogenin, CUGBP1 may inhibit myogenesis and delay muscle development
in DM1 [296]. CUGBP1 may also stabilize its target mRNA species by binding AREs
(AU‐rich elements), or mediate mRNA decay of short‐lived transcripts by interaction
with GU‐rich elements [192, 308, 331].
4.4.2.6.3. Role of DMPK in DM1
Soon after the discovery of the DM1 mutation, decreased levels of both DMPK
mRNA and protein were reported in muscle of DM1 patients, suggesting that DMPK
is directly involved in DM1 pathophysiology [84]. Accordingly, knockout mice lacking
Dmpk show progressive late‐onset myopathy [84], and heart conduction
abnormalities [27]. In DM1, the repeat expansion is incompletely cleaved from the
DMPK pre‐mRNA, leading to both viable and aberrant DMPK mRNA species. A large
proportion of the (CUG)DM1‐containing mutant transcript is retained in the nucleus as
the ribonuclear foci [63]; however, it is also to some extent exported from the
nucleus to the cytoplasm [283, 297]. Defects in the posttranslational processing of
the mutant DMPK transcript, and its nuclear retention as a constituent of the nuclear
foci, cause lower DMPK protein levels and hence, haploinsufficiency in DM1 [141]
[63].
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4.4.2.6.4. Role of ZNF9 in DM2
Originally, the role of ZNF9 in DM2 pathogenesis was thought to be less relevant,
based on studies with cell culture models, which failed to indicate any effect of the
(CCTG)DM2 expansion on ZNF9 mRNA or protein expression [37, 175]. However,
recently accumulating data indicates that these models may have been insufficient
for such conclusions. Chen et al. (2007) reported that Znf9 haploinsufficiency in Znf+/‐
mice produces a multiorgan phenotype reminiscent of myotonic dystrophy, including
muscle weakness, cataracts, myotonic discharges, and cardiac conduction defects
[55]. Subsequently, reduction of ZNF9 protein was reported in DM2 myoblasts and
muscle biopsies [114, 219]. Later, it was shown that both ZNF9 mRNA and protein
are reduced in DM2, and that the mechanism underlying this is the impaired splicing
of ZNF9, due to the mutant (CCUG)DM2 sequence in the first intron. In the same
study, altered subcellular localization of ZNF9 in DM2 muscle was reported [234].
ZNF9 is expressed in a variety of tissues, most abundantly in heart and
skeletal muscle [158]. ZNF9 contains seven zinc‐finger (CCHC) motifs common to
transcription factors and ribosomal proteins, and its protein structure is highly
conserved, suggestive of an important biological role [81, 312]. Znf9‐/‐ knockout mice
are embryonic lethal due to defective forebrain development [56]. ZNF9 binds to
several gene regulatory elements [80, 185, 235], and it has been suggested to be
involved in a variety of transcription and splicing events. However, accumulating
evidence is pointing to translation as the major regulatory role for ZNF9: it has been
shown to regulate both cap‐dependent and cap‐independent translation initiation
[96, 220], as well as internal ribosome entry site sequence (IRES) –mediated
translation, the activation of which was reduced in DM2 [256]. Importantly, ZNF9
binds to terminal oligo‐pyrimidine tract (TOP) containing mRNAs, including those
encoding ribosomal proteins and elongation factors. This has been specifically
indicated in DM2 pathomechanisms, as down‐regulation of ZNF9 reduces the global
protein synthesis in DM2 [114].
58 |
4.4.2.6.5. The nuclear foci
Despite the nuclear foci being a characteristic finding in myotonic dystrophies, there
is evidence that the ribonuclear inclusions are neither toxic nor mandatory for the
disease pathogenesis. They could even be protective, as a mouse model
overexpressing the normal DMPK 3´ UTR mRNA, containing (CTG)5, developed many
of the core features seen in DM1, including myotonia, cardiac conduction defects,
and defective splicing [168]. This is in agreement with a Drosophila model with
nuclear foci formation but no phenotype [112]; the same study suggested that
MBNL1 may, in fact, stabilize the CTG‐repeat secondary structures, and hence, the
nuclear foci [112]. Nuclear foci in DM2 muscle cells are generally larger compared to
foci in DM1, possibly due to larger repeats in DM2 and/or more abundant expression
of ZNF9 versus DMPK [171].
4.4.2.6.6. Neighboring genes
Neighboring genes may well modify pathophysiology in DMs, which further
complicates the molecular pathomechanisms and clinical outcome. Six5 knockout
mice develop cataracts, even in the heterozygous state (although without a muscle
phenotype) [261]. Haploinsufficiency of SIX5 and DMWD have been implicated in
some DM1 patients, which may be due either to transcriptional defects, or failed
nuclear export of the transcripts [4, 136, 139, 289].
It has been suggested that the CTG repeat affects the chromatin
structure at the mutation locus (3´), making it more condensed and less accessible to
nucleases [213]. Subsequently, it was shown that this affects the transcription of the
neighboring genes [136, 289].
4.4.2.6.7. Other pathomechanisms
Another pathomechanism suggested in DM2 involves protein turnover; the catalytic
core subunits of the proteasome were identified as (CCUG)‐interacting proteins, and
| 59
inhibition of protein degradation, or increased stability of short‐lived proteins, was
observed in DM2 myoblasts [254].
Myogenic differentiation has been shown to be defective in DM1: the
satellite cells are able to proliferate normally, but they show delay in fusion and
differentiation [88]. A role of CUGBP1 has been implicated, as it impairs the
transcriptional activation of myogenic markers necessary for differentiation [219,
295]. In congenital DM1, the satellite cells are defective also in proliferation [90],
whereas in DM2, no abnormalities in myogenic program or satellite cells have been
described [50, 219].
4.4.2.7. Animal models
Several mouse models have been developed which display features of DM1, some of
the most important ones are described below. Interestingly, none of the models so
far have reproduced the multisystem disease completely. This may be due to several
reasons, including the mutation length and level of expression, whether the
mutation is expressed from its native genetic context, or from an unrelated
localization, the genetic background of the model animal species, and variation
between different strains. In addition, there is less somatic and intergenerational
instability of long repeat tracts in mouse. DM2 mouse models have not been
reported as yet.
Transgenic mice have been developed to carry human actin‐gene
modified to include a short repeat (CTG)5 or a long repeat (CTG)250, called HSASR and
HSALR, respectively [173]. The transgene was selectively expressed in skeletal muscle,
and the HSALR mice displayed aberrant splicing events similar to DM1 with developed
myotonia and mild histopathological alterations of the skeletal muscle, but not
weakness and wasting.
A French group generated a transgenic mouse carrying a large (>45 kD)
human genomic fragment containing DMPK with more than 300 CTG repeats, and
60 |
also the neighboring genes DMWD and SIX5 [273]. The repeat expansion in this
model showed both somatic and intergenerational instability, although the rate of
instability is generally lower in mice. The phenotypes were variable between strains
and also between individual mice in the same strain. Importantly, the mouse strains
with a phenotype showed myopathic changes, atrophy and myotonia, and findings in
the brain suggested similar tau pathology as encountered in DM1 patients [274].
Because of the well established role of MBNL1 in DM pathogenesis, a
MBNL1 knockout mouse was developed [126]. It showed a very similar phenotype as
the HSALR model, including splicing abnormalities, myotonia and mild muscle
pathology [126]. CUGBP1 overexpressing mouse models presented with delayed
skeletal muscle differentiation and muscular dystrophy, with translational
dysregulation [296], in addition to aberrant splicing in both heart and skeletal muscle
[109].
Subsequently, an adeno‐associated virus 2 (AAV2)‐vector mediated
inducible mouse model was described, harbouring a longer (CTG)960 repeat,
corresponding to its native locus in the 3´UTR of DMPK [210]. The construct was
expressed selectively in skeletal muscle. In addition to replicating the myotonia and
abnormal splicing of the HSALR and MBNL1 ko, this (CTG)960 mouse displayed
upregulation of CUGBP1 protein. The mice displayed severe muscle wasting and
weakness, along with dystrophic changes in muscle histopathology [210]. A mouse
model (EpA960/MCM) expressing a similar inducible (CTG)960 cassette, specifically in
heart, fully recapitulated the heart phenotype in DM1, including arrhythmias,
cardiomyopathy, and systolic/diastolic dysfunction, with a key molecular pathology
of nuclear foci formation and aberrant splicing [310].
Drosophila models for DM1 indicated that longer expansions are
needed for pathological effects in fly; a model expressing (CTG)162 from human
DMPK 3´‐UTR did develop ribonuclear foci, but no disease phenotype [112], whereas
expressing sole interrupted (CTG)480 without DMPK flanking sequences resulted in
foci formation, muscle wasting and eye phenotype [67].
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4.4.2.8. Management and gene therapy
There is no cure for myotonic dystrophies, however medication and physiotherapy
may relieve some of the symptoms. Cardiorespiratory disorders account for 70% of
the mortality in DM1, emphasizing the need for active monitoring and interventions
at some stage of the disease. Even though malignant cardiac arrhythmias are more
rare in DM2, it has been suggested that annual ECG should be also performed on
DM2 patients [302]. Early insertion of pacemakers in DM1 patients could prevent
some of the sudden deaths due to arrhythmia. Respiratory complications and apnea
in DM1 may require non‐invasive ventilation, and daytime sleepiness may be
relieved using CNS stimulant drugs [164, 298]. These measures are usually not
needed for DM2 patients.
Cataracts may require surgical removal. Gastrointestinal symptoms,
such as abdominal pain, dysphagia, vomiting and diarrhoea, are common in DM1
patients, and may also occur in DM2 patients [245, 246, 291, 292]. Some of the
symptoms can be relieved medically, however, there is no treatment for dysphagia
in DM1 except for parenteral feeding [298]. Myalgic pains in DM2 do not respond
well to conventional analgesics.
There have been some reports concerning the treatment of muscle
wasting in DM1, that suggest insulin‐like growth factor 1 (IGF1) therapy to improve
insulin insensitivity in DM1 myoblasts [87]. Treatment with creatine may moderately
improve muscle strength in DM2 patients, but not in DM1 patients [265]. Training
does not relieve muscle symptoms in DMs, but moderate exercise is not harmful,
and it may have other beneficial effects on patients [309]. Myotonia is often not a
major complaint, however, DM1 patients may be treated with mexiletine [161].
The continued search for molecular pathomechanisms in myotonic
dystrophies has enabled the development of gene and molecular therapy strategies
at several levels. These include (I) elimination of the expanded C/CUG RNAs; (II)
preventing harmful protein binding to the mutant RNA; (III) reversing the aberrant
62 |
splicing events [320]; (IV) forcing nuclear export of the mutant RNA; (V)
overexpression of MBNL1 [125]; (VI) restoring normal CUGBP1 levels by PKC
dephosphorylation [311]. Most attention has been directed to strategies designed
for specific targeting of the toxic expanded RNA [197]. Silencing of mutant (CUG)n
RNA can be achieved by several methods. Using ribozymes or artificial trans‐splicing
molecules, 3´part of the mutant transcript can be exchanged to reduce the number
of CTG repeats [54, 226]. Antisense strategies inducing degradation of mutant
transcripts have been applied by introducing complementary antisense RNA
molecules directed against DMPK 3´UTR. This resulted in partial reduction of
pathological effects in DM1 patient myoblasts [89]. Antisense oligonucleotides
(AONs), which induce the degradation of the DNA‐RNA hybrid by RNase H [144],
have been used successfully in both patient myoblasts and two different mouse
models for DM1 [198]. RNA interference pathways have also been utilized in order
to eliminate the formation of hairpin structures in mutant RNA, reminiscent of A‐
form RNA. Trials using synthetic RNA oligos and lentiviral vector‐delivered short
hairpin RNA targeting sequences resulted in reduction of mutant DMPK transcripts
[150]. Another approach to avoid the pathological effects of the mutant repeat RNA
is to inhibit detrimental protein binding to the hairpin structures formed by
(C/CUG)n. Use of morpholino‐type AONs, which bind to and inhibit activity of, but do
not induce degradation of the target RNA [279], resulted in release of MBNL1
protein from the expanded (CUG)n, and subsequent restoration of aberrant splicing
in HSALR mice [321]. Recently, a group of small molecules, including pentamidine
[313] have been described, which release MBNL1 from (C/CUG) repeats [7, 94, 313].
Several studies on DM1 mouse models suggest that the pathological
effects are reversible [106, 143] [311, 320], which makes development of gene
therapies that may be clinically useful, very important. In addition to overcoming
harmful side‐effects, developing an efficient means for delivering the active
compounds to the target tissues, which are versatile in DMs, continues to be a
challenge in DM molecular therapy.
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5. AIMS OF THE STUDY
At the beginning of this study, the DM2 mutation had just been discovered, and tools
for accurate mutation verification were not widely available. In addition, using
routine muscle histopathological methods, the muscle histopathology of DM2
showed mostly non‐specific, mild myopathic findings, similar to DM1, which offered
no possibility for differential diagnostics. The principal goal of this work was to
develop new methods for DM2 diagnostic purposes utilizing muscle biopsies
frequently taken from patients with undefined muscular symptoms. In the second
part of the study, the differences in mRNA and protein expression between DM1 and
DM2 were assessed in order to resolve the specific pathways involved in the
molecular pathology of the myotonic dystrophies, with special emphasis on the
differences between the two similar but distinct diseases.
The specific objectives in this study were:
1. To improve differential diagnostics of DM2 by
a. identifying specific differences in the muscle histopathology, for
screening of patients for eventual mutation verification
b. developing an accurate DM2 mutation verification method using
muscle biopsy material
2. To gain understanding of the defective molecular pathways underlying DM2
muscle pathology by
a. studying expression of muscle‐specific proteins using standard methods
b. studying mRNA expression of muscle‐specific genes using microarrays,
and differential splicing using splice variant analysis
64 |
6. MATERIALS AND METHODS
6.1. Patients and samples
Sixty‐two DM2 patients participated in this study (publications I‐IV). The majority of
them were Finnish (37) and the remaining patients originated from North America,
France, Spain, and Italy. All Finnish DM2 skeletal muscles used were vastus lateralis
(VL), whereas the foreign cohort consisted mostly of biceps brachii and deltoid
muscle biopsies.
Thirty DM1 patients from various countries and ethnic origins (Swedish,
French, North American, Spanish, and Finnish) served as controls. The muscles used
were tibialis anterior (TA), VL, biceps brachii and deltoid. Disease controls, with AR
congenital myotonia (CLCN1‐channelopathy) patient (I), TMD, BMD and DMD (III, IV)
were also included. In addition, more than 20 muscle biopsies from healthy controls
from Finland and North America were used.
In experiments where DM1 samples were analyzed to assess specific
differences in comparison to DM2, muscle biopsies matched by age, sex, muscle site,
and stage of pathology were preferably used. Both muscle biopsy and blood samples
were obtained from all patients and control individuals in this study.
6.1.1. Consent and ethics committee permissions
The majority of the DM2 biopsies had previously been taken for
diagnostic purposes. Enrollment of patients was approved by the local institutional
review boards, and the muscle biopsies and blood samples were used in this study
only after informed consent from the patients.
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6.1.2. Patients included in the histopathological study (I)
Nine biopsies from DM2 patients were included as follows: Finnish VL (3), French
deltoid (3), and Italian biceps (3). Biopsies from age‐matched French DM1 patients,
VL (3), deltoid (3), biceps (2), with mild pathology, and one recessive congenital
myotonia patient with a homozygous mutation in the CLCN1 gene served as controls.
6.1.3. Patients included in the study of new diagnostic methods (II)
Altogether 13 Finnish DM2 patients with previous molecular genetic diagnosis, and a
cohort of patients with an undefined (at the time of the study) neuromuscular
disorder (n = 31) served as subjects. Additionally, four healthy individuals, and three
DM1 patients were used as (negative) controls in CISH and FISH.
6.1.4. Patients included in the studies of sarcomeric and Ca2+ handling proteins and
mRNA expression (III, IV)
We used two distinct sample sets for the protein (set 1) and mRNA (set 2) expression
studies. In set 1 (publication III), 20 DM2 and 9 DM1 patients were included,
together with 2 healthy individuals and 1 TMD‐patient. Set 2 (III) was used for mRNA
expression profiling and splice variant analysis and consisted of 20 DM2, 10 DM1
patients and 6 healthy controls. In publication IV, the same samples were used, with
additional DM1 (n = 2) and healthy controls (n = 12) in set 1, and neuromuscular
disease controls with TMD (n = 1), BMD (n = 3) and DMD (n = 3) in set 2.
6.1.5. Patients with neuromuscular diseases included in the study of protein
expression in the highly atrophic fibers (U)
The same DM2 patients as in set 1 (III) were used for a broader protein expression
study in muscle samples by immunohistochemistry, including an additional 9 Finnish
patients with neurogenic disorders as disease controls: Charcot‐Marie‐Tooth disease
66 |
(1); amyotrophic lateral sclerosis (2); familial amyotrophic lateral sclerosis (3); lower
motor neuron syndrome (2); spinal and bulbar muscular atrophy (1).
6.2. Methods
The methods used in this study are summarized in Table 4. The methods are
described briefly herein; for details, please see the corresponding article where full
descriptions of methods can be found. In addition, Table 5 summarizes all primary
antibodies used in this study.
Table 4. Methods used in this study.
Method Study
Immunohistochemistry I, III, IV
Immunofluorescence III, IV
Enzyme histochemistry I
Muscle morphometry I
Bright and dark field microscopy I‐IV
Chromogenic in situ hybridization II
Fiber‐FISH II
PCR‐based allele sizing II
Repeat‐primed (RP‐)PCR II
Field‐inversion gel electrophoresis II
Western blotting III, IV
Microarray expression profiling III, IV
Splice variant analysis III, IV
Confocal microscopy IV
| 67
Table 5. Primary antibodies used in this study.
Antibody Gene Protein Clone Supplier Ref. WB-MHCf MYH2
(/MYH1) MyHC-IIa(/x) WB-MHCf NC [73]
A4.74 MYH2 MyHC-IIa A4.74 DSHB [316] MHCf(ast) MYH2
(/MYH1) MyHC-IIa(/x) MY-32 SA [107]
MHCs(low) MYH7 MyHC-beta/slow NOQ7.5.4D SA [69] MHC-beta MYH7 MyHC-beta/slow WB-MHCs NC -
MHCd MYH3 MyHC-emb RNMy2/9D2 NC [73] MHCn MYH8 MyHC-pn WB-MHCn NC [73]
SERCA-1 ATP2A1 SERCA1 VE121G9 RDI [204] SERCA2 ATP2A2 SERCA2 goat pAb SCB [184]
TropT TNNT3 fTnT T1/61 NC [30] NCAM NCAM1 NCAM-1 UJ13A DAKO [217] nNOS NOS1 NOS1 NOS-B1 SA [230] Bcl-2 BCL2 BCL2 124 DAKO [223]
Cleaved Casp-3 CASP3 CASP3 5A1 CS [118] Dystrophin C-
term. DMD DYS rabbit pAb NM
-
Myogenin MYOG Myogenin F12B SA [215] Vimentin VIM Vimentin 3B4 DAKO [33]
ZASP LDB3 ZASP mouse pAb G. F. [79] RyR RYR1/2 RYR1/2* 34C NB [123]
DHPR alpha-1 CACNA1S DHPR 1A NB [193] Triadin TRDN TRDN (pAb) SCB -
Junctophilin JPH1 JPH1 (pAb) ABN - CASQ1 N-16 CASQ1 CASQ1 goat pAb SCB -
CASQ2 CASQ2 CASQ2 rabbit pAb ABC [159] Calcineurin PPP3CA CaN CN-A1 SA [207]
NFATc3 NFATC3 NFATC3 mouse pAb ABN - Pax7 PAX7 PAX7 Pax7 DSHB [130]
Alpha-actinin ACTN2/3 ACTN2/3 EA-53 SA -
NC, Novocastra Laboratories, Newcastle Upon Tyne, UK; DSHB, Developmental Studies Hybridoma Bank, University of Iowa, Iowa City, IA, USA; SA, Sigma‐Aldrich, Saint Louis, MO, USA; RDI, RDI Division of Fitzgerald Industries Intl, North Acton, MA, USA; SCB, Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA.; DAKO, Glostrup, Denmark; CS, Cell Signaling Technology, Inc., Danvers, MA, USA; NM, NeoMarkers, Thermo Fisher Scientific, Inc., Fremont, CA, USA; G. F., ZASP mouse pAb was a generous gift from Dr. Georgine Faulkner, Trieste, Italy; NB, Novus Biologicals, Inc., Littleton, CA, USA; ABN, Abnova Corporation, Taipei, Taiwan; ABC, Abcam, Cambridge, UK. pAb, polyclonal antibody. *The RYR antibody is reported to recognize RYR1 and RYR2; however, we conclude that this antibody detects mainly the dominant RYR1 isoform in skeletal muscle, based on expression data and the immunohistochemical findings: localization to the SR junctional membranes with DHPR, and higher signal in the fast fibers.
In order to to verify that each antibody recognizes its intended target, they have been carefully tested in immunohistochemistry and/or Western blotting using healthy control muscle material, and compared the obtained results with the information provided by the Ab manufacturer and literature. We have confirmed that subcellular distribution (IHC) and molecular weight (WB) are in concordance with published data of each protein.
68 |
6.2.1. Immunohistochemistry (I, III, IV), immunofluorescence (III, IV) and microscopy (I‐IV)
All muscle biopsies used in this study were snap frozen in isopentane cooled with
liquid nitrogen, and stored at ‐80°C – ‐135°C until use. Either 6 or 10 μm sections
were cut on glass slides using a cryomicrotome, and the sections were stored frozen
at ‐80°C.
For DAB immunohistochemistry, either BenchMark (Ventana Medical
Systems, Tucson, AZ, USA) or TechMate (DakoCytomation, Glostrup, Denmark)
automated immunostainer was used, according to the manufacturer´s instructions.
Immunofluorescent stainings were performed manually using the following
procedure. The sections were fixed in 4% PFA for 10 min, at room temperature (RT),
or in methanol at ‐20°C for 2 min. The sections were then permeabilized using 0.2%
Triton‐X‐100, and blocked with 1% bovine serum albumin (BSA). The Ab incubations
were performed at RT for 1 – 2 h. The slides were rinsed with PBS, and mounted in
mounting medium containing an anti‐fade agent. The slides were viewed under a
light microscope, or using a Zeiss LSM 510 confocal microscope (Carl Zeiss, Inc.,
Göttingen, Germany).
6.2.2. Enzyme histochemistry (I)
The ATPase staining used for fiber type differentiation was performed by pre‐
incubating the slides in either alkaline Barbital‐Na (pH 10.4), or acidic Ba‐acetate (pH
4.3/pH 4.6). After rinsing the slides, ATP solution was added, followed by a substrate
solution. Counterstaining with Herovici was performed, and the slides were mounted
in HistoMountTM. [71]
6.2.3. Muscle morphometry (I)
Randomly selected regions on muscle sections stained with ATPase or MyHC‐IHC
were analyzed by measuring the “smallest diameter” [71]. Approximately 500 –
| 69
5,000 fibers were measured in each specimen, and the data was analyzed using the
Microcal Origin software (Microcal Software Inc., Northampton, MA, USA). Atrophy
and hypertrophy factors were calculated, and metahistograms were generated.
6.2.4. Chromogenic in situ‐hybridization (II)
Frozen muscle biopsy sections were prepared as in 2.1. In addition, some formalin‐
fixed (12 – 24 h), paraffin‐embedded sections were used. Digoxigenin end‐labeled
sense (CCTG)8 and antisense (CAGG)8 probes (synthesized by Sigma‐Genosys,
Cambridge, UK) complementary to the DM2 mutation were allowed to hybridize to
the target overnight at 37°C. The next day, the slides were rinsed with 1 x SSC at
45°C, and endogenous peroxidase was quenched using H2O2 in methanol. The bound
probe was visualized using Zymed Spot‐Light CISH Polymer Detection kit (Zymed
Laboratories, San Francisco, CA, USA), according to the manufacturer´s instructions.
The slides were counterstained with Gill II hematoxylin and mounted in
HistoMountTM solution. For the paraffin‐embedded sections, a similar procedure was
applied, with two additional steps in the beginning of the procedure: Spot‐Light
enzyme pretreatment was used, followed by heating in HIER pretreatment buffer
(both from Zymed).
6.2.5. Fiber‐FISH (II)
Peripheral blood lymphocytes were used to prepare extended DNA fibers on glass
microscope slides. The same digoxigenin‐labeled sense and antisense probes were
used for the mutation detection as in 2.4. In addition, BAC clone RP11‐814L21
(AC022944; BACPAC Resource Center) was nick translation labeled with biotin 16‐
dUTP (Roche Diagnostics GmbH), in order to detect the mutation flanking sequences.
The the RP11‐probe was denatured at 90°C for 5 min, after which the
(CCTG)8/(CAGG)8 probes were added, and the hybridization mix was applied on
slides. The hybridization was allowed to take place for 48 h at 37°C. The biotinylated
RP11‐probe was detected using TRITC‐conjugated avidin D followed by biotinylated
goat anti‐avidin D and another layer of TRITC‐conjugated avidin (Vector Laboratories,
70 |
Burlingame, CA, USA), in order to increase detection sensitivity. Mouse anti‐
digoxigenin (Roche Diagnostics, Mannheim, Germany) was used to detect the bound
(CCTG)8/CAGG)8 probes, followed by immunofluorescent detection with anti‐mouse
fluorescein. The slides were mounted in mounting medium containing an anti‐fade
agent. The mutation length was measured using the centromeric portion of the BAC
814L21 for calibration, which is known to be approximately 155 kb in length.
6.2.6. PCR‐based allele sizing (II)
The DM2 mutation locus was amplified by polymerase chain reaction (PCR) using
genomic DNA derived from blood leucocytes and primers CL3N58‐D F/R, which bind
to the mutation flanking sequences [158]. The resulting PRC‐products were
separated by one of the following methods. 1) 32P –dCTP labeled PCR‐products were
separated by polyacrylamide gel electrophoresis (PAGE), and detected from the gel
using autoradiography. 2) Fluorescently (Cy‐) labeled PCR‐products were separated
by electrophoresis on an ALFexpress Automated DNA Sequencer (Amersham
Pharmacia Biotech, Uppsala, Sweden), and the bands were analyzed using AlleleLinks
1.0 software (Amersham). 3) Fluorescently (FAM‐) labeled PCR‐products were
separated using capillary electrophoresis on an ABI 3100 Genetic Analyzer, and the
results were analyzed using the instrument´s software (Applied Biosystems, Foster
City, CA, USA).
6.2.7. Repeat‐primed PCR (RP‐PCR) (II)
A similar forward primer as in 2.6., but with an additional 5´ tail (5´‐GTTTCTT‐3´), was
used for RP‐PCR, together with a reverse primer consisting of (CAGG)5 and an
universal tail sequence. Genomic DNA was used for amplification of the DM2
mutation locus, and the resulting PCR products were analyzed using capillary
electrophoresis, ABI 3100 and Genotyper software (Applied Biosystems).
| 71
6.2.8. Field‐inversion gel electrophoresis (FIGE) (II)
Field‐inversion gel electrophoresis is a modified Southern blot analysis, particularly
suitable for detecting long repeat tracts. Genomic DNA was digested with EcoRI, and
the digested samples were separated in agarose gels on the FIGE Mapper (Bio‐Rad
Laboratories, Hercules, CA, USA). The samples were blotted on membranes for
detection with a 32P‐labeled probe recognizing the DM2 repeat expansion sequence,
and the membranes were analyzed using autogradiography.
6.2.9. Western blotting (III, IV)
Frozen muscle biopsies were used for Western blotting (WB) sample preparation by
mechanical homogenization in 20 – 40 fold (mg/µl) volume of sample buffer
containing a reducing agent (beta‐mercaptoethanol) and sodium dodecyl sulphate
(SDS) for polypeptide linearization, after which the samples were heated at 95°C for
5 min. Alternatively, to preserve and solubilize the fragile membrane proteins, the
muscle biopsies were sonicated 4 x 5 sec in lysis buffer containing a protease
inhibitors mix (Complete; Sigma‐Aldrich, Saint Louis, MO, USA), before adding
sample buffer, and heating at 35°C for 15 min. Usually 5 – 20 µl of samples,
depending on the protein to be detected, were separated electrophoretically using
SDS‐PAGE gels, and transferred onto PVDF membranes for detection. Primary and
secondary Ab incubations were performed for 1 – 2 h at RT, and enhanced
chemiluminescence (ECL) was used for the detection of bands. The obtained bands
were most often normalized to the Coomassie brilliant blue stained MyHC bands in
gels after blotting. Alternatively, ECL‐detected beta‐actin or alpha‐tubulin bands
were sometimes used.
6.2.10. Microarray expression profiling (III, IV)
Frozen muscle biopsies were homogenized, and the total RNA was extracted and
purified with RNeasy kit (Qiagen, Valencia, CA, USA). Using the RNA as template,
cDNA was synthesized using the Superscript II system (GIBCO/BRL). In vitro
72 |
transcription labeling was performed with biotinylated UTP and CTP (Enzo
Diagnostics, Farmingdale, NY, USA), and the labeled, amplified cRNA was fragmented
before applying on the whole human genome expression array U133Plus 2.0
GeneChip (Affymetrix, Santa Clara, CA, USA). After hybridization, the chips were
scanned according to the manufacturer´s protocol, and data analysis was performed
using the DChip software package (DNA‐Chip Analyzer).
6.2.11. Splice variant analysis (III, IV)
cDNA derived from muscle biopsies was also used for splice variant analysis (SVA). In
order to estimate the proportions of different splice isoforms in each sample,
reverse transcriptase‐PCR (RT‐PCR) was performed with fluorescently (FAM‐) labeled
universal primer. Specific primers for the exons flanking the alternatively spliced
exon were used. The RT‐PCR products were separated and analyzed using capillary
electrophoresis and ABI 3100 sequencer, and the resultant peak heights were added
together to obtain the total signal. Student´s t‐test was used to test the statistical
significance of the results.
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7. RESULTS AND DISCUSSION
7.1. Histopathology and protein expression in DM2 (I, III, IV, U)
Nine DM2 muscle biopsies originating from three European countries and obtained
from three different muscles (VL, deltoid, biceps brachii) showed rather non‐specific
myopathic changes when stained with routine histopathological procedures (HE,
Gomori´s trichrome and NADH): increased fiber size variation, including scattered
round or thin atrophic fibers, internal nuclei, and nuclear clump fibers (Table 6). The
findings were consistent with previously reported findings [243, 287]. However,
nuclear clump fibers, with and without pyknotic nuclei, occurred in most biopsies in
high numbers, also in clinically asymptomatic muscles. Infrequent findings included
increased connective tissue and fat replacement, necrotic fibers, moth‐eaten fibers,
rimmed vacuoles and ragged red fibers. On the other hand, ring fibers, targetoid
fibers, and sarcoplasmic masses, more common in DM1, were extremely rare.
When the enzyme histochemical ATPase method was used for fiber
type differentiation, the previously reported fiber type 1 predominance [66] was
observed in many DM2 specimens, particularly in VL muscles. However, when
immunohistochemistry was applied, using an antibody against the fast myosin heavy
chain, numerous very atrophic fibers (diameter ≤ 6 µm) were revealed (Figure 11A).
They were mostly scattered, without grouping, as in neurogenic atrophy. In contrast
to the nuclear clump fibers, the smallest population simply showed sarcolemma and
a scarce amount of sarcoplasm with or without nuclei. ATPase staining was not
sensitive enough to visualize the majority of these very small diameter fibers (Figure
11C). IHC using an Ab against type 1 (slow) fibers did neither recognize such highly
atrophic fibers, nor the nuclear clumps, confirming them solely as fast type in DM2.
Subsequently, we verified that these mini‐fibers in DM2 represent the fast subtype
2A (III) [233]. The presence of type 2B fibers (expressing MyHC‐IIx) in DM2 was also
confirmed, as well as an increased number of 2C hybrid fibers expressing both fast
(MyHC‐IIa) and slow (MyHC‐I) myosin heavy chain isoforms [233]. Even though the
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presence of the very small fast fibers is a typical feature of DM2, it is not specific as
they can also occur in congenital myopathies, although without nuclear clumps. Fast
fiber type atrophy may also be encountered in cortisone‐induced myopathy and
myasthenias. [71]. However, to our knowledge, selective atrophy of this degree in a
subpopulation of type 2A fibers has not been reported in any other muscle disease
apart from DM2. Its relevance is further implicated by the fact that it provides a clear
distinction to fiber atrophy in DM1.
Table 6. Summary of muscle histopathology in DM1 and DM2.
DM1 DM2
Fiber size variation +++ +++
Internal nuclei +++ +++
More in type 1 fibers in DM1, more in type 2 fibers in DM2
Type 1 fiber atrophya + to ++ -
Type 2 fiber atrophya + ++(+)
Type 2 fiber hypertrophy
- +
Nuclear clump fibers early
- ++
Nuclear clump fibers late
+ +++
Very atrophic fibers (diameter ≤ 6 μm)
(+) Type 1 and 2
+++ Type 2 only
In DM1 only at advanced stage
Necrotic fibers + (+)
Ring fibers ++(+) +
Sarcoplasmic masses ++(+) (+)
Targetoid fibers ++ (+)
Moth-eaten fibers (+)
Swirled sarcomeres (+)
Small group atrophy + - At advanced stage only
Fibrosis +++ ++ Late pathology only
Fatty replacement +++ ++ Late pathology only
Rimmed vacuoles (+) (+)
Ragged-red fibers (+) (+)
COX-negative fibers (+) (+)
Finding present +++, in >75% of biopsies; ++, in 25‐50% of biopsies; + in 10‐24 % of biopsies; (+) occasional finding, or restricted to few families; ‐ absent aas indicated by atrophy factor
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Figure 11. Serial sections of a VL muscle, DM2 patient, man, age 38. (A, B) Brown immunolabeling is DAB; blue is modified hematoxylin counterstain. (A) IHC with MHCfast Ab reveals characteristic high number of very atrophic fibers (black arrows), which are exclusively of type 2, as MHCslow Ab, in (B), does not recognize them. (C) ATPase staining at pH 10,4, which stains large type 2 fibers in dark brown, fails to detect the small fibers. (D) ATPase at pH 4,3 stains type 1 fibers dark brown. Note one necrotic fiber (white arrow), which expresses both fast and slow myosins as revealed by IHC.
In order to analyze the distribution of fast and slow fiber types in
myotonic dystrophies, we performed morphometric analyses, on 9 DM2 and 8 DM1
muscle biopsies, immunostained for fast and slow myosin isoforms. The “lesser
diameter” of each fiber was measured [71]. The results of approximately 500 – 5000
fibers in each sample were normalized to the mean diameters of each muscle type
analyzed (60 µm in VL, 50 µm in deltoid and biceps [228]). The results were
presented in metahistograms, and atrophy and hypertrophy factors in each group
were calculated [71]. The findings clearly indicated preferential fiber type 2 atrophy
in all DM2 specimens, in contrast to DM1, where marked fiber type 1 atrophy was
encountered with minor fiber type 2 atrophy occurring at advanced stages (Figure
12). It should be noted that a subpopulation of type 2 fibers was hypertrophic in
DM2 Biceps brachii muscle. In addition, atrophy of type 1 fibers occurs in DM2,
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however, to lesser extent: the percentage of atrophic type 1 fibers is lower than that
of type 2 fibers (I).
Interestingly, the presence of the very atrophic fibers in DM2 may
explain the previously reported fiber type 1 predominance, which according to our
study is not a true finding. The normal fiber type distribution in each muscle [122]
was restored when the very small fibers, visualized in IHC but missed in ATPase
staining, were included in the count. The reason why a subpopulation of type 2A
fibers undergoes severe atrophy in DM2 has not been clarified.
7.1.1. The atrophy of type II fibers in DM2 (I)
It was only after we used the immunohistochemical method for fiber type
differentiation in the muscle sections, that the preferential fiber type 2 atrophy was
fully recognized in DM2 [307], even though the possibility of it had been suggested
previously [21]. Schoser et al. later confirmed the finding with a larger set of samples
[268].
Figure 12. A representative metahistogram, showing marked type 2 fiber atrophy, and, on the other hand, slight type 2 fiber hypertrophy in DM2 patients. Note the bimodal fast fiber distribution. (710 fast and 635 slow fibers in Biceps were measured in three Italian DM2 patients. Figure kindly provided by Dr. Guillaume Bassez.)
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In this thesis work, the concept of very/highly atrophic fibers is used.
They are characteristic for DM2 where they appear very early in the disease course,
even in clinically unaffected muscles. Morphologically, they fall into at least three
categories: (1) nuclear clump fibers, (2) other extremely small fibers (≤ 6 µm) with
very little cytoplasm and none to few nuclei (in the plane of sectioning), and (3) very
thin, angulated fibers. Most of these fibers are round and scattered. They seem to
represent a unique pool, because they form a distinct population apart from regular
type 2A fibers (Figure 12). They may be encountered in DM1, but only in later stages
of pathology, less numerous compared to DM2, and in DM1 they most often co‐
express slow MyHC.
It is not clear if the nuclear clumps and other very atrophic fibers
represent distinct populations. Examination of longitudinal sections suggests that
they may represent a single population of fibers, which show differential
morphology depending on the number of nuclei trapped inside the slim fibers at the
plane of sectioning (Figure 13).
Figure 13. DM2 muscle. (A) immunofluorescent staining (MyHC‐II) showing two highly atrophic fibers. One of them is devoid of nuclei (arrow 1), whereas the another (arrow 2) is filled with nuclei, shown as black here due to lack of nuclear counterstain. (B) DAB staining (MyHC‐II) of a highly atrophic type 2 fiber (arrows) filled with nuclei (blue). The twisty fibers appear fragmented in the 6 µm longitudinal sections.
The nuclear clump fibers in DM2 are morphologically indistinguishable
from those seen in neurogenic atrophy, where they form as an end product of
denervation atrophy in the absence of re‐innervation. However, in neurogenic
conditions the nuclei may be pyknotic whereas this feature may be absent in nuclear
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clump fibers in DM2. However, no primary neurogenic involvement has been shown
in myotonic dystrophies, which is supported by the lack of fiber type grouping.
7.1.2. Protein expression in highly atrophic fibers (III, U)
We carried out further studies on protein expression in DM2 and DM1 muscles to
identify possible differences between them, which could lead to better
understanding of their differential muscle and fiber type involvement. In addition,
due to the presence of nuclear clump fibers in myotonic dystrophies, similar to
neurogenic atrophy, we also included a group of patients with various neurogenic
disorders. The results of immunohistochemical analysis of the very atrophic fibers in
DM2, DM1 and neurogenic atrophy are summarized in Table 7 and Figures 14‐16.
Altogether, we characterized the expression of 24 proteins, which are
involved in muscle contraction and Ca2+ handling, regeneration, denervation, and
apoptosis, by immunohistochemistry. A subset of the proteins show preferential
expression in type 2 fibers as described below.
Myosin heavy chain (MyHC)‐encoding genes (MYH1, ‐2, ‐3, ‐7 and ‐8)
are differentially regulated in fast and slow fibers, and during muscle development.
The expression of MyHC‐IIa isoforms in the highly atrophic fibers of DM2 was
confirmed with three different monoclonal antibodies raised against fast type
myosin heavy chains (Table 5), however, only the results obtained with MHCf are
shown. As a new finding, we showed that the population of very small fibers in DM1
co‐expressed myosins fast IIa and beta/slow, in contrast to DM2, where they
expressed only type IIa. In neurogenic atrophies, virtually all small atrophic fibers
expressed MyHC‐IIa, showing moderate to high co‐expression of MyHC‐beta/slow.
The MyHC‐pn is widely used as a marker of regeneration, but is not exclusive to
regenerating fibers [71]. There was high expression of the MyHC‐pn isoform in very
atrophic fibers in all groups. Since the majority of very atrophic fibers in progressive
neurogenic atrophy without any regeneration activity express MyHC‐pn, this may be
turned on and expressed also by other mechanisms than regeneration. MyHC‐emb
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positive fibers were absent in most biopsies, showing expression in a few DM
biopsies only, with advanced muscle pathology. This observation is congruent with
its reported expression mostly in small basophilic regenerating fibers having survived
severe injury, in muscular dystrophies and other necrotic myopathies [71]. A similar
finding was observed with the transcription factor myogenin (MYOG), and the
intermediate filament protein vimentin (VIM), which appear early during myogenesis
and serve as regeneration markers in muscular dystrophies [71, 209].
Neural cell adhesion molecule (NCAM1) has distinct roles in
myogenesis, synaptogenesis, and synaptic maintenance [60]. It is transiently re‐
expressed after denervation and during re‐innervation, decreasing progressively as
regeneration proceeds [324]. Hence, it has been a candidate marker for recently
occurred denervation atrophy [71]. Interestingly, we found that one third of the
atrophic fibers in DM2 abundantly expressed NCAM1, outnumbering the small fibers
in DM1, and even more intriguingly, the severely atrophic fibers in neurogenic
atrophies. In fact, the pattern of NCAM1 expression was different in the latter group,
showing positive signal more frequently in intermediate ‐ large fibers, which was not
encountered in any specimens in myotonic dystrophies. The usage of NCAM1
expression as a marker of neurogenic atrophies thus seems to be very limited.
Because we were preoccupied with the question of why the highly
atrophic type 2 fibers persist in the DM2 muscle, we investigated the expression of
the oncoprotein BCL2 (BCL2) in the muscle biopsies. Apoptosis is required during
normal tissue remodeling, development and regeneration. BCL2 inhibits cells from
entering apoptosis, but it also has a distinct effect on cell cycle [59]. There are rather
controversial reports on the role of apoptosis in muscular dystrophies [285];
however, it seems not to be generally involved as a pathomechanism, at least not in
myotonic dystrophies [186]. In order to verify this we used an Ab raised against
activated Caspase‐3 (CASP3), a cysteine protease playing a key role in executing
apoptosis in many cell types, and a commonly used marker for apoptosis [258]. To
our surprise, the majority of the very atrophic fibers in DM2 expressed BCL2.
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Table 7. Protein expression in highly atrophic fibers a by IHC.
Protein Gene DM2 DM1 NA
MyHC-IIa MYH2 +++ ++ ++
MyHC-beta MYH7 (+) ++ ++
MyHC-pn MYH8 +++ +++ ++
MyHC-emb MYH3 (+) (+) -
SERCA1 ATP2A1 +++ +++ +++
fTnT TNNT3 +++ ++ +++
NCAM NCAM1 ++ + +
nNOS NOS1 + n/a n/a
BCL2 BCL2 +++ + ++
Caspase-3 CASP3 (+) n/a n/a
Dystrophin DYS ++ n/a n/a
Myogenin MYOG (+) (+) (+)
Vimentin VIM (+) + (+) aThe population of very atrophic fibers (≤ 6 µm in diameter) and nuclear clump fibers, not expected to be functional. NA, neurogenic atrophy; n/a, not assessed. Protein expression: +++, in >75% of highly atrophic fibers; ++, in 30‐74%; +, in 1‐15%; (+), in <1%; ‐, absent. The results indicate the proportion of the highly atrophic fibers expressing the given antigen in DM2 (n = 20), DM1 (n = 5), and NA (n = 9).
In contrast, in DM1 BCL2 immunoreactivity was present in only one biopsy, in which
approximately 5% of the very small fibers were positive. In neurogenic atrophies,
BCL2 positive fibers were moderately frequent, albeit with a lot of dispersion among
the samples (range 1‐100% positive cells). Caspase‐3 was not detected in DM2
biopsies, supporting the prevailing concept of apoptosis not playing a major role.
However, the combined high BCL2 expression and absent Caspase‐3 could offer a
partial explanation to the persistence of the very atrophic, non‐functional type 2A
fibers in DM2: the anti‐apoptotic BCL2 might prevent the cells from entering
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Figure 14. Immunohistochemical panel showing key findings in DM2. Upper panel (I): (A) H&E; (B) MyHC‐IIa/x; (C) MyHC‐beta/slow; (D) MHCd; (E) NCAM1; (F) SERCA1; (G) fTnT; (H) DYS; (I) nNOS. Lower panel (II): (A1,2) MyHC‐IIa/x; (B1,2) MHCn; (C1,2) BCL2, showing the mutually exclusive staining of MyHC‐pn and BCL2. (A‐G) and (A1‐C2) DAB immunohistochemistry; (H, I) immunofluorescence with (H) Alexa 488 and (I) Alexa 564 labeled secondary antibodies.
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Figure 15. Panel showing key immunohistochemical findings in DM1 (DAB). (A) MyHC‐IIa/x; (B) MyHC‐beta/slow; (C) MHCpn; (D) MHCd; (E) NCAM1; (F) BCL2.
Figure 16. Panel showing key immunohistochemical findings in neurogenic atrophy (DAB). (A) MyHC‐IIa/x; (B) MyHC‐beta/slow; (C) MHCpn; (D) MHCd; (E) NCAM1; (F) BCL2.
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apoptosis. Another interesting finding in DM2 was revealed when serial sections,
stained for MHC‐pn and BCL2, were examined. Mutually exclusive expression of
these markers was often observed (Figure 14, panel II). We do not have a specific
explanation to this finding at the moment.
We also observed reduced sarcolemmal labeling of dystrophin and
neuronal nitric oxide synthase (nNOS) in the very atrophic fibers in DM2 (Figures 14H
and 14I). Reduction of nNOS was prominent, and present also in a subset of
large/regular fibers. These findings, although not reported with DM2 before, may be
non‐specific given that several sarcolemmal proteins are reported to show low
immunoreactivity in MyHC‐pn positive, small atrophic fibers [71]. However, nNOS
has been suggested to affect excitation‐contraction coupling, its activity is regulated
by Ca2+/calmodulin, and its dysregulation has been implicated in some muscular
dystrophies, which makes it an interesting targer for further investigation in DMs.
We also studied two proteins preferentially expressed in, but not
restricted to fast type 2 fibers: the fast skeletal muscle troponin T (fTnT/TNNT3), a
sarcomeric thin filament component, and the Ca2+ ATPase SERCA1 (ATP2A1), both of
which are involved in regulation of muscle contraction [40, 328]. SERCA1 was
expressed in a majority of the atrophic fibers in all groups, whereas fTnT showed
moderate to high expression in atrophic fibers in DM2 and DM1, and high expression
in neurogenic atrophies. Unfortunately, even though not strictly following the fast‐
slow fiber distribution, neither of these proteins were differentially expressed in very
atophic and regular type 2 fibers, thus giving no further insight into the question of
why a subpopulation of type 2A fibers are subject to severe atrophy in DM2.
It should be noted that the neurogenic atrophies was the most
heterogeneous group, relative to MyHC‐pn, NCAM‐1 and BCL2 expression in the very
small, atrophic fibers, showing more dispersion in results and greater standard
deviation values. This is understandable because of their divergent etiologies.
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7.1.3. Using fiber type II atrophy as a predictive factor for DM2 diagnosis
The highly atrophic type 2 fibers appear very early in DM2 muscle pathology,
whereas in DM1, they occur only later with severe general muscle pathology. In
addition, as an important distinctive feature, the very atrophic fibers in DM1 express
both slow and fast MyHC isoforms. Hence, the early appearance of highly atrophic
type 2 fibers, with or without nuclear clumps, can very well be used as a diagnostic
tool, for screening of patients for molecular diagnostic testing. This finding has been
successfully used for this purpose in Finland for almost a decade, resulting in several
DM2 diagnoses in patients, who otherwise lacked key features of the disease, such
as myotonia (Udd, personal communication). The number of internalized nuclei is
greater in type 2 fibers in DM2, and in type 1 fibers in DM1, further reinforcing the
observation that the distinct fiber populations are differentially affected in DM1 and
DM2 [20, 227]. In France, the combination of atrophy and central nuclei in type 2
fibers was found to predict DM2 even more efficiently [20].
7.1.4. Satellite cells (U)
Defective differentiation in primary myoblast cultures derived from satellite cells
have been reported with DM1 [88]. Similar findings have not been observed in DM2
[219]. However, repeatedly in our DM2 cell cultures, myoblast proliferation after 4‐5
passages has been limited even if differentiation seemed to be normal (Vihola et al.
unpublished results).
Pax7 is considered an appropriate protein marker and expressed in
quiescent skeletal muscle satellite cells [241], the number of which has been
reported to decline with age [252]. Consequently, as a separate experiment, the
number of skeletal muscle satellite cells in DM2 (n = 4), DM1 (n = 5), neurogenic
atrophy (n = 4), and healthy control (n = 1) biopsies was assessed. Approximately 200
fibers were included per biopsy, counting all fibers in randomly selected fields. The
number of Pax7 positive cells in satellite cell position was proportioned to the total
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number of fibers; the results are indicated as the number of Pax7+ cells (in satellite
cell position) per 100 muscle fibers counted (Table 8 and Figure 17).
Table 8. Relative abundance of Pax7+ cells.
C 7
DM1 20
DM2 25
NA 20
NA, neurogenic atrophy; C, healthy control
Figure 17. Pax7 staining (DAB IHC) in DM2 muscle showing several satellite cells (black arrows) and nuclear clumps (N, white arrows).
Other Pax7+ cells were present within the interstitial space of the
biopsies examined, which are likely to represent a distinct pool of progenitor cells.
These were not included in the number of satellite cells. The results indicate that the
number of spindle‐shaped, Pax7+ cells in satellite cell positions was increased in all
disease groups, compared to control biopsy. We observed the highest number of
Pax7+ cells in DM2. The reported proportion of satellite cells in human muscle
normally varies between 3‐6% of all muscle fiber nuclei [252], which is in accordance
with our control muscle. The number of satellite cells is affected by age and muscle
type [31]. The increase of satellite cells in DM2, DM1 and neurogenic atrophy is
probably caused by basic attempts to regenerate in response to muscle injury. These
results agree with previous reports of DM2 and DM1 that indicated unaffected
satellite cell proliferation [88, 219].
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7.1.5. Expression of Ca2+ handling proteins in DM (IV)
Because several genes encoding proteins involved in Ca2+ handling showed
dysregulation at the mRNA expression level for both DM1 and DM2 (see chapter
7.2.5.2.), we analyzed a subset of them for protein expression, using
immunohistochemistry and Western blotting. The analyzed proteins were selected
due to their central roles in Ca2+ release (RYR1, DHPR, TRDN, JPH1) and re‐uptake
(SERCA1, SERCA2); Ca2+ storage (CASQ1, CASQ2); and Ca2+‐dependent signaling
(PPP3CA, NFATC3). The results are summarized in Table 9.
RYR1 and CASQ2 were clearly reduced by IHC for DM2, but WB failed to
reproduce the finding, showing no statistically significant difference between the
groups (Normal, DM1 and DM2). However, CASQ2 did show a trend towards being
decreased in DM1 and DM2, but high variability between samples in all groups gave
p‐values < 0.05. DM1 samples showed a similar finding for RYR1 and CASQ2, with
additional reduction in DHPR and JPH1 staining. DHPR showed no change in WB,
whereas JPH1 was significantly reduced in DMs as a group. Notably, total protein
levels of RYR1 and SERCA1, both of which show aberrant splicing in DM, showed no
dysregulation in WB. We do not have a definitive explanation for the discrepancy
between IHC and WB for RYR1 and CASQ2. However, both show fiber‐type specific
expression (RYR1 shows higher expression in fast fibers and CASQ2 in slow fibers),
which makes quantitative WB difficult to interpret. In addition, both accumulate in
the non‐functional highly atrophic fibers in DMs, which may contribute to the result.
Seven of the ten proteins analyzed showed up‐regulation of total
mRNA level (Table 9). However, only four of them were dysregulated at total protein
level, including SERCA2, TRDN, JPH1 and calcineurin. Furthermore, none of the
protein expression profiles were completely in accordance with mRNA expression,
with ATP2A2 and JPH1 showing reduced total protein amounts in DM, whereas
TRDN and calcineurin were comparatively increased in DM2 versus DM1. The
observed contradiction between mRNA and protein expression is probably caused by
translational block or dysregulation of mRNA processing in DMs [114, 254, 256], or
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other, yet unidentified regulatory events. It is worth noting that mRNA levels alone
are often not sufficient to predict the steady‐state protein levels, and therefore the
changes in mRNA expression should be validated at protein level to be able to
evaluate the true physiological relevance of the findings.
The reduced expression of the Ca2+ release complex constituents, RYR1
and CASQ2 in both DMs, and also JPH1 in DM1 by IHC may constitute the most
relevant findings in this part of the investigation, as it may affect muscle contraction.
This finding also emphasizes the need for using several methods in parallel, in order
to obtain information of expression at multiple levels, including mRNA and protein
expression, and the subcellular localization of proteins.
Table 9. Expression of Ca2+ handling proteins in DM1 and DM2.
Protein IHC WB EP
DM1 DM2
RYR1 ↓ ↓ ‐ ‐
DHPR ↓ ‐ ‐ ‐
SERCA1 ‐ ‐ ‐ ↑
SERCA2 na ‐ N > DM2 ↑
CASQ1 na ‐ ‐ ↑
CASQ2 ↓ ↓ ‐ ↑
TRDN na ‐ DM2 > DM1 ↑
JPH1 ↓ ‐ N > DM ↑
Calcineurin na ‐ DM2 > DM1 ↑
NFATC3 na na DM2 > DM1 ‐
IHC, immunohistochemistry; WB, western blotting; EP, mRNA in expression profiling; ‐, no change; ↑, increased; ↓, decreased; na, not assessed
88 |
7.2. Molecular genetic tools for DM2 mutation identification and size
determination (II)
7.2.1. PCR‐based allele‐sizing (II)
All the PCR‐based allele‐sizing methods, using primers in mutation‐flanking exons,
were consistent in showing only a single allele in all DM2 patients at the ZNF9 locus.
However, the PCR methods using fluorescent detection, automated laser fluorescent
(ALF) DNA analysis (Cy5‐label) and capillary electrophoresis (FAM‐label), were able to
detect alleles differing by only 2 bp, which were usually missed by conventional
PAGE using 32P‐dCTP‐labeling.
7.2.2. RP‐PCR and FIGE (II)
RP‐PCR and FIGE methods both identified all 12 known DM2 patients in group 1, and
14 new DM2 mutations in cohort 2 with previously undefined muscular disorders (II).
7.2.3. In situ –hybridization (II)
Due to the huge size, repeat nature, and somatic instability of the DM2 mutation,
conventional molecular genetic methods are insufficient for its identification [18,
66]. When the DM2 mutation was identified in 2001, and for some years following,
there was no reliable genetic testing widely available. The development of an
accurate genetic test for DM2 mutation verification based on muscle biopsy material
was thus pursued here; bearing in mind that conventional methodology and imaging,
available in any medical laboratory or pathology department could be employed.
Furthermore, it was considered that this approach should be able to determine the
DM2 mutation expansion size, which had continued to prove extremely difficult to
assess.
To meet these requirements, two novel in situ‐hybridization (ISH)
methods were developed for the direct visualization of the DM2 mutation in
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genomic DNA: chromogenic ISH (CISH) on muscle biopsies, and fiber‐FISH on
extended DNA fibers derived from peripheral blood lymphocytes. The former was
found suitable for routine molecular diagnostics, and the latter was especially
efficient for estimating the mutation size.
The DM2 mutation detection in the two methods was based on
digoxigenin end‐labeled oligonucleotide sense probes [(CCTG)8]. Antisense (CAG)10
probes had previously been used for showing the accumulated mutant mRNA in the
ribonuclear inclusions in DM1 [283] and (CAGG)8 in DM2 [158]; however, using signal
amplification methods, polymer amplification technique in CISH, and two layers of
fluorescent labels in fiber‐FISH, we were able to increase the sensitivity and directly
visualize the mutation.
In order to validate the new ISH methods; to investigate the
concordance between the high‐accuracy methods RP‐PCR and FIGE; and between
three different PCR‐based allele sizing methods for primary screening, a blinded
study involving four laboratories was performed. Two sets of samples were used:
previously confirmed DM2‐patients (n = 12), and a cohort of patients with undefined
neuromuscular disorder, with biased selection towards myotonia symptoms (n = 31).
7.2.3.1. CISH (II)
When the sense (CCTG)8 probe was used for hybridization, the chromogenic in situ‐
hybridization method successfully identified the DM2 mutations directly as a single‐
spot signal per nucleus in all known DM2 patients´ frozen muscle biopsies (Figure
18D). Similarly, the same 14 novel DM2 mutations in group 2, as determined with
RP‐PCR and FIGE, were readily identified by CISH. In contrast, CISH failed to detect
the genomic mutation directly in paraffin‐embedded specimens (n = 9), most likely
due to loss of sensitivity during the long fixation and embedding procedure.
However, the ribonuclear inclusions were identified with the antisense probe in all
DM2 specimens, including the paraffin sections. Frozen muscle samples from healthy
90 |
individuals and DM1 patients remained negative with both sense and antisense
probes, confirming the specificity of the protocol.
This method provides an improvement in specificity compared to a
previously described FISH‐method, where the (CAG)10 antisense probe, directed
against the DM1 mutant transcript, detected the ribonuclear foci in both DM1 and
DM2 [174]. In theory, by lowering the stringency conditions in the hybridization,
similar (CCTG/CTG)n or related expansions could be screened. However, such trials
have been unsuccessful so far.
Molecular genetic testing, based on genomic DNA derived from
peripheral blood, is currently the standard diagnostic method. However, the CISH
method we developed here was used in Finland as a primary diagnostic method for 5
years (2003 – 2007), during which time 105 patients were confirmed with DM2
mutation. At the same time, an RP‐PCR test was being optimized, and used side by
side with CISH. As quality control of DM2 mutation verification, all single‐allele
samples determined to be negative with RP‐PCR/CISH were sent to Prof. Krahe´s
laboratory in Houston, Texas for FIGE analysis. In 8 years, only 1 false negative result
was obtained with CISH, in a patient carrying a very short expansion of (CCTG)55 (Udd
et al. unpublished results). Also RP‐PCR has produced 1 false negative result among
180 DM2 patients in 8 years, signifying 99 % sensitivity for both methods. The exact
limit for CISH sensitivity is not known. The very short expansions, like (CCTG)55, do
not necessarily form ribonuclear foci, resulting in negative outcome with antisense
probe as well. It has been reported with DM1 that ≤ 100 CTG repeats do not form
detectebale foci with FISH [35].
CISH method is still valid as a research and diagnostic tool. For patients
with inconclusive first results with RP‐PCR and clinical features of DM2, CISH is still
applied as part of the diagnostic setting. In addition, since CISH is capable of
detecting accumulated mutant RNA in paraffin‐embedded tissues, this method can
and has been used for retrospective studies on several kinds of tissue, as old as 17
years [19]. CISH has certain advantages over FISH in some situations. Performing
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FISH on paraffin‐embedded specimens is challenging because of strong background
fluorescence due to the procedure. Old surgical specimens taken for other reasons
are usually formaldehyde‐fixed, paraffin‐embedded and stored at room temperature
for decades. Even in such specimens, the RNA in ribonuclear inclusions has proven
robust.
Figure 18. showing the in situ ‐hybridization findings in DM2. (A) Fiber‐FISH: the DM2 expansion mutation is revealed by binding of the sense (CCTG)8 probe (fluorescein, green), while the flanking sequences are visualized by 814L21 (TRITC, red). (B) Muscle biopsy with macrophage invasion in a necrotic fiber. Accumulated RNA is visualized (dark brown) using antisense (CAGG)8 probe. (C) Fibroblast nuclei showing ribonuclear foci with (CAGG)8 probe. (D) Myonuclei in section detected with the sense probe, revealing one spot per nucleus, corresponding to the mutation. (E) Nuclear clump fiber detected with antisense probe, showing accumulated mutant RNA.
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7.2.3.2. Fiber‐FISH (II)
In fiber‐FISH, co‐hybridization of the oligonucteotide probe (CCTG)8/(CAGG)8, and
the BAC‐clone 814L21 covering the DM2 locus, hence flanking the repeat mutation,
resulted in visualization of the mutation directly in its locus in all 5 DM2 patients
analyzed from groups 1 and 2 (Figure 18A). The hybridization signals for both sense
and antisense oligonucleotide probes were always localized to a gap at the telomeric
end of the BAC814L21 signal. The mutation size measurements were performed by
using the centromeric 115 kb portion of the 814L21 for calibration. The estimated
mutation lengths in the Finnish patients, obtained with this method varied between
2.4 kb and 23.6 kb within the whole group, while the mean size of all mutations
measured was approximately 10 kb, which was slightly smaller compared to
estimations done with FIGE. The extreme variation observed between the individual
DNA‐molecules measured may be largely explained by an intrinsic feature of the
DM2 mutation: the profound somatic instability and mosaicism.
The Finnish patients all had an expansion mutation approximately 17‐
19 kb in size, as estimated by FIGE that apparently was more sensitive for larger
expansions. In addition, an Italian DM2 patient with a relatively short expansion
mutation (approximately 4 kb) was used to test the sensitivity of the method, with a
positive result, hence establishing the sensitivity boundary to (CCTG)1000 so far.
Fluorescent in situ hybridization (FISH) offers an alternative means for
the repeat length measurement, when performed on single extended DNA
molecules (fiber‐FISH). However, it requires highly specialized methodology and
equipment for analysis, thus PCR‐based Southern modifications such as FIGE and
others have remained the major methods applied for size estimation [18, 34, 51, 66,
119, 199].
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7.3. Splice variant analysis (III, IV)
Splice variant analysis (SVA) for detecting aberrant splicing events of TNNT3 (fTnT),
LDB3 (ZASP), RYR1 (RYR1) and ATP2A1 (SERCA1) was performed using RT‐PCR and
cDNA prepared from total muscle RNA. The primers were designed to have binding
sites either in exons flanking the differentially spliced exon, or in the differentially
spliced exon itself.
TNNT3 encodes the sarcomeric thin filament‐associated troponin
complex subunit T, involved in the execution of muscle contraction. The TNNT3
gene, expressed predominantly in fast type 2 fibers, shows extensive
developmentally regulated differential splicing [328]. The fetal (F) exon, located
between exons 8a and 9a, is included in the two fetal mRNA isoforms (A, 225 bp and
D, 188 bp). Three adult isoforms are expressed in skeletal muscle [328]. In
agreement with this, we detected 3 isoforms both at mRNA and protein
(approximately 30 kD) levels, expressed in relatively equal amounts in all samples. In
DM1 and DM2 samples, two additional isoforms, corresponding to A and D, were
identified at the mRNA level, the isoform A being dominant (Figure 19). When the
proteins were analyzed using WB the presence of a fourth, slightly larger isoform
corresponding to the fetal isoform A, was recognized in 60% of the DM2 samples.
This fetal isoform was not present at the protein level in healthy control or DM1
samples, suggesting that it may have a greater role in DM2 pathogenesis compared
to DM1.
The roles of the Z‐disc alternatively spliced PDZ‐domain containing
protein (ZASP) are not fully elucidated. It interacts with α‐actinin, and is localized to
the sarcomeric Z‐disc of both skeletal and cardiac muscles [14]. Mutations in LDB3
cause diverse muscular disorders, including dilated cardiomyopathy [304] and
myofibrillar myopathy [99, 270]. Two LDB3 exons (4a and 7) show tissue‐dependent
expression patterns with predominant expression in the heart muscle [113], and the
mutations causing skeletal myopathies are found in exon 6.
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When analyzing ZASP expression by WB, we observed two bands of
MWs approximately 32 and 78 kD in healthy controls, corresponding to the short
and long skeletal muscle isoforms previously reported [113]. In all DM2 patients and
most DM1 patients analyzed, an additional band of approximately 95 kD was
detected, which was more pronounced in DM2 patient samples. Altogether four RP‐
PCR assays were designed to identify the larger ZASP isoform at the mRNA level. The
results revealed the presence of a long cardiac isoform, including both exons 4a and
7, in DM1 and DM2 skeletal muscle biopsies. Quantitative analysis showed that the
relative proportion of both cardiac exons was greater in DM2 compared to DM1. The
retention of exon 7 has previously been reported for both DM1 and DM2 at the
mRNA level [156, 165]. A novel DM specific isoform encompassing both exon 7 and
4a was demonstrated. Furthermore, the presence of DM specific abnormal protein
products appeared to be greater in DM2, although also evident for DM1 samples.
The Ca2+ release channel RYR1 is also expressed preferentially in fast
muscle fibers, and alternative splicing of its two exons, 70 and 83, is under
developmental regulation [91, 225]. SVA showed the exclusion of the RYR1 exon 70
in DM2 samples, giving rise to the ASI(‐) isoform, lacking part the inhibitory domain,
as previously reported only for DM1 [133, 134]. In this study, the difference between
Figure 19. Distribution of TNNT3 isoforms. Proportions of the different TNNT3 mRNA isoforms detected in DM1, DM2 and controls. The fetal isoform A (225 bp; green) is increased in DM1, and even more pronounced in DM2. (DMx, non‐DM1/DM2 muscular dystrophy; N, normal/healthy control)
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DM1 and healthy controls was not statistically significant, probably due to high
variability among samples, and small number of DM1 samples (n = 3) analyzed.
Unfortunately, the RYR1 protein was too large (565 kD) for distinguishing the
isoforms by WB. Therefore, we failed to show the presence of the aberrant RYR1
isoform at protein level.
SERCA1 ATPase Ca2+ reuptake pump shows, similarly to RYR1,
preferential expression in fast muscle fibers. ATP2A1 contains an alternatively
spliced exon 22, which is present in the adult isoform, but excluded from the fetal
isoform [39]. With RP‐PCR, we confirmed the presence of the fetal isoform
(ATP2A1B) in skeletal muscle of DM1 patients [134], and, as a novel finding, in DM2
patients as well. ATP2A1 also showed a greater relative proportion of the fetal
isoform in DM2 patients compared to DM1. As SERCA1 is also large (110 kD), the
exclusion of a single exon could not be demonstrated by WB, due to the limited
resolution of the method.
In summary, SVA showed that the expression of fetal isoforms of four
genes (TNNT3, LDB3, RYR1 and ATP2A1) were increased in both DM1 and DM2, and
that the relative proportions of the immature forms were even higher in DM2
patient muscle (even though a statistically significant difference could not be
confirmed for RYR1, possibly due to small samples number). The splicing of all these
four genes is regulated by MBNL1, and given that MBNL1 has been shown to
accumulate more heavily in the ribonuclear inclusions of DM2 compared to DM1
[156], we suggest that MBNL1 loss‐of‐function may play a greater role in DM2 than
in DM1. In case of the sarcomeric proteins fTnT (TNNT3) and ZASP (LDB3), the
corresponding fetal protein levels were also increased, showing prominent steady‐
state protein levels in muscle. This strengthens our hypothesis that these changes
may have biological and pathological relevance. Because of (dys)regulation at the
level of translation and protein degradation, the mRNA expression of a given gene or
isoform alone does not necessarily indicate the presence of the corresponding
protein product. The developmental protein isoforms of RYR1 and SERCA1 could be
indicated, for example, by raising isoform‐specific antibodies.
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The findings concerning fTNT, RYR1 and SERCA1 were particularly
interesting, as their expression is concentrated in the fast fibers. In addition, all of
them show relatively high expression in the population of very atrophic fibers in
DM2 and DM1. However, the impact of the highly atrophic fibers on mRNA and
protein expression results obtained with expression profiling, splice variant analysis,
and Western blotting is not known. The methods used in this study were not able to
show whether the very atrophic fibers expressed relatively different proportions of
the fetal isoforms compared to the regular, functional fibers. Thus, the question of
why a particular subpopulation of type 2 fibers is susceptible to atrophy remains
unsolved. A method capable of analyzing mRNA and protein expression in the
atrophic fibers separately would be warranted, such as laser micro‐dissection.
Reports on aberrant splicing in DM1 and DM2 patients demonstrate
extreme variation between samples. In mouse models such variation is often less
prominent, due to more homogenous genetic background [134]. In a conclusion, the
significance of aberrant splicing is expected to be highly variable between individual
patients.
Although extensive aberrant splicing has been identified in DM, recent
research indicated that it is not a completely DM‐specific pathomechanism. In a
study of Mbnl1‐deficient and CUGBP1‐overexpressing mouse models, up to
approximately 50% of aberrant splicing events were found to be caused by neither
Mbln1 nor CUGBP1 [124]. Similarly, Bachinski et al. (2010) showed that the abnormal
splicing of MEF2A and MEF2C was not restricted to DM, but was also observed in
other muscular dystrophies [16].
7.4. Expression profiling (III, IV)
In order to investigate mRNA expression in myotonic dystrophies, a global
expression profliling was performed using Affymetrix U133 Plus2.0 chips and muscle
biopsies from healthy control individuals and DM1 and DM2 patients. Instead of
profiling the entire set of genes on the array, a list of 327 probe sets comprising a
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muscle‐specific gene array was generated. Subsequently, a second list of 2,490
probe sets was created, consisting of genes involved in Ca2+ regulation, and their
expression was analyzed in DM. The entire gene expression data can be found at
www.ncbi.nlm.nih.gov/geo/ (GEO series numbers GSE7014).
7.4.1. Muscle‐specific gene array (III)
Comparing DM1 and DM2 patients to controls, 11% of the probe sets covering 28
unique genes showed differential regulation. In the supervised two‐way hierarchical
cluster analysis, where DM1 and DM1 patients clustered together, the majority of
the dysregulated genes (86%) showed increased mRNA expression in DMs compared
to controls. Three genes showed differential regulation between DM1 and DM2:
CAMKK2 (downregulated in DM2), CALML6 and MEF2C (both upregulated in DM2).
Cluster analysis revealed four major gene groups: one group with
down‐regulated genes, and three groups with up‐regulation. Group 1 consisted of
four down‐regulated genes. In groups 2 and 3, the MYH genes (ten altogether)
showed over‐expression at the mRNA level. The group 4 included several
transcription factor encoding genes, as well as genes encoding for proteins involved
in Ca2+ handling.
Interestingly, the MEF2 family of transcriptional effectors of diverse
Ca2+ signaling pathways [179] showed major dysregulation. One isoform of MEF2B
was down‐regulated, whereas MEF2A and MEF2C, as well as the homebox protein
SIX1 were up‐regulated in both DM1 and DM2. A few differences were seen between
DM1 and DM2, including higher up‐regulation of NFATC4 and one MEF2C isoform in
DM2. NFATC4 has been shown to drive MyHC‐2B expression (the rat homolog of
human MYH4) [47]. SIX 1 is implicated in establishing and maintaining the fast fiber
phenotype [98], similarly to MEF2A [2, 3]. MEF2C, on the other hand, plays an
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Figure 20. LDB3 exon‐specific microarray. The heatmap shows inclusion of the fetal exons 4a (red arrow on left) and 7 (red arrow on right) in DM1 and DM2 (4a also in non‐DM disease controls, DMx), but not in healthy controls (Normal). (Figure kindly provided from Prof. Ralf Krahe, Dr. Linda L. Bachinski and Dr. Keith Baggerly, all from the University of Texas M.D. Anderson Cancer Center, TX, USA.)
important role in adult muscle, as it drives the transcription of several structural
genes, including those encoding sarcomeric proteins [229]. As a conclusion, the
extensive dysregulation of the myogenic transcription factors may be of major
importance in DM pathology, as they regulate several downstream effector genes.
LDB3 was not among the genes showing dysregulation at the total
mRNA level in the first analyzed set of sarcomeric and other muscle‐specific protein
encoding genes. However, in a separate expression profiling experiment, the exon‐
specific expression array, a novel aberrant splice event was identified for LDB3 in
DMs, the inclusion of fetal exon 4a (Figure 20). This aberrant splice variant was
further validated using SVA and WB (Results 7.3.).
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7.4.2. Ca2+ gene array (IV)
Several genes encoding Ca2+ handling proteins were among the dysregulated genes
in the first EP analysis of muscle‐specific genes. When Ingenuity Pathway Analysis
was performed on all genes showing dysregulation in the the entire EP data set, Ca2+
signaling was identified as the single most significant canonical pathway (Sirito et al.,
unpublished results). To investigate the expression of these genes in DMs, we
generated a second list of 2,490 probe sets (1,039 unique genes) involved in Ca2+
metabolism. Of them, 200 (8%) probe sets were dysregulated in DM1 and DM2
compared to controls. Of the dysregulated probe sets 82% showed increased mRNA
expression in DMs. When the 200 dysregulated probe sets were compared between
DM1 and DM2, 21 of them, representing 16 unique genes, showed differences in
mRNA expression.
The expression of a subset of the dysregulated genes playing pivotal
roles in muscle Ca2+ metabolism was studied at the protein level (Table 9). Many of
the genes involved in regulating the Ca2+ release from SR showed up‐regulation in
both DMs, including TRDN (triadin), ASPH (junctin), JPH1 (junctophilin), CASQ1 and
CASQ2 (Table 9). The Ca2+ re‐uptake channel encoding genes ATP2A1 (SERCA1) and
ATP2A2 (SERCA2) showed up‐regulation in both DM1 and DM2. Of these, ATP2A2,
TRDN abd JPH1 showed differential expression between DM1 and DM2, with higher
expression in DM1.
Similar to aberrant splicing, dysregulation at the level of transcription
may also be a more common phenomenon in muscular dystrophies than previously
estimated. An extensive study comparing total mRNA expression by EP in HSALR and
Clc1‐deficient mice suggested that a significant proportion of the transcriptional
dysregulation could be induced by unspecific secondary changes, such as myotonia
[212]. In the same study, dysregulation of several genes involved in Ca2+ signaling
was reported, similar to our results. Given the central role of Ca2+ in muscle signaling
and contraction, its strict regulation is highly important, and the aberrant gene
expression reported herein may well be significant for DM pathophysiology.
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However, the expression should be also validated at protein level. A high‐throughput
method, such as protein expression array, would be more suitable for large‐scale
quantification. When analyzing single proteins in detail, however, the IHC and WB
methods represent a good basic set of tools.
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8. CONCLUDING REMARKS AND FUTURE PROSPECTS
Our recent epidemiological work suggests that DM2 is far more common than DM1,
at least in Finland. Despite adequate attention to the search for and molecular
testing of DM2 in many centers in Finland, it is obvious that the large majority of
patients are still not identified. We are currently aware of approximately 300
patients in Finland, whereas the number of symptomatic patients based on the
mutation frequency in the population should be more than 1000. This discrepancy
may be explained in part by the fact that the mean clinical phenotype is much less
severe than the phenotype for DM2 reported thus far. Many patients may have a
very late onset of symptoms, after age 60 or 70, when symptoms of myalgia and/or
mild proximal weakness are taken for normal aging and do not necessarily lead to
neuromuscular examinations. Furthermore, whether the DM2 mutation in all
circumstances is 100% penetrant is not fully settled.
In light of these facts, the diagnosis of DM2 is an enormous challenge
for the clinician. However, there is a clear rationale as to why DM2 diagnostics
should be enhanced. DM2 patients may present with severe cardiac conduction
defects, with risk of sudden cardiac death, thus a cardiac surveillance similar to DM1
patients is recommended. DM2 patients are frequently misdiagnosed as having
polymyositis, unexplained hepatopathy, chest pain, fibromyalgia, and other
conditions, which may lead to extended immunosuppressive treatments or
unnecessary liver biopsy. Myalgic pains may be disabling, sometimes leading to
incapacity for work, which may cause socio‐economical problems. Many of the
known DM2 patients in Finland had long histories of seeking medical help and
advice, before reaching final diagnosis by DM2 genetic testing. The definite diagnosis
in these cases is a therapeutic relief, explaining the enigmatic symptoms and
correcting the management of the patient.
Major issues persist concerning the need for a better understanding of
the disease pathogenesis in DMs. The mutations are of course the sole cause of the
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diseases, but which of the downstream effects of the mutations contribute to the
pathology, is less clear. For a while the toxic mutant RNA, accumulating as
ribonuclear foci sequestering proteins needed for proper alternative splicing,
seemed to have clarified the molecular basis of disease. However, nuclear foci are
not mandatory for the disease, and there are also other shortcomings in trying to
explain all the pathology through this mechanism. MBNL1 is more heavily
sequestered in DM2 nuclear foci, rendering the soluble MBNL1 fraction smaller in
DM2. This would be expected to lead to a more severe muscle phenotype in DM2,
including myotonia based on greater CLC‐1 splicing abnormality in DM2 compared to
DM1, but this is not the case. On the contrary, myotonia is much less severe and can
be absent even in EMGs of DM2 patients, indicating other mechanisms in addition to
MBNL1 sequestration are of importance. The appearance of these large ribonuclear
foci in all cell types very early at developmental stages is also in some disagreement
with the late onset of disease in DM2.
There is no conclusive evidence of CUGBP1 involvement in DM2
pathology, although it apparently plays a role in DM1. The cause of congenital and
early onset forms in DM1 and the lack of these in DM2 have no definite molecular
explanation as yet.
The muscle wasting, considered to be the major cause of disability in
DM1 and DM2 is not very well understood either. Even though we have shown
abnormal splicing of sarcomeric protein coding genes, it is far from clear that these
abnormalities form the major cause of progressive muscle loss. The distinctive
histopathological difference between DM1 and DM2, the presence of the highly
atrophic type 2 fibers early in DM2, remains unsolved. In this study, we did not find
any conclusive explanation to the question of why a subpopulation of fast type 2A
fibers is preferentially vulnerable in DM2.
The new exciting findings of abnormal translation and protein turnover
in DMs are of major interest and require further investigation. With more research
done, it becomes clearer that the DM mutations seem to affect a large number of
| 103
molecular pathways more globally within the muscle cell. On the other hand, if a
therapeutic cure to abolish the dominant gain‐of‐function of the mutant transcript
can be achieved, these known and unknown downstream effects should also be
corrected.
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9. ACKNOWLEDGEMENTS
This thesis work was carried out at the Folkhälsan Institute of Genetics, the
Department of Medical Genetics, University of Helsinki, and Vaasa Central Hospital,
Department of Pathology, during the years 2002 – 2011. The former and present
heads of the departments, professors Anna‐Elina Lehesjoki and Päivi Peltomäki,
Docent Irma Järvelä, as well as Gustav Granroth and Kalle Salo are thanked for
providing excellent facilities and working environments for scientific research. I am
especially grateful to Anna‐Elina Lehesjoki for signing numerous recommendations
for grant applications, Irma Järvelä for serving as the custodian, and Gustav Granroth
and Kalle Salo for their flexibility and positive attitude towards biomedical research.
The following foundations are warmly thanked for funding this thesis
project: the Emil Aaltonen Foundation, the Biomedicum Helsinki Foundation, the
Folkhälsan Research Foundation, the Helsinki University Research Funds, the Liv och
Hälsa Foundation, the Paulo Foundation and the Vaasa Central Hospital District
Research Funds. In addition, the Academy of Finland, Association Française Contre
les Myopathies, the Helsinki University Chancellor´s Travel Grants, the Juselius
Foundation and the Oskar Öflund Foundation are acknowledged for financially
supporting my participation in international congresses, and the neuromuscular
research projects in general.
It has been a privilege to work for many years in the research group of
my supervisor, Professor Bjarne Udd, whose truly enthusiastic attitude towards
science has served as a constant motivating power over the years. Bjarne has an
incredible capability to pull the strings regardless of the ever growing number of
projects he is involved in, not to mention the vast amount of knowledge he
possesses in the field of neuromuscular diseases. In addition, he is the one to remind
us of the fundamental reason what we are doing this research for: the patients.
I own my gratitude to all my co‐authors for their contributions in the
original publications: performing laboratory experiments, providing patient samples
and helping to prepare the manuscripts. Especially, Professor Ralf Krahe and his
research group in Houston, Texas are thanked for introducing a completely new
world of expression profiling, and for their indispensable scientific contribution to
the shared publications. Professor Hannu Kalimo is particularly thanked for fruitful
discussions which significantly improved the readability of the publication III.
| 105
The external reviewers of the thesis manuscript, Docent Katarina Pelin
and Professor Caroline Sewry are thanked for their constructive and useful
comments on the thesis manuscript.
I would like to thank the Vaasa Central Hospital and the Department of
Pathology, where the first part of this study was carried out. Marcus Svartbäck,
Edmond Nushi, Margaretha Pada, Ulla Frejman, Tiina Högström, Camilla Österman,
Tiina Tammenpää, Anette Wikberg, Maikki Mäki and of course Raimo Brax are
thanked for creating a pleasant working atmosphere despite the sometimes
depressing work items. I truly enjoyed working at VKS.
The Udd neuromuscular group co‐workers are warmly thanked: Jaakko
Sarparanta for invaluable help with image processing during the preparation of this
thesis, as well as for serving as an endless source for ideas and answers concerning,
well, any questions; Marjut Ritala and Helena Luque for excellent technical
assistance and positive attitude; pH Jonson for PC‐support, and for always being so
kind and helpful. Also, thanks to my other neuromuscular group‐mates, Peter
Hackman, Merja Soininen, Mark Screen, Sara Hollo, Anni Riihiaho, Jeanette
Holmlund‐Hampf and all the former members of the group at FIG. The Tampere
division: Yinka Raheem, Tiina Suominen, Sanna Huovinen, Sini Penttilä, Satu
Luhtasela, Henna‐Riikka Koskinen and all the others, warm thanks for scientific
collaboration and cheerful travel company. Huge thanks to you all!
My long‐term C305b (“tarkkis”) roommates Saara Tegelberg, Hanna
Västinsalo and Maria Kousi are thanked for sharing the ups and downs of being a
Ph.D. student, assistance in various practical scientific issues, but most of all, for
friendship and off‐science discussions concerning everyday life, sometimes very odd
issues, and everything in between. Without Hanna I would not be able to track
myself in IMDb! Warm thanks also to the more recent member of the gang, Kaisa
Kettunen, and the former inhabitants: Anna‐Kaisa Anttonen and Juha Isosomppi: it
has been a pleasure to share an office with you. In addition, I own my warmest
gratitude to all the people at the Folkhälsan Institute of Genetics, past and present,
for on‐ and off‐science discussions, company during lunch and coffee breaks, and for
creating a cheerful atmosphere at FIG all these years: Mervi Kuronen, Vilma‐Lotta
Lehtokari, Anni Laari, Maria Sandbacka, Anne Saarinen (special thanks for last minute
advice and support!), Otto Manninen, Ann‐Liz Träskelin, Mubashir Hanif, Tarja
Joensuu, Outi Kopra, Anne Polvi, Inken Körber, Olesya Okuneva, Paula Hakala,
Marilotta Turunen (thanks for help with cell lines!), Hanna Hellgren and Teija
Toivonen. Warm thanks also to the former FIG co‐workers Eija Siintola, Kirsi
Alakurtti, Reetta Vainionpää, Kati Donner, Nina Aula, Riikka Hämäläinen, Hanne
Ahola, Laura Mustaniemi, Jukka Kallijärvi and Liina Nevalaita.
106 |
Jaana Welin‐Haapamäki, Marjatta Valkama, Aila Riikonen, Sinikka
Lindh, Stephan Keskinen and Madeleine Avellan are thanked for helping with all
those little practical things over the years.
A huge thanks to all my “old” (no offence!) friends – Heli, Burse, Maarit,
Itu, Leila and Eve, for bringing all the good things in my life over the past years – I
would not be me without you.
Finally, I would like to thank my parents Pirjo and Tapio for always
letting me choose my own path, and supporting me in my decisions. Without the
empathy, care and babysitting help of “mummi” this would not have been possible.
A very special person in my life has been my grandmother Aune, who was almost like
a second mom in my childhood. My younger sister and brother, Eeva and Matti,
thanks for being there always when needed, regardless of the distance. Leena is
thanked for friendship, sharing the ups and downs of being a mom, and the honor of
being Laura´s godmother. Also, my warmest thanks to my relatives‐in‐law: my
mother‐in‐law Raili, for everything that you have done for us; Ulla, Emma and Carla;
Antti and Elina – I have always enjoyed your cheerful company during the holidays.
My husband Pekka is thanked for running the everyday life with me, and sometimes
for me, thus making the finalizing of this long project possible. And of course for
building the house we live in. My precious daughter Veera, you are my greatest
achievement, the funniest person in the world, and the best reminder of what really
is important in life.
Thank you!
Klaukkala, March 2011
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10. REFERENCES
1. Alberts B, Bray D, Lewis J, Raff M, Roberts J, Watson JD. Molecular biology of the cell. Garland Publishing 1994, London, UK.
2. Allen DL, Leinwand LA. Intracellular calcium and myosin isoform transitions. Calcineurin and calcium‐calmodulin kinase pathways regulate preferential activation of the IIa myosin heavy chain promoter. J Biol Chem 2002; 277(47): 45323‐30.
3. Allen DL, Weber JN, Sycuro LK, Leinwand LA. Myocyte enhancer factor‐2 and serum response factor binding elements regulate fast myosin heavy chain transcription in vivo. J Biol Chem 2005; 280(17): 17126‐34.
4. Alwazzan M, Newman E, Hamshere MG, Brook JD. Myotonic dystrophy is associated with a reduced level of RNA from the DMWD allele adjacent to the expanded repeat. Hum Mol Genet 1999; 8(8): 1491‐7.
5. 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(9): 1397‐400.
6. Arahata K, Hoffman EP, Kunkel LM, Ishiura S, Tsukahara T, Ishihara T, Sunohara N, Nonaka I, Ozawa E, Sugita H. Dystrophin diagnosis: Comparison of dystrophin abnormalities by immunofluorescence and immunoblot analyses. Proc Natl Acad Sci U S A 1989; 86(18): 7154‐8.
7. Arambula JF, Ramisetty SR, Baranger AM, Zimmerman SC. A simple ligand that selectively targets CUG trinucleotide repeats and inhibits MBNL protein binding. Proc Natl Acad Sci U S A 2009; 106(38): 16068‐73.
8. Artero R, Prokop A, Paricio N, Begemann G, Pueyo I, Mlodzik M, Perez‐Alonso M, Baylies MK. The muscleblind gene participates in the organization of Z‐bands and epidermal attachments of drosophila muscles and is regulated by Dmef2. Dev Biol 1998; 195(2): 131‐43.
9. Ashizawa T, Baiget M. New nomenclature and DNA testing guidelines for myotonic dystrophy type 1 (DM1). the international myotonic dystrophy consortium (IDMC). Neurology 2000; 54(6): 1218‐21.
10. Ashizawa T, Epstein HF. Ethnic distribution of myotonic dystrophy gene. Lancet 1991; 338(8767): 642‐3.
11. Ashizawa T, Dunne CJ, Dubel JR, Perryman MB, Epstein HF, Boerwinkle E, Hejtmancik JF. Anticipation in myotonic dystrophy. I. Statistical verification based on clinical and haplotype findings Neurology 1992; 42(10): 1871‐7.
12. Ashizawa T, Anvret M, Baiget M, Barcelo JM, Brunner H, Cobo AM, Dallapiccola B, Fenwick RG,Jr, Grandell U, Harley H. Characteristics of intergenerational contractions of the CTG repeat in myotonic dystrophy. Am J Hum Genet 1994; 54(3): 414‐23.
13. Ashizawa T, Dubel JR, Dunne PW, Dunne CJ, Fu YH, Pizzuti A, Caskey CT, Boerwinkle E, Perryman MB, Epstein HF. Anticipation in myotonic dystrophy. II. Complex relationships between clinical findings and structure of the GCT repeat. Neurology 1992; 42(10): 1877‐83.
14. Au Y, Atkinson RA, Guerrini R, Kelly G, Joseph C, Martin SR, Muskett FW, Pallavicini A, Faulkner G, Pastore A. Solution structure of ZASP PDZ domain; implications for sarcomere ultrastructure and enigma family redundancy. Structure (Camb) 2004; 12(4): 611‐22.
15. Auvinen S, Suominen T, Hannonen P, Bachinski LL, Krahe R, Udd B. Myotonic dystrophy type 2 found in two of sixty‐three persons diagnosed as having fibromyalgia. Arthritis Rheum 2008; 58(11): 3627‐31.
108 |
16. Bachinski LL, Sirito M, Bohme M, Baggerly KA, Udd B, Krahe R. Altered MEF2 isoforms in myotonic dystrophy and other neuromuscular disorders. Muscle Nerve 2010; 42: 856‐63.
17. Bachinski LL, Czernuszewicz T, Ramagli LS, Suominen T, Shriver MD, Udd B, Siciliano MJ, Krahe R. Premutation allele pool in myotonic dystrophy type 2. Neurology 2009; 72(6): 490‐7.
18. Bachinski LL, Udd B, Meola G, Sansone V, Bassez G, Eymard B, Thornton CA, Moxley RT, Harper PS, Rogers MT, Jurkat‐Rott K, Lehmann‐Horn F, Wieser T, Gamez J, Navarro C, Bottani A, Kohler A, Shriver MD, Sallinen R, Wessman M, Zhang S, Wright FA, Krahe R. Confirmation of the type 2 myotonic dystrophy (CCTG)n expansion mutation in patients with proximal myotonic myopathy/proximal myotonic dystrophy of different european origins: A single shared haplotype indicates an ancestral founder effect. Am J Hum Genet 2003; 73(4): 835‐48.
19. Bassez G, Radvanyi H, Chapoy E, Authier FJ, Eymard B, Gherardi R. Assessment of in situ hybridization feasibility and efficiency for molecular diagnosis of myotonic dystrophy type 2 (DM2). Myology 2005; (Abstract ): 215.
20. Bassez G, Chapoy E, Bastuji‐Garin S, Radvanyi‐Hoffman H, Authier FJ, Pellissier JF, Eymard B, Gherardi RK. Type 2 myotonic dystrophy can be predicted by the combination of type 2 muscle fiber central nucleation and scattered atrophy. J Neuropathol Exp Neurol 2008; 67(4): 319‐25.
21. Bassez G, Attarian S, Laforet P, Azulay JP, Rouche A, Ferrer X, Urtizberea JA, Pellissier JF, Duboc D, Fardeau M, Pouget J, Eymard B. Proximal myotonial myopathy (PROMM): Clinical and histology study. Rev Neurol (Paris) 2001; 157(2): 209‐18.
22. Beam KG, Horowicz P. Exitation‐contraction coupling in skeletal muscle. In: Engel AG, Franzini‐Armstrong C, editors. Myology. McGraw‐Hill 2004, USA: p. 257‐80.
23. Beard NA, Wei L, Dulhunty AF. Ca(2+) signaling in striated muscle: The elusive roles of triadin, junctin, and calsequestrin. Eur Biophys J 2009; 39(1): 27‐36.
24. Beard NA, Laver DR, Dulhunty AF. Calsequestrin and the calcium release channel of skeletal and cardiac muscle. Prog Biophys Mol Biol 2004; 85(1): 33‐69.
25. Begemann G, Paricio N, Artero R, Kiss I, Perez‐Alonso M, Mlodzik M. Muscleblind, a gene required for photoreceptor differentiation in drosophila, encodes novel nuclear Cys3His‐type zinc‐finger‐containing proteins. Development 1997; 124(21): 4321‐31.
26. Berchtold MW, Brinkmeier H, Muntener M. Calcium ion in skeletal muscle: Its crucial role for muscle function, plasticity, and disease. Physiol Rev 2000; 80: 1215‐65.
27. Berul CI, Maguire CT, Aronovitz MJ, Greenwood J, Miller C, Gehrmann J, Housman D, Mendelsohn ME, Reddy S. DMPK dosage alterations result in atrioventricular conduction abnormalities in a mouse myotonic dystrophy model. J Clin Invest 1999; 103(4): R1‐7.
28. Bhargava A, Fuentes FF. Mutational dynamics of microsatellites. Mol Biotechnol 2010; 44(3): 250‐66.
29. Biral D, Volpe P, Damiani E, Margreth A. Coexistence of two calsequestrin isoforms in rabbit slow‐twitch skeletal muscle fibers. FEBS Lett 1992; 299(2): 175‐8.
30. Bird IM, Dhoot GK, Wilkinson JM. Identification of multiple variants of fast muscle troponin T in the chicken using monoclonal antibodies. Eur J Biochem 1985; 150(3): 517‐25.
31. Bischoff R, Franzini‐Armstrong C. Satellite and stem cells in muscle regeneration. In: Engel AG, Franzini‐Armstrong C, editors. Myology. McGraw‐Hill 2004,USA: p. 66‐86.
32. Blake DJ, Martin‐Rendon E. Intermediate filaments and the function of the dystrophin‐protein complex. Trends Cardiovasc Med 2002; 12(5): 224‐8.
33. Bohn W, Wiegers W, Beuttenmuller M, Traub P. Species‐specific recognition patterns of monoclonal antibodies directed against vimentin. Exp Cell Res 1992; 201(1): 1‐7.
| 109
34. Bonifazi E, Vallo L, Giardina E, Botta A, Novelli G. A long PCR‐based molecular protocol for detecting normal and expanded ZNF9 alleles in myotonic dystrophy type 2. Diagn Mol Pathol 2004; 13(3): 164‐6.
35. Bonifazi E, Gullotta F, Vallo L, Iraci R, Nardone AM, Brunetti E, Botta A, Novelli G. Use of RNA fluorescence in situ hybridization in the prenatal molecular diagnosis of myotonic dystrophy type I. Clin Chem 2006; 52(2): 319‐22.
36. Borycki AG, Emerson CP,Jr. Multiple tissue interactions and signal transduction pathways control somite myogenesis. Curr Top Dev Biol 2000; 48: 165‐224.
37. Botta A, Caldarola S, Vallo L, Bonifazi E, Fruci D, Gullotta F, Massa R, Novelli G, Loreni F. Effect of the [CCTG]n repeat expansion on ZNF9 expression in myotonic dystrophy type II (DM2). Biochim Biophys Acta 2006; 1762(3): 329‐34.
38. Branden C, Tooze J. Introduction to protein structure. Garland Publishing, Inc. 1999, NY, USA.
39. Brandl CJ, deLeon S, Martin DR, MacLennan DH. Adult forms of the Ca2+ATPase of sarcoplasmic reticulum. expression in developing skeletal muscle. J Biol Chem 1987; 262(8): 3768‐74.
40. Brandl CJ, Green NM, Korczak B, MacLennan DH. Two Ca2+ ATPase genes: Homologies and mechanistic implications of deduced amino acid sequences. Cell 1986; 44(4): 597‐607.
41. Brook JD, McCurrach ME, Harley HG, Buckler AJ, Church D, Aburatani H, Hunter K, Stanton VP, Thirion JP, Hudson T. 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(4): 799‐808.
42. Brouwer JR, Willemsen R, Oostra BA. Microsatellite repeat instability and neurological disease. Bioessays 2009; 31(1): 71‐83.
43. Buj‐Bello A, Furling D, Tronchere H, Laporte J, Lerouge T, Butler‐Browne GS, Mandel JL. Muscle‐specific alternative splicing of myotubularin‐related 1 gene is impaired in DM1 muscle cells. Hum Mol Genet 2002; 11(19): 2297‐307.
44. Burk SE, Lytton J, MacLennan DH, Shull GE. cDNA cloning, functional expression, and mRNA tissue distribution of a third organellar Ca2+ pump. J Biol Chem 1989; 264(31): 18561‐8.
45. Burke HB. Discovering patterns in microarray data. Mol Diagn 2000; 5(4): 349‐57. 46. Burnette WN. "Western blotting": Electrophoretic transfer of proteins from sodium
dodecyl sulfate‐‐polyacrylamide gels to unmodified nitrocellulose and radiographic detection with antibody and radioiodinated protein A. Anal Biochem 1981; 112(2): 195‐203.
47. Calabria E, Ciciliot S, Moretti I, Garcia M, Picard A, Dyar KA, Pallafacchina G, Tothova J, Schiaffino S, Murgia M. NFAT isoforms control activity‐dependent muscle fiber type specification. Proc Natl Acad Sci U S A 2009; 106(32): 13335‐40.
48. Calarco JA, Saltzman AL, Ip JY, Blencowe BJ. Technologies for the global discovery and analysis of alternative splicing. Adv Exp Med Biol 2007; 623: 64‐84.
49. Capetanaki Y, Bloch RJ, Kouloumenta A, Mavroidis M, Psarras S. Muscle intermediate filaments and their links to membranes and membranous organelles. Exp Cell Res 2007; 313(10): 2063‐76.
50. Cardani R, Baldassa S, Botta A, Rinaldi F, Novelli G, Mancinelli E, Meola G. Ribonuclear inclusions and MBNL1 nuclear sequestration do not affect myoblast differentiation but alter gene splicing in myotonic dystrophy type 2. Neuromuscul Disord 2009; 19(5): 335‐43.
51. Carson NL. Analysis of repetitive regions in myotonic dystrophy type 1 and 2. Curr Protoc Hum Genet 2009; Chapter 9: Unit 9.6.
110 |
52. Catalli C, Morgante A, Iraci R, Rinaldi F, Botta A, Novelli G. Validation of sensitivity and specificity of tetraplet‐primed PCR (TP‐PCR) in the molecular diagnosis of myotonic dystrophy type 2 (DM2). J Mol Diagn 2010; 12(5): 601‐6.
53. Charlet‐B N, Savkur RS, Singh G, Philips AV, Grice EA, Cooper TA. Loss of the muscle‐specific chloride channel in type 1 myotonic dystrophy due to misregulated alternative splicing. Mol Cell 2002; 10(1): 45‐53.
54. Chen HY, Kathirvel P, Yee WC, Lai PS. Correction of dystrophia myotonica type 1 pre‐mRNA transcripts by artificial trans‐splicing. Gene Ther 2009; 16(2): 211‐7.
55. Chen W, Wang Y, Abe Y, Cheney L, Udd B, Li YP. Haploinsuffciency for Znf9 in Znf9+/‐ mice is associated with multiorgan abnormalities resembling myotonic dystrophy. J Mol Biol 2007; 368(1): 8‐17.
56. Chen W, Liang Y, Deng W, Shimizu K, Ashique AM, Li E, Li YP. The zinc‐finger protein CNBP is required for forebrain formation in the mouse. Development 2003; 130(7): 1367‐79.
57. Coons AH. The development of immunohistochemistry. Ann N Y Acad Sci 1971; 177: 5‐9. 58. Coons AH, Creech HJ, Jones RN. Immunological properties of an antibody containing a
fluorescent group. Proc Soc Exp Biol Med 1941; 47: 200‐2. 59. Cory S, Huang DC, Adams JM. The bcl‐2 family: Roles in cell survival and oncogenesis.
Oncogene 2003; 22(53): 8590‐607. 60. Covault J, Sanes JR. Distribution of N‐CAM in synaptic and extrasynaptic portions of
developing and adult skeletal muscle. J Cell Biol 1986; 102(3): 716‐30. 61. Craig RW, Padron R. Molecular structure of the sarcomere. In: Engel AG, Franzini‐
Armstrong C, editors. Myology. McGraw‐Hill 2004, USA: p. 129‐66. 62. Damiani E, Margreth A. Characterization study of the ryanodine receptor and of
calsequestrin isoforms of mammalian skeletal muscles in relation to fibre types. J Muscle Res Cell Motil 1994; 15(2): 86‐101.
63. 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 U S A 1997; 94(14): 7388‐93.
64. Day JW, Ranum LP. RNA pathogenesis of the myotonic dystrophies. Neuromuscul Disord 2005; 15(1): 5‐16.
65. Day JW, Roelofs R, Leroy B, Pech I, Benzow K, Ranum LP. Clinical and genetic characteristics of a five‐generation family with a novel form of myotonic dystrophy (DM2). Neuromuscul Disord 1999; 9(1): 19‐27.
66. Day JW, Ricker K, Jacobsen JF, Rasmussen LJ, Dick KA, Kress W, Schneider C, Koch MC, Beilman GJ, Harrison AR, Dalton JC, Ranum LP. Myotonic dystrophy type 2: Molecular, diagnostic and clinical spectrum. Neurology 2003; 60(4): 657‐64.
67. de Haro M, Al‐Ramahi I, De Gouyon B, Ukani L, Rosa A, Faustino NA, Ashizawa T, Cooper TA, Botas J. MBNL1 and CUGBP1 modify expanded CUG‐induced toxicity in a drosophila model of myotonic dystrophy type 1. Hum Mol Genet 2006; 15(13): 2138‐45.
68. Deries M, Schweitzer R, Duxson MJ. Developmental fate of the mammalian myotome. Dev Dyn 2010; 239(11): 2898‐910.
69. Draeger A, Weeds AG, Fitzsimons RB. Primary, secondary and tertiary myotubes in developing skeletal muscle: A new approach to the analysis of human myogenesis. J Neurol Sci 1987; 81(1): 19‐43.
70. Du H, Cline MS, Osborne RJ, Tuttle DL, Clark TA, Donohue JP, Hall MP, Shiue L, Swanson MS, Thornton CA, Ares M,Jr. Aberrant alternative splicing and extracellular matrix gene expression in mouse models of myotonic dystrophy. Nat Struct Mol Biol 2010; 17(2): 187‐93.
71. Dubowitz V, Sewry CA. Muscle biopsy ‐ A practical approach. Saunders Elsevier 2007, UK.
| 111
72. Early P, Rogers J, Davis M, Calame K, Bond M, Wall R, Hood L. Two mRNAs can be produced from a single immunoglobulin mu gene by alternative RNA processing pathways. Cell 1980; 20(2): 313‐9.
73. Ecob‐Prince M, Hill M, Brown W. Immunocytochemical demonstration of myosin heavy chain expression in human muscle. J Neurol Sci 1989; 91(1‐2): 71‐8.
74. Edmondson DG, Lyons GE, Martin JF, Olson EN. Mef2 gene expression marks the cardiac and skeletal muscle lineages during mouse embryogenesis. Development 1994; 120(5): 1251‐63.
75. Emerson CP, Hauschka SD. Embryonic origins of skeletal muscles. In: Engel AG, Franzini‐Armstrong C, editors. Myology. McGraw‐Hill 2004, USA: p. 3‐44.
76. Engel AG. The neuromuscular junction. In: Engel AG, Franzini‐Armstrong C, editors. Myology. McGraw‐Hill 2004, USA: p. 325‐72.
77. Engel WK, Cunningham GG. Rapid examination of muscle tissue: an improved trichrome method for fresh‐frozen biopsy sections. Neurology 1963; 13: 919‐23.
78. Fardaei M, Rogers MT, Thorpe HM, Larkin K, Hamshere MG, Harper PS, Brook JD. Three proteins, MBNL, MBLL and MBXL, co‐localize in vivo with nuclear foci of expanded‐repeat transcripts in DM1 and DM2 cells. Hum Mol Genet 2002; 11(7): 805‐14.
79. Faulkner G, Pallavicini A, Formentin E, Comelli A, Ievolella C, Trevisan S, Bortoletto G, Scannapieco P, Salamon M, Mouly V, Valle G, Lanfranchi G. ZASP: A new Z‐band alternatively spliced PDZ‐motif protein. J Cell Biol 1999; 146(2): 465‐75.
80. Flink IL, Morkin E. Alternatively processed isoforms of cellular nucleic acid‐binding protein interact with a suppressor region of the human beta‐myosin heavy chain gene. J Biol Chem 1995; 270(12): 6959‐65.
81. Flink IL, Blitz I, Morkin E. Characterization of cellular nucleic acid binding protein from xenopus laevis: Expression in all three germ layers during early development. Dev Dyn 1998; 211(2): 123‐30.
82. Franzini‐Armstrong C. The membrane systems of muscle cells. In: Engel AG, Franzini‐Armstrong C, editors. Myology. McGraw‐Hill 2004, USA: p. 232‐56.
83. Franzini‐Armstrong C, Horwitz AR. The cytoskeleton: Maintenance of muscle fiber integrity. In: Engel AG, Franzini‐Armstrong C, editors. Myology. McGraw‐Hill 2004, USA. p. 443‐54.
84. Fu YH, Friedman DL, Richards S, Pearlman JA, Gibbs RA, Pizzuti A, Ashizawa T, Perryman MB, Scarlato G, Fenwick RG,Jr. Decreased expression of myotonin‐protein kinase messenger RNA and protein in adult form of myotonic dystrophy. Science 1993; 260(5105): 235‐8.
85. Fu YH, Pizzuti A, Fenwick RG,Jr, King J, Rajnarayan S, Dunne PW, Dubel J, Nasser GA, Ashizawa T, de Jong P. An unstable triplet repeat in a gene related to myotonic muscular dystrophy. Science 1992; 255(5049): 1256‐8.
86. Fu YH, Kuhl DP, Pizzuti A, Pieretti M, Sutcliffe JS, Richards S, Verkerk AJ, Holden JJ, Fenwick RG,Jr, Warren ST. Variation of the CGG repeat at the fragile X site results in genetic instability: Resolution of the Sherman paradox. Cell 1991; 67(6): 1047‐58.
87. Furling D, Marette A, Puymirat J. Insulin‐like growth factor I circumvents defective insulin action in human myotonic dystrophy skeletal muscle cells. Endocrinology 1999; 140(9): 4244‐50.
88. Furling D, Lemieux D, Taneja K, Puymirat J. Decreased levels of myotonic dystrophy protein kinase (DMPK) and delayed differentiation in human myotonic dystrophy myoblasts. Neuromuscul Disord 2001; 11(8): 728‐35.
89. Furling D, Doucet G, Langlois MA, Timchenko L, Belanger E, Cossette L, Puymirat J. Viral vector producing antisense RNA restores myotonic dystrophy myoblast functions. Gene Ther 2003; 10(9): 795‐802.
112 |
90. Furling D, Coiffier L, Mouly V, Barbet JP, St Guily JL, Taneja K, Gourdon G, Junien C, Butler‐Browne GS. Defective satellite cells in congenital myotonic dystrophy. Hum Mol Genet 2001; 10(19): 2079‐87.
91. Futatsugi A, Kuwajima G, Mikoshiba K. Tissue‐specific and developmentally regulated alternative splicing in mouse skeletal muscle ryanodine receptor mRNA. Biochem J 1995; 305(Pt 2): 373‐8.
92. Gall JG, Pardue ML. Formation and detection of RNA‐DNA hybrid molecules in cytological preparations. Proc Natl Acad Sci U S A 1969; 63(2): 378‐83.
93. Garbino A, van Oort RJ, Dixit SS, Landstrom AP, Ackerman MJ, Wehrens XH. Molecular evolution of the junctophilin gene family. Physiol Genomics 2009; 37(3): 175‐86.
94. Gareiss PC, Sobczak K, McNaughton BR, Palde PB, Thornton CA, Miller BL. Dynamic combinatorial selection of molecules capable of inhibiting the (CUG) repeat RNA‐MBNL1 interaction in vitro: Discovery of lead compounds targeting myotonic dystrophy (DM1). J Am Chem Soc 2008; 130(48): 16254‐61.
95. George A, Schneider‐Gold C, Zier S, Reiners K, Sommer C. Musculoskeletal pain in patients with myotonic dystrophy type 2. Arch Neurol 2004; 61(12): 1938‐42.
96. Gerbasi VR, Link AJ. The myotonic dystrophy type 2 protein ZNF9 is part of an ITAF complex that promotes cap‐independent translation. Mol Cell Proteomics 2007; 6(6): 1049‐58.
97. Green KJ, Böhringer M, Gocken T, Jones JCR. ScienceDirect ‐ advances in protein chemistry: Intermediate filament associated proteins. 2005; 2011(2/16/2011).
98. Grifone R, Laclef C, Spitz F, Lopez S, Demignon J, Guidotti JE, Kawakami K, Xu PX, Kelly R, Petrof BJ, Daegelen D, Concordet JP, Maire P. Six1 and Eya1 expression can reprogram adult muscle from the slow‐twitch phenotype into the fast‐twitch phenotype. Mol Cell Biol 2004; 24(14): 6253‐67.
99. Griggs R, Vihola A, Hackman P, Talvinen K, Haravuori H, Faulkner G, Eymard B, Richard I, Selcen D, Engel A, Carpen O, Udd B. Zaspopathy in a large classic late‐onset distal myopathy family. Brain 2007; 130(Pt 6): 1477‐84.
100. Grounds MD, White JD, Rosenthal N, Bogoyevitch MA. The role of stem cells in skeletal and cardiac muscle repair. J Histochem Cytochem 2002; 50(5): 589‐610.
101. Haller RG, Vissing J. Functional evaluation of metabolic myopathies. In: Engel AG, Franzini‐Armstrong C, editors. Myology. McGraw‐Hill 2004, USA: p. 665‐80.
102. 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(1): 59‐61.
103. Hao M, Akrami K, Wei K, De Diego C, Che N, Ku JH, Tidball J, Graves MC, Shieh PB, Chen F. Muscleblind‐like 2 (Mbnl2) ‐deficient mice as a model for myotonic dystrophy. Dev Dyn 2008; 237(2): 403‐10.
104. Harley HG, Rundle SA, Reardon W, Myring J, Crow S, Brook JD, Harper PS, Shaw DJ. Unstable DNA sequence in myotonic dystrophy. Lancet 1992; 339(8802): 1125‐8.
105. Harlow E, Lane D. Antibodies: A laboratory manual. Cold Spring Harbor Laboratory 1988, NY, USA.
106. Harper PS. Myotonic dystrophy. Saunders 2001, London. 107. Havenith MG, Visser R, Schrijvers‐van Schendel JM, Bosman FT. Muscle fiber typing in
routinely processed skeletal muscle with monoclonal antibodies. Histochemistry 1990; 93(5): 497‐9.
108. Hess KR, Zhang W, Baggerly KA, Stivers DN, Coombes KR. Microarrays: Handling the deluge of data and extracting reliable information. Trends Biotechnol 2001; 19(11): 463‐8.
| 113
109. Ho TH, Bundman D, Armstrong DL, Cooper TA. Transgenic mice expressing CUG‐BP1 reproduce splicing mis‐regulation observed in myotonic dystrophy. Hum Mol Genet 2005; 14(11): 1539‐47.
110. Ho TH, Charlet‐B N, Poulos MG, Singh G, Swanson MS, Cooper TA. Muscleblind proteins regulate alternative splicing. EMBO J 2004; 23(15): 3103‐12.
111. Horne Z, Hesketh J. Immunological localization of ribosomes in striated rat muscle. Evidence for myofibrillar association and ontological changes in the subsarcolemmal myofibrillar distribution. Biochem J 1990; 268(1): 231‐6.
112. Houseley JM, Wang Z, Brock GJ, Soloway J, Artero R, Perez‐Alonso M, O'Dell KM, Monckton DG. Myotonic dystrophy associated expanded CUG repeat muscleblind positive ribonuclear foci are not toxic to drosophila. Hum Mol Genet 2005; 14(6): 873‐83.
113. Huang C, Zhou Q, Liang P, Hollander MS, Sheikh F, Li X, Greaser M, Shelton GD, Evans S, Chen J. Characterization and in vivo functional analysis of splice variants of cypher. J Biol Chem 2003; 278(9): 7360‐5.
114. Huichalaf C, Schoser B, Schneider‐Gold C, Jin B, Sarkar P, Timchenko L. Reduction of the rate of protein translation in patients with myotonic dystrophy 2. J Neurosci 2009; 29(28): 9042‐9.
115. Hund E, Jansen O, Koch MC, Ricker K, Fogel W, Niedermaier N, Otto M, Kuhn E, Meinck HM. Proximal myotonic myopathy with MRI white matter abnormalities of the brain. Neurology 1997; 48(1): 33‐7.
116. International Human Genome Sequencing Consortium. Finishing the euchromatic sequence of the human genome. Nature 2004; 431(7011): 931‐45.
117. Jacobs AE, Benders AA, Oosterhof A, Veerkamp JH, van Mier P, Wevers RA, Joosten EM. The calcium homeostasis and the membrane potential of cultured muscle cells from patients with myotonic dystrophy. Biochim Biophys Acta 1990; 1096(1): 14‐9.
118. Jakob S, Corazza N, Diamantis E, Kappeler A, Brunner T. Detection of apoptosis in vivo using antibodies against caspase‐induced neo‐epitopes. Methods 2008; 44(3): 255‐61.
119. Jakubiczka S, Vielhaber S, Kress W, Kupferling P, Reuner U, Kunath B, Wieacker P. Improvement of the diagnostic procedure in proximal myotonic myopathy/myotonic dystrophy type 2. Neurogenetics 2004; 5(1): 55‐9.
120. Jasinska A, Michlewski G, de Mezer M, Sobczak K, Kozlowski P, Napierala M, Krzyzosiak WJ. Structures of trinucleotide repeats in human transcripts and their functional implications. Nucleic Acids Res 2003; 31(19): 5463‐8.
121. Jiang H, Mankodi A, Swanson MS, Moxley RT, Thornton CA. Myotonic dystrophy type 1 is associated with nuclear foci of mutant RNA, sequestration of muscleblind proteins and deregulated alternative splicing in neurons. Hum Mol Genet 2004; 13(24): 3079‐88.
122. Johnson MA, Polgar J, Weightman D, Appleton D. Data on the distribution of fibre types in thirty‐six human muscles. An autopsy study. J Neurol Sci 1973; 18(1): 111‐29.
123. Jones PP, Jiang D, Bolstad J, Hunt DJ, Zhang L, Demaurex N, Chen SR. Endoplasmic reticulum Ca2+ measurements reveal that the cardiac ryanodine receptor mutations linked to cardiac arrhythmia and sudden death alter the threshold for store‐overload‐induced Ca2+ release. Biochem J 2008; 412(1): 171‐8.
124. Kalsotra A, Xiao X, Ward AJ, Castle JC, Johnson JM, Burge CB, Cooper TA. A postnatal switch of CELF and MBNL proteins reprograms alternative splicing in the developing heart. Proc Natl Acad Sci U S A 2008; 105(51): 20333‐8.
125. Kanadia RN, Shin J, Yuan Y, Beattie SG, Wheeler TM, Thornton CA, Swanson MS. Reversal of RNA missplicing and myotonia after muscleblind overexpression in a mouse poly(CUG) model for myotonic dystrophy. Proc Natl Acad Sci U S A 2006; 103(31): 11748‐53.
114 |
126. Kanadia RN, Johnstone KA, Mankodi A, Lungu C, Thornton CA, Esson D, Timmers AM, Hauswirth WW, Swanson MS. A muscleblind knockout model for myotonic dystrophy. Science 2003; 302(5652): 1978‐80.
127. Karpati G, Hilton‐Jones D, Bushby K, Griggs RC. Disorders of voluntary muscle. Cambridge University Press 2010, Cambridge, UK.
128. Karpati G, Charuk J, Carpenter S, Jablecki C, Holland P. Myopathy caused by a deficiency of Ca2+‐adenosine triphosphatase in sarcoplasmic reticulum (brody's disease). Ann Neurol 1986; 20(1): 38‐49.
129. Kassubek J, Juengling FD, Hoffmann S, Rosenbohm A, Kurt A, Jurkat‐Rott K, Steinbach P, Wolf M, Ludolph AC, Lehmann‐Horn F, Lerche H, Weber YG. Quantification of brain atrophy in patients with myotonic dystrophy and proximal myotonic myopathy: A controlled 3‐dimensional magnetic resonance imaging study. Neurosci Lett 2003; 348(2): 73‐6.
130. Kawakami A, Kimura‐Kawakami M, Nomura T, Fujisawa H. Distributions of PAX6 and PAX7 proteins suggest their involvement in both early and late phases of chick brain development. Mech Dev 1997; 66(1‐2): 119‐30.
131. Kierszenbaum A. Histology and cell biology: An introduction to pathology. Mosby Elsevier 2007, PA, USA.
132. Kimura T, Pace SM, Wei L, Beard NA, Dirksen RT, Dulhunty AF. A variably spliced region in the type 1 ryanodine receptor may participate in an inter‐domain interaction. Biochem J 2007; 401(1): 317‐24.
133. Kimura T, Lueck JD, Harvey PJ, Pace SM, Ikemoto N, Casarotto MG, Dirksen RT, Dulhunty AF. Alternative splicing of RyR1 alters the efficacy of skeletal EC coupling. Cell Calcium 2009; 45(3): 264‐74.
134. Kimura T, Nakamori M, Lueck JD, Pouliquin P, Aoike F, Fujimura H, Dirksen RT, Takahashi MP, Dulhunty AF, Sakoda S. Altered mRNA splicing of the skeletal muscle ryanodine receptor and sarcoplasmic/endoplasmic reticulum Ca2+‐ATPase in myotonic dystrophy type 1. Hum Mol Genet 2005; 14(15): 2189‐200.
135. Kirzinger L, Schmidt A, Kornblum C, Schneider‐Gold C, Kress W, Schoser B. Side effects of anesthesia in DM2 as compared to DM1: A comparative retrospective study. Eur J Neurol 2010; 17(6): 842‐5.
136. Klesert TR, Otten AD, Bird TD, Tapscott SJ. Trinucleotide repeat expansion at the myotonic dystrophy locus reduces expression of DMAHP. Nat Genet 1997; 16(4): 402‐6.
137. Kobayashi YM, Alseikhan BA, Jones LR. Localization and characterization of the calsequestrin‐binding domain of triadin 1. evidence for a charged beta‐strand in mediating the protein‐protein interaction. J Biol Chem 2000; 275(23): 17639‐46.
138. Kohler G, Milstein C. Derivation of specific antibody‐producing tissue culture and tumor lines by cell fusion. Eur J Immunol 1976; 6(7): 511‐9.
139. Korade‐Mirnics Z, Tarleton J, Servidei S, Casey RR, Gennarelli M, Pegoraro E, Angelini C, Hoffman EP. Myotonic dystrophy: Tissue‐specific effect of somatic CTG expansions on allele‐specific DMAHP/SIX5 expression. Hum Mol Genet 1999; 8(6): 1017‐23.
140. Kornblum C, Reul J, Kress W, Grothe C, Amanatidis N, Klockgether T, Schroder R. Cranial magnetic resonance imaging in genetically proven myotonic dystrophy type 1 and 2. J Neurol 2004; 251(6): 710‐4.
141. Krahe R, Ashizawa T, Abbruzzese C, Roeder E, Carango P, Giacanelli M, Funanage VL, Siciliano MJ. Effect of myotonic dystrophy trinucleotide repeat expansion on DMPK transcription and processing. Genomics 1995; 28(1): 1‐14.
142. Kress W, Mueller‐Myhsok B, Ricker K, Schneider C, Koch MC, Toyka KV, Mueller CR, Grimm T. Proof of genetic heterogeneity in the proximal myotonic myopathy syndrome (PROMM) and its relationship to myotonic dystrophy type 2 (DM2). Neuromuscul Disord 2000; 10(7): 478‐80.
| 115
143. Kruse B, Wohrle D, Steinbach P, Gal A. Does proximal myotonic myopathy show anticipation? Hum Mutat 2008; 29(8): E100‐2.
144. Kurreck J. Antisense technologies. Improvement through novel chemical modifications. Eur J Biochem 2003; 270(8): 1628‐44.
145. Kuyumcu‐Martinez NM, Wang GS, Cooper TA. Increased steady‐state levels of CUGBP1 in myotonic dystrophy 1 are due to PKC‐mediated hyperphosphorylation. Mol Cell 2007; 28(1): 68‐78.
146. La Spada AR, Wilson EM, Lubahn DB, Harding AE, Fischbeck KH. Androgen receptor gene mutations in X‐linked spinal and bulbar muscular atrophy. Nature 1991; 352(6330): 77‐9.
147. Labeit S, Kolmerer B. Titins: Giant proteins in charge of muscle ultrastructure and elasticity. Science 1995; 270(5234): 293‐6.
148. Laemmli UK. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 1970; 227(5259): 680‐5.
149. Lange S, Xiang F, Yakovenko A, Vihola A, Hackman P, Rostkova E, Kristensen J, Brandmeier B, Franzen G, Hedberg B, Gunnarsson LG, Hughes SM, Marchand S, Sejersen T, Richard I, Edstrom L, Ehler E, Udd B, Gautel M. The kinase domain of titin controls muscle gene expression and protein turnover. Science 2005; 308(5728): 1599‐603.
150. Langlois MA, Boniface C, Wang G, Alluin J, Salvaterra PM, Puymirat J, Rossi JJ, Lee NS. Cytoplasmic and nuclear retained DMPK mRNAs are targets for RNA interference in myotonic dystrophy cells. J Biol Chem 2005; 280(17): 16949‐54.
151. Lavedan C, Hofmann‐Radvanyi H, Shelbourne P, Rabes JP, Duros C, Savoy D, Dehaupas I, Luce S, Johnson K, Junien C. Myotonic dystrophy: Size‐ and sex‐dependent dynamics of CTG meiotic instability, and somatic mosaicism. Am J Hum Genet 1993; 52(5): 875‐83.
152. Le Ber I, Martinez M, Campion D, Laquerriere A, Betard C, Bassez G, Girard C, Saugier‐Veber P, Raux G, Sergeant N, Magnier P, Maisonobe T, Eymard B, Duyckaerts C, Delacourte A, Frebourg T, Hannequin D. A non‐DM1, non‐DM2 multisystem myotonic disorder with frontotemporal dementia: Phenotype and suggestive mapping of the DM3 locus to chromosome 15q21‐24. Brain 2004; 127(Pt 9): 1979‐92.
153. Lee JE, Cooper TA. Pathogenic mechanisms of myotonic dystrophy. Biochem Soc Trans 2009; 37(Pt 6): 1281‐6.
154. Leroy O, Wang J, Maurage CA, Parent M, Cooper T, Buee L, Sergeant N, Andreadis A, Caillet‐Boudin ML. Brain‐specific change in alternative splicing of tau exon 6 in myotonic dystrophy type 1. Biochim Biophys Acta 2006; 1762(4): 460‐7.
155. Lewin B. Genes VII. Oxford University Press and Cell Press 2000, NY, USA. 156. Lin X, Miller JW, Mankodi A, Kanadia RN, Yuan Y, Moxley RT, Swanson MS, Thornton
CA. Failure of MBNL1‐dependent post‐natal splicing transitions in myotonic dystrophy. Hum Mol Genet 2006; 15(13): 2087‐97.
157. Liquori CL, Ikeda Y, Weatherspoon M, Ricker K, Schoser BG, Dalton JC, Day JW, Ranum LP. Myotonic dystrophy type 2: Human founder haplotype and evolutionary conservation of the repeat tract. Am J Hum Genet 2003; 73(4): 849‐62.
158. Liquori CL, Ricker K, Moseley ML, Jacobsen JF, Kress W, Naylor SL, Day JW, Ranum LP. Myotonic dystrophy type 2 caused by a CCTG expansion in intron 1 of ZNF9. Science 2001; 293(5531): 864‐7.
159. Liu J, Fu JD, Siu CW, Li RA. Functional sarcoplasmic reticulum for calcium handling of human embryonic stem cell‐derived cardiomyocytes: Insights for driven maturation. Stem Cells 2007; 25(12): 3038‐44.
116 |
160. Logigian EL, Moxley RT,4th, Blood CL, Barbieri CA, Martens WB, Wiegner AW, Thornton CA, Moxley RT,3rd. Leukocyte CTG repeat length correlates with severity of myotonia in myotonic dystrophy type 1. Neurology 2004; 62(7): 1081‐9.
161. Logigian EL, Martens WB, Moxley RT,4th, McDermott MP, Dilek N, Wiegner AW, Pearson AT, Barbieri CA, Annis CL, Thornton CA, Moxley RT,3rd. Mexiletine is an effective antimyotonia treatment in myotonic dystrophy type 1. Neurology 2010; 74(18): 1441‐8.
162. Luke S, Shepelsky M. FISH: Recent advances and diagnostic aspects. Cell Vis 1998; 5(1): 49‐53.
163. Lyfenko AD, Goonasekera SA, Dirksen RT. Dynamic alterations in myoplasmic Ca2+ in malignant hyperthermia and central core disease. Biochem Biophys Res Commun 2004; 322(4): 1256‐66.
164. MacDonald JR, Hill JD, Tarnopolsky MA. Modafinil reduces excessive somnolence and enhances mood in patients with myotonic dystrophy. Neurology 2002; 59(12): 1876‐80.
165. Machuca‐Tzili L, Thorpe H, Robinson TE, Sewry C, Brook JD. Flies deficient in muscleblind protein model features of myotonic dystrophy with altered splice forms of Z‐band associated transcripts. Hum Genet 2006; 120(4): 487‐99.
166. MacLennan DH, Rice WJ, Green NM. The mechanism of Ca2+ transport by sarco(endo)plasmic reticulum Ca2+‐ATPases. J Biol Chem 1997; 272(46): 28815‐8.
167. Mahadevan M, Tsilfidis C, Sabourin L, Shutler G, Amemiya C, Jansen G, Neville C, Narang M, Barcelo J, O'Hoy K. Myotonic dystrophy mutation: An unstable CTG repeat in the 3' untranslated region of the gene. Science 1992; 255(5049): 1253‐5.
168. Mahadevan MS, Yadava RS, Yu Q, Balijepalli S, Frenzel‐McCardell CD, Bourne TD, Phillips LH. Reversible model of RNA toxicity and cardiac conduction defects in myotonic dystrophy. Nat Genet 2006; 38(9): 1066‐70.
169. Mancini N, Carletti S, Perotti M, Canducci F, Mammarella M, Sampaolo M, Burioni R. Phage display for the production of human monoclonal antibodies against human pathogens. New Microbiol 2004; 27(4): 315‐28.
170. Mankodi A, Lin X, Blaxall BC, Swanson MS, Thornton CA. Nuclear RNA foci in the heart in myotonic dystrophy. Circ Res 2005; 97(11): 1152‐5.
171. Mankodi A, Teng‐Umnuay P, Krym M, Henderson D, Swanson M, Thornton CA. Ribonuclear inclusions in skeletal muscle in myotonic dystrophy types 1 and 2. Ann Neurol 2003; 54(6): 760‐8.
172. Mankodi A, Takahashi MP, Jiang H, Beck CL, Bowers WJ, Moxley RT, Cannon SC, Thornton CA. Expanded CUG repeats trigger aberrant splicing of ClC‐1 chloride channel pre‐mRNA and hyperexcitability of skeletal muscle in myotonic dystrophy. Mol Cell 2002; 10(1): 35‐44.
173. Mankodi A, Logigian E, Callahan L, McClain C, White R, Henderson D, Krym M, Thornton CA. Myotonic dystrophy in transgenic mice expressing an expanded CUG repeat. Science 2000; 289(5485): 1769‐73.
174. Mankodi A, Urbinati CR, Yuan QP, Moxley RT, Sansone V, Krym M, Henderson D, Schalling M, Swanson MS, Thornton CA. Muscleblind localizes to nuclear foci of aberrant RNA in myotonic dystrophy types 1 and 2. Hum Mol Genet 2001; 10(19): 2165‐70.
175. Margolis JM, Schoser BG, Moseley ML, Day JW, Ranum LP. DM2 intronic expansions: Evidence for CCUG accumulation without flanking sequence or effects on ZNF9 mRNA processing or protein expression. Hum Mol Genet 2006; 15(11): 1808‐15.
176. Matlin AJ, Clark F, Smith CW. Understanding alternative splicing: Towards a cellular code. Nat Rev Mol Cell Biol 2005; 6(5): 386‐98.
| 117
177. Maurage CA, Udd B, Ruchoux MM, Vermersch P, Kalimo H, Krahe R, Delacourte A, Sergeant N. Similar brain tau pathology in DM2/PROMM and DM1/Steinert disease. Neurology 2005; 65(10): 1636‐8.
178. Mauro A. Satellite cell of skeletal muscle fibers. J Biophys Biochem Cytol 1961; 9: 493‐5. 179. McKinsey TA, Zhang CL, Olson EN. MEF2: A calcium‐dependent regulator of cell
division, differentiation and death. Trends Biochem Sci 2002; 27(1): 40‐7. 180. Meola G, Sansone V, Radice S, Skradski S, Ptacek L. A family with an unusual myotonic
and myopathic phenotype and no CTG expansion (proximal myotonic myopathy syndrome): A challenge for future molecular studies. Neuromuscul Disord 1996; 6(3): 143‐50.
181. Meola G, Sansone V, Perani D, Scarone S, Cappa S, Dragoni C, Cattaneo E, Cotelli M, Gobbo C, Fazio F, Siciliano G, Mancuso M, Vitelli E, Zhang S, Krahe R, Moxley RT. Executive dysfunction and avoidant personality trait in myotonic dystrophy type 1 (DM‐1) and in proximal myotonic myopathy (PROMM/DM‐2). Neuromuscul Disord 2003; 13(10): 813‐21.
182. Michalowski S, Miller JW, Urbinati CR, Paliouras M, Swanson MS, Griffith J. Visualization of double‐stranded RNAs from the myotonic dystrophy protein kinase gene and interactions with CUG‐binding protein. Nucleic Acids Res 1999; 27(17): 3534‐42.
183. Michel RN, Dunn SE, Chin ER. Calcineurin and skeletal muscle growth. Proc Nutr Soc 2004; 63(2): 341‐9.
184. Micheletti R, Palazzo F, Barassi P, Giacalone G, Ferrandi M, Schiavone A, Moro B, Parodi O, Ferrari P, Bianchi G. Istaroxime, a stimulator of sarcoplasmic reticulum calcium adenosine triphosphatase isoform 2a activity, as a novel therapeutic approach to heart failure. Am J Cardiol 2007; 99(2A): 24A‐32A.
185. Michelotti EF, Tomonaga T, Krutzsch H, Levens D. Cellular nucleic acid binding protein regulates the CT element of the human c‐myc protooncogene. J Biol Chem 1995; 270(16): 9494‐9.
186. Migheli A, Mongini T, Doriguzzi C, Chiado‐Piat L, Piva R, Ugo I, Palmucci L. Muscle apoptosis in humans occurs in normal and denervated muscle, but not in myotonic dystrophy, dystrophinopathies or inflammatory disease. Neurogenetics 1997; 1(2): 81‐7.
187. Miller JW, Urbinati CR, Teng‐Umnuay P, Stenberg MG, Byrne BJ, Thornton CA, Swanson MS. Recruitment of human muscleblind proteins to (CUG)(n) expansions associated with myotonic dystrophy. EMBO J 2000; 19(17): 4439‐48.
188. Minnerop M, Luders E, Specht K, Ruhlmann J, Schneider‐Gold C, Schroder R, Thompson PM, Toga AW, Klockgether T, Kornblum C. Grey and white matter loss along cerebral midline structures in myotonic dystrophy type 2. J Neurol 2008; 255(12): 1904‐9.
189. Molkentin JD, Firulli AB, Black BL, Martin JF, Hustad CM, Copeland N, Jenkins N, Lyons G, Olson EN. MEF2B is a potent transactivator expressed in early myogenic lineages. Mol Cell Biol 1996; 16(7): 3814‐24.
190. Monckton DG, Wong LJ, Ashizawa T, Caskey CT. Somatic mosaicism, germline expansions, germline reversions and intergenerational reductions in myotonic dystrophy males: Small pool PCR analyses. Hum Mol Genet 1995; 4(1): 1‐8.
191. Moore KL, Agur AM, Dalley AF. Essential clinical anatomy ‐ international edition. Lippincott Williams & Wilkins 2010, USA.
192. Moraes KC, Wilusz CJ, Wilusz J. CUG‐BP binds to RNA substrates and recruits PARN deadenylase. RNA 2006; 12(6): 1084‐91.
193. Morton ME, Froehner SC. Monoclonal antibody identifies a 200‐kDa subunit of the dihydropyridine‐sensitive calcium channel. J Biol Chem 1987; 262(25): 11904‐7.
118 |
194. Moxley RT,3rd. Proximal myotonic myopathy: Mini‐review of a recently delineated clinical disorder. Neuromuscul Disord 1996; 6(2): 87‐93.
195. Moxley RT,3rd, Udd B, Ricker K. 54th ENMC international workshop: PROMM (proximal myotonic myopathies) and other proximal myotonic syndromes. 10‐12th October 1997, Naarden, the Netherlands. Neuromuscul Disord 1998; 8(7): 508‐18.
196. Mu X, Brown LD, Liu Y, Schneider MF. Roles of the calcineurin and CaMK signaling pathways in fast‐to‐slow fiber type transformation of cultured adult mouse skeletal muscle fibers. Physiol Genomics 2007; 30(3): 300‐12.
197. Mulders SA, van Engelen BG, Wieringa B, Wansink DG. Molecular therapy in myotonic dystrophy: Focus on RNA gain‐of‐function. Hum Mol Genet 2010; 19(R1): R90‐7.
198. Mulders SA, van den Broek WJ, Wheeler TM, Croes HJ, van Kuik‐Romeijn P, de Kimpe SJ, Furling D, Platenburg GJ, Gourdon G, Thornton CA, Wieringa B, Wansink DG. Triplet‐repeat oligonucleotide‐mediated reversal of RNA toxicity in myotonic dystrophy. Proc Natl Acad Sci U S A 2009; 106(33): 13915‐20.
199. Nakamori M, Sobczak K, Moxley RT,3rd, Thornton CA. Scaled‐down genetic analysis of myotonic dystrophy type 1 and type 2. Neuromuscul Disord 2009; 19(11): 759‐62.
200. Nakamori M, Kimura T, Fujimura H, Takahashi MP, Sakoda S. Altered mRNA splicing of dystrophin in type 1 myotonic dystrophy. Muscle Nerve 2007; 36(2): 251‐7.
201. Nakamori M, Kimura T, Kubota T, Matsumura T, Sumi H, Fujimura H, Takahashi MP, Sakoda S. Aberrantly spliced alpha‐dystrobrevin alters alpha‐syntrophin binding in myotonic dystrophy type 1. Neurology 2008; 70(9): 677‐85.
202. Naya FJ, Olson E. MEF2: A transcriptional target for signaling pathways controlling skeletal muscle growth and differentiation. Curr Opin Cell Biol 1999; 11(6): 683‐8.
203. Newman B, Meola G, O'Donovan DG, Schapira AH, Kingston H. Proximal myotonic myopathy (PROMM) presenting as myotonia during pregnancy. Neuromuscul Disord 1999; 9(3): 144‐9.
204. Nguyen T, Rubinstein NA, Vijayasarathy C, Rome LC, Kaiser LR, Shrager JB, Levine S. Effect of chronic obstructive pulmonary disease on calcium pump ATPase expression in human diaphragm. J Appl Physiol 2005; 98(6): 2004‐10.
205. Odermatt A, Becker S, Khanna VK, Kurzydlowski K, Leisner E, Pette D, MacLennan DH. Sarcolipin regulates the activity of SERCA1, the fast‐twitch skeletal muscle sarcoplasmic reticulum Ca2+‐ATPase. J Biol Chem 1998; 273(20): 12360‐9.
206. Odermatt A, Taschner PE, Khanna VK, Busch HF, Karpati G, Jablecki CK. Mutations in the gene‐encoding SERCA1, the fast‐twitch skeletal muscle sarcoplasmic reticulum Ca2+ ATPase, are associated with Brody disease. Nat Genet 1996; 14: 191‐4.
207. Oishi Y, Ogata T, Ohira Y, Taniguchi K, Roy RR. Calcineurin and heat shock protein 72 in functionally overloaded rat plantaris muscle. Biochem Biophys Res Commun 2005; 330(3): 706‐13.
208. Okazaki K, Holtzer H. Myogenesis: Fusion, myosin synthesis, and the mitotic cycle. Proc Natl Acad Sci U S A 1966; 56(5): 1484‐90.
209. Olive M, Martinez‐Matos JA, Pirretas P, Povedano M, Navarro C, Ferrer I. Expression of myogenic regulatory factors (MRFs) in human neuromuscular disorders. Neuropathol Appl Neurobiol 1997; 23(6): 475‐82.
210. Orengo JP, Chambon P, Metzger D, Mosier DR, Snipes GJ, Cooper TA. Expanded CTG repeats within the DMPK 3' UTR causes severe skeletal muscle wasting in an inducible mouse model for myotonic dystrophy. Proc Natl Acad Sci U S A 2008; 105(7): 2646‐51.
211. Orr HT, Zoghbi HY. Trinucleotide repeat disorders. Annu Rev Neurosci 2007; 30: 575‐621.
212. Osborne RJ, Lin X, Welle S, Sobczak K, O'Rourke JR, Swanson MS, Thornton CA. Transcriptional and post‐transcriptional impact of toxic RNA in myotonic dystrophy. Hum Mol Genet 2009; 18(8): 1471‐81.
| 119
213. Otten AD, Tapscott SJ. Triplet repeat expansion in myotonic dystrophy alters the adjacent chromatin structure. Proc Natl Acad Sci U S A 1995; 92(12): 5465‐9.
214. Ottenheijm CA, Heunks LM, Dekhuijzen RP. Diaphragm adaptations in patients with COPD. Respir Res 2008; 9: 12.
215. Ozasa S, Kimura S, Ito K, Ueno H, Ikezawa M, Matsukura M, Yoshioka K, Araki K, Yamamura KI, Abe K, Niwa H, Miike T. Efficient conversion of ES cells into myogenic lineage using the gene‐inducible system. Biochem Biophys Res Commun 2007; 357(4): 957‐63.
216. Park H, Park IY, Kim E, Youn B, Fields K, Dunker AK, Kang C. Comparing skeletal and cardiac calsequestrin structures and their calcium binding: A proposed mechanism for coupled calcium binding and protein polymerization. J Biol Chem 2004; 279(17): 18026‐33.
217. Patel K, Rossell RJ, Bourne S, Moore SE, Walsh FS, Kemshead JT. Monoclonal antibody UJ13A recognizes the neural cell adhesion molecule (NCAM). Int J Cancer 1989; 44(6): 1062‐8.
218. Pelin K, Wallgren‐Pettersson C. Nebulin‐‐a giant chameleon. Adv Exp Med Biol 2008; 642: 28‐39.
219. Pelletier R, Hamel F, Beaulieu D, Patry L, Haineault C, Tarnopolsky M, Schoser B, Puymirat J. Absence of a differentiation defect in muscle satellite cells from DM2 patients. Neurobiol Dis 2009; 36(1): 181‐90.
220. Pellizzoni L, Lotti F, Maras B, Pierandrei‐Amaldi P. Cellular nucleic acid binding protein binds a conserved region of the 5' UTR of Xenopus laevis ribosomal protein mRNAs. J Mol Biol 1997; 267(2): 264‐75.
221. Penault‐Llorca F, Bilous M, Dowsett M, Hanna W, Osamura RY, Ruschoff J, van de Vijver M. Emerging technologies for assessing HER2 amplification. Am J Clin Pathol 2009; 132(4): 539‐48.
222. Petersen OH, Michalak M, Verkhratsky A. Calcium signalling: Past, present and future. Cell Calcium 2005; 38: 161‐9.
223. Pezzella F, Tse AG, Cordell JL, Pulford KA, Gatter KC, Mason DY. Expression of the bcl‐2 oncogene protein is not specific for the 14;18 chromosomal translocation. Am J Pathol 1990; 137(2): 225‐32.
224. Philips AV, Timchenko LT, Cooper TA. Disruption of splicing regulated by a CUG‐binding protein in myotonic dystrophy. Science 1998; 280(5364): 737‐41.
225. Phillips MS, Fujii J, Khanna VK, DeLeon S, Yokobata K, de Jong PJ, MacLennan DH. The structural organization of the human skeletal muscle ryanodine receptor (RYR1) gene. Genomics 1996; 34(1): 24‐41.
226. Phylactou LA, Darrah C, Wood MJ. Ribozyme‐mediated trans‐splicing of a trinucleotide repeat. Nat Genet 1998; 18(4): 378‐81.
227. Pisani V, Panico MB, Terracciano C, Bonifazi E, Meola G, Novelli G, Bernardi G, Angelini C, Massa R. Preferential central nucleation of type 2 myofibers is an invariable feature of myotonic dystrophy type 2. Muscle Nerve 2008; 38(5): 1405‐11.
228. Polgar J, Johnson MA, Weightman D, Appleton D. Data on fibre size in thirty‐six human muscles, an autopsy study. J Neurol Sci 1973; 19(3): 307‐18.
229. Potthoff MJ, Arnold MA, McAnally J, Richardson JA, Bassel‐Duby R, Olson EN. Regulation of skeletal muscle sarcomere integrity and postnatal muscle function by Mef2c. Mol Cell Biol 2007; 27(23): 8143‐51.
230. Powers‐Martin K, Phillips JK, Biancardi VC, Stern JE. Heterogeneous distribution of basal cyclic guanosine monophosphate within distinct neuronal populations in the hypothalamic paraventricular nucleus. Am J Physiol Regul Integr Comp Physiol 2008; 295(4): R1341‐50.
120 |
231. Proenza C, O'Brien J, Nakai J, Mukherjee S, Allen PD, Beam KG. Identification of a region of RyR1 that participates in allosteric coupling with the alpha(1S) (ca(V)1.1) II‐III loop. J Biol Chem 2002; 277(8): 6530‐5.
232. Radvansky J, Kadasi L. The expanding world of myotonic dystrophies: How can they be detected? Genet Test Mol Biomarkers 2010; 14(6): 733‐41.
233. Raheem O, Huovinen S, Suominen T, Haapasalo H, Udd B. Novel myosin heavy chain immunohistochemical double staining developed for the routine diagnostic separation of I, IIA and IIX fibers. Acta Neuropathol 2010; 119(4): 495‐500.
234. Raheem O, Olufemi SE, Bachinski LL, Vihola A, Sirito M, Holmlund‐Hampf J, Haapasalo H, Li YP, Udd B, Krahe R. Mutant (CCTG)n expansion causes abnormal expression of zinc finger protein 9 in myotonic dystrophy type 2. Am J Pathol 2010; 177(6):3025‐36.
235. Rajavashisth TB, Taylor AK, Andalibi A, Svenson KL, Lusis AJ. Identification of a zinc finger protein that binds to the sterol regulatory element. Science 1989; 245(4918): 640‐3.
236. Ramos‐Vara JA. Technical aspects of immunohistochemistry. Vet Pathol 2005; 42(4): 405‐26.
237. Rantala I, Lounatmaa K. Biologinen valomikroskopia. Helsinki University Press 1998, Helsinki, Finland.
238. Ranum LP, Cooper TA. RNA‐mediated neuromuscular disorders. Annu Rev Neurosci 2006; 29: 259‐77.
239. Ranum LP, Rasmussen PF, Benzow KA, Koob MD, Day JW. Genetic mapping of a second myotonic dystrophy locus. Nat Genet 1998; 19(2): 196‐8.
240. Rauch C, Loughna PT. Static stretch promotes MEF2A nuclear translocation and expression of neonatal myosin heavy chain in C2C12 myocytes in a calcineurin‐ and p38‐dependent manner. Am J Physiol Cell Physiol 2005; 288(3): C593‐605.
241. Relaix F, Rocancourt D, Mansouri A, Buckingham M. A Pax3/Pax7‐dependent population of skeletal muscle progenitor cells. Nature 2005; 435(7044): 948‐53.
242. Ricker K, Koch MC, Lehmann‐Horn F, Pongratz D, Otto M, Heine R, Moxley RT,3rd. Proximal myotonic myopathy: A new dominant disorder with myotonia, muscle weakness, and cataracts. Neurology 1994; 44(8): 1448‐52.
243. Ricker K, Koch MC, Lehmann‐Horn F, Pongratz D, Speich N, Reiners K, Schneider C, Moxley RT,3rd. Proximal myotonic myopathy. clinical features of a multisystem disorder similar to myotonic dystrophy. Arch Neurol 1995; 52(1): 25‐31.
244. Roitt IM, Delves PI. Essential immunology. Blackwell Science 2001, London, UK. 245. Ronnblom A, Forsberg H, Danielsson A. Gastrointestinal symptoms in myotonic
dystrophy. Scand J Gastroenterol 1996; 31(7): 654‐7. 246. Ronnblom A, Andersson S, Hellstrom PM, Danielsson A. Gastric emptying in myotonic
dystrophy. Eur J Clin Invest 2002; 32(8): 570‐4. 247. Rossi AE, Dirksen RT. Sarcoplasmic reticulum: The dynamic calcium governor of muscle.
Muscle Nerve 2006; 33(6): 715‐31. 248. Round JM, Matthews Y, Jones DA. A quick, simple and reliable histochemical method
for ATPase in human muscle preparations. Histochem J 1980; 12(6): 707‐10. 249. Rubinstein NA, Kelly AM. The diversity of muscle fiber types and its origin during
development. In: Engel AG, Franzini‐Armstrong C, editors. Myology. McGraw‐Hill 2004, USA: p. 87‐103.
250. Ryan AM, Matthews E, Hanna MG. Skeletal‐muscle channelopathies: Periodic paralysis and nondystrophic myotonias. Curr Opin Neurol 2007; 20(5): 558‐63.
251. Saito T, Amakusa Y, Kimura T, Yahara O, Aizawa H, Ikeda Y, Day JW, Ranum LP, Ohno K, Matsuura T. Myotonic dystrophy type 2 in japan: Ancestral origin distinct from Caucasian families. Neurogenetics 2008; 9(1): 61‐3.
| 121
252. Sajko S, Kubinova L, Cvetko E, Kreft M, Wernig A, Erzen I. Frequency of M‐cadherin‐stained satellite cells declines in human muscles during aging. J Histochem Cytochem 2004; 52(2): 179‐85.
253. Salisbury E, Sakai K, Schoser B, Huichalaf C, Schneider‐Gold C, Nguyen H, Wang GL, Albrecht JH, Timchenko LT. Ectopic expression of cyclin D3 corrects differentiation of DM1 myoblasts through activation of RNA CUG‐binding protein, CUGBP1. Exp Cell Res 2008; 314(11‐12): 2266‐78.
254. Salisbury E, Schoser B, Schneider‐Gold C, Wang GL, Huichalaf C, Jin B, Sirito M, Sarkar P, Krahe R, Timchenko NA, Timchenko LT. Expression of RNA CCUG repeats dysregulates translation and degradation of proteins in myotonic dystrophy 2 patients. Am J Pathol 2009; 175(2): 748‐62.
255. Salvatori S, Furlan S, Fanin M, Picard A, Pastorello E, Romeo V, Trevisan CP, Angelini C. Comparative transcriptional and biochemical studies in muscle of myotonic dystrophies (DM1 and DM2). Neurol Sci 2009; 30(3): 185‐92.
256. Sammons MA, Antons AK, Bendjennat M, Udd B, Krahe R, Link AJ. ZNF9 activation of IRES‐mediated translation of the human ODC mRNA is decreased in myotonic dystrophy type 2. PLoS One 2010; 5(2): e9301.
257. Sander HW, Tavoulareas GP, Chokroverty S. Heat‐sensitive myotonia in proximal myotonic myopathy. Neurology 1996; 47(4): 956‐62.
258. Sandri M, El Meslemani AH, Sandri C, Schjerling P, Vissing K, Andersen JL, Rossini K, Carraro U, Angelini C. Caspase 3 expression correlates with skeletal muscle apoptosis in duchenne and facioscapulo human muscular dystrophy. A potential target for pharmacological treatment? J Neuropathol Exp Neurol 2001; 60(3): 302‐12.
259. Sanger JW, Sanger JM, Franzini‐Armstrong C. Assembly of the skeletal muscle cell. In: Engel AG, Franzini‐Armstrong C, editors. Myology. McGraw‐Hill 2004, USA: p. 45‐65.
260. Sansone V, Griggs RC, Moxley RT,3rd. Hypothyroidism unmasking proximal myotonic myopathy. Neuromuscul Disord 2000; 10(3): 165‐72.
261. Sarkar PS, Appukuttan B, Han J, Ito Y, Ai C, Tsai W, Chai Y, Stout JT, Reddy S. Heterozygous loss of Six5 in mice is sufficient to cause ocular cataracts. Nat Genet 2000; 25(1): 110‐4.
262. Savkur RS, Philips AV, Cooper TA. Aberrant regulation of insulin receptor alternative splicing is associated with insulin resistance in myotonic dystrophy. Nat Genet 2001; 29(1): 40‐7.
263. Savkur RS, Philips AV, Cooper TA, Dalton JC, Moseley ML, Ranum LP, Day JW. Insulin receptor splicing alteration in myotonic dystrophy type 2. Am J Hum Genet 2004; 74(6): 1309‐13.
264. Schneider C, Ziegler A, Ricker K, Grimm T, Kress W, Reimers CD, Meinck H, Reiners K, Toyka KV. Proximal myotonic myopathy: Evidence for anticipation in families with linkage to chromosome 3q. Neurology 2000; 55(3): 383‐8.
265. Schneider‐Gold C, Beck M, Wessig C, George A, Kele H, Reiners K, Toyka KV. Creatine monohydrate in DM2/PROMM: A double‐blind placebo‐controlled clinical study. Neurology 2003; 60(3): 500‐2.
266. Schoser B, Timchenko L. Myotonic dystrophies 1 and 2: Complex diseases with complex mechanisms. Curr Genomics 2010; 11(2): 77‐90.
267. Schoser BG, Ricker K, Schneider‐Gold C, Hengstenberg C, Durre J, Bultmann B, Kress W, Day JW, Ranum LP. Sudden cardiac death in myotonic dystrophy type 2. Neurology 2004; 63(12): 2402‐4.
268. Schoser BG, Schneider‐Gold C, Kress W, Goebel HH, Reilich P, Koch MC, Pongratz DE, Toyka KV, Lochmuller H, Ricker K. Muscle pathology in 57 patients with myotonic dystrophy type 2. Muscle Nerve 2004; 29(2): 275‐81.
122 |
269. Seale P, Ishibashi J, Scime A, Rudnicki MA. Pax7 is necessary and sufficient for the myogenic specification of CD45+:Sca1+ stem cells from injured muscle. PLoS Biol 2004; 2(5): E130.
270. Selcen D, Engel AG. Mutations in ZASP define a novel form of muscular dystrophy in humans. Ann Neurol 2005; 57(2): 269‐76.
271. Sergeant N, Sablonniere B, Schraen‐Maschke S, Ghestem A, Maurage CA, Wattez A, Vermersch P, Delacourte A. Dysregulation of human brain microtubule‐associated tau mRNA maturation in myotonic dystrophy type 1. Hum Mol Genet 2001; 10(19): 2143‐55.
272. Serrano AL, Murgia M, Pallafacchina G, Calabria E, Coniglio P, Lomo T, Schiaffino S. Calcineurin controls nerve activity‐dependent specification of slow skeletal muscle fibers but not muscle growth. Proc Natl Acad Sci U S A 2001; 98(23): 13108‐13.
273. Seznec H, Lia‐Baldini AS, Duros C, Fouquet C, Lacroix C, Hofmann‐Radvanyi H, Junien C, Gourdon G. Transgenic mice carrying large human genomic sequences with expanded CTG repeat mimic closely the DM CTG repeat intergenerational and somatic instability. Hum Mol Genet 2000; 9(8): 1185‐94.
274. Seznec H, Agbulut O, Sergeant N, Savouret C, Ghestem A, Tabti N, Willer JC, Ourth L, Duros C, Brisson E, Fouquet C, Butler‐Browne G, Delacourte A, Junien C, Gourdon G. Mice transgenic for the human myotonic dystrophy region with expanded CTG repeats display muscular and brain abnormalities. Hum Mol Genet 2001; 10(23): 2717‐26.
275. Simmerman HK, Jones LR. Phospholamban: Protein structure, mechanism of action, and role in cardiac function. Physiol Rev 1998; 78(4): 921‐47.
276. Southern EM. Detection of specific sequences among DNA fragments separated by gel electrophoresis. J Mol Biol 1975; 98(3): 503‐17.
277. Stoll G, von Giesen HJ, Koch MC, Arendt G, Benecke R. Proximal myotonic myopathy syndrome in the absence of trinucleotide repeat expansions. Muscle Nerve 1995; 18(7): 782‐3.
278. Strachan T, Read AP. Human molecular genetics. BIOS Scientific Publishers Ltd, 1999, Oxford, UK.
279. Summerton J. Morpholino antisense oligomers: The case for an RNase H‐independent structural type. Biochim Biophys Acta 1999; 1489(1): 141‐58.
280. Suominen T, Schoser B, Raheem O, Auvinen S, Walter M, Krahe R, Lochmuller H, Kress W, Udd B. High frequency of co‐segregating CLCN1 mutations among myotonic dystrophy type 2 patients from Finland and Germany. J Neurol 2008; 255(11): 1731‐6.
281. Tajsharghi H, Hilton‐Jones D, Raheem O, Saukkonen AM, Oldfors A, Udd B. Human disease caused by loss of fast IIa myosin heavy chain due to recessive MYH2 mutations. Brain 2010; 133(Pt 5): 1451‐9.
282. Takada F, Vander Woude DL, Tong HQ, Thompson TG, Watkins SC, Kunkel LM, Beggs AH. Myozenin: An alpha‐actinin‐ and gamma‐filamin‐binding protein of skeletal muscle Z lines. Proc Natl Acad Sci U S A 2001; 98(4): 1595‐600.
283. 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(6): 995‐1002.
284. Terwilliger JD, Ott J. Handbook of human genetic linkage. The John Hopkins University Press 1994, Baltimore, USA.
285. Tews DS. Muscle‐fiber apoptosis in neuromuscular diseases. Muscle Nerve 2005; 32(4): 443‐58.
286. Thayer MJ, Tapscott SJ, Davis RL, Wright WE, Lassar AB, Weintraub H. Positive autoregulation of the myogenic determination gene MyoD1. Cell 1989; 58(2): 241‐8.
287. Thornton CA, Griggs RC, Moxley RT,3rd. Myotonic dystrophy with no trinucleotide repeat expansion. Ann Neurol 1994; 35(3): 269‐72.
| 123
288. Thornton CA, Johnson K, Moxley RT,3rd. Myotonic dystrophy patients have larger CTG expansions in skeletal muscle than in leukocytes. Ann Neurol 1994; 35(1): 104‐7.
289. Thornton CA, Wymer JP, Simmons Z, McClain C, Moxley RT,3rd. Expansion of the myotonic dystrophy CTG repeat reduces expression of the flanking DMAHP gene. Nat Genet 1997; 16(4): 407‐9.
290. Tieleman AA, den Broeder AA, van de Logt AE, van Engelen BG. Strong association between myotonic dystrophy type 2 and autoimmune diseases. J Neurol Neurosurg Psychiatry 2009; 80(11): 1293‐5.
291. Tieleman AA, Knuijt S, van Vliet J, de Swart BJ, Ensink R, van Engelen BG. Dysphagia is present but mild in myotonic dystrophy type 2. Neuromuscul Disord 2009; 19(3): 196‐8.
292. Tieleman AA, van Vliet J, Jansen JB, van der Kooi AJ, Borm GF, van Engelen BG. Gastrointestinal involvement is frequent in myotonic dystrophy type 2. Neuromuscul Disord 2008; 18(8): 646‐9.
293. Tijo JH, Levan A. The chromosome number in man. Hereditas 1956; 42: 1‐6. 294. Timchenko LT, Salisbury E, Wang GL, Nguyen H, Albrecht JH, Hershey JW, Timchenko
NA. Age‐specific CUGBP1‐eIF2 complex increases translation of CCAAT/enhancer‐binding protein beta in old liver. J Biol Chem 2006; 281(43): 32806‐19.
295. Timchenko NA, Iakova P, Cai ZJ, Smith JR, Timchenko LT. Molecular basis for impaired muscle differentiation in myotonic dystrophy. Mol Cell Biol 2001; 21(20): 6927‐38.
296. Timchenko NA, Patel R, Iakova P, Cai ZJ, Quan L, Timchenko LT. Overexpression of CUG triplet repeat‐binding protein, CUGBP1, in mice inhibits myogenesis. J Biol Chem 2004; 279(13): 13129‐39.
297. Timchenko NA, Cai ZJ, Welm AL, Reddy S, Ashizawa T, Timchenko LT. RNA CUG repeats sequester CUGBP1 and alter protein levels and activity of CUGBP1. J Biol Chem 2001; 276(11): 7820‐6.
298. Turner C, Hilton‐Jones D. The myotonic dystrophies: Diagnosis and management. J Neurol Neurosurg Psychiatry 2010; 81(4): 358‐67.
299. Udd B. Lihastautipotilaita on suomessa yli 10000. Suomen Lääkärilehti 2007; 6: 519‐22. 300. Udd B, Krahe R, Wallgren‐Pettersson C, Falck B, Kalimo H. Proximal myotonic
dystrophy‐‐a family with autosomal dominant muscular dystrophy, cataracts, hearing loss and hypogonadism: Heterogeneity of proximal myotonic syndromes? Neuromuscul Disord 1997; 7(4): 217‐28.
301. Udd B, Meola G, Krahe R, Thornton C, Ranum L, Day J, Bassez G, Ricker K. Report of the 115th ENMC workshop: DM2/PROMM and other myotonic dystrophies. 3rd workshop, 14‐16 February 2003, Naarden, the Netherlands. Neuromuscul Disord 2003; 13(7‐8): 589‐96.
302. Udd B, Meola G, Krahe R, Thornton C, Ranum LP, Bassez G, Kress W, Schoser B, Moxley R. 140th ENMC international workshop: Myotonic dystrophy DM2/PROMM and other myotonic dystrophies with guidelines on management. Neuromuscul Disord 2006; 16(6): 403‐13.
303. van den Broek WJ, Nelen MR, Wansink DG, Coerwinkel MM, te Riele H, Groenen PJ, Wieringa B. Somatic expansion behaviour of the (CTG)n repeat in myotonic dystrophy knock‐in mice is differentially affected by Msh3 and Msh6 mismatch‐repair proteins. Hum Mol Genet 2002; 11(2): 191‐8.
304. Vatta M, Mohapatra B, Jimenez S, Sanchez X, Faulkner G, Perles Z, Sinagra G, Lin JH, Vu TM, Zhou Q, Bowles KR, Di Lenarda A, Schimmenti L, Fox M, Chrisco MA, Murphy RT, McKenna W, Elliott P, Bowles NE, Chen J, Valle G, Towbin JA. Mutations in Cypher/ZASP in patients with dilated cardiomyopathy and left ventricular non‐compaction. J Am Coll Cardiol 2003; 42(11): 2014‐27.
124 |
305. Vicart P, Caron A, Guicheney P, Li Z, Prevost MC, Faure A, Chateau D, Chapon F, Tome F, Dupret JM, Paulin D, Fardeau M. A missense mutation in the alphaB‐crystallin chaperone gene causes a desmin‐related myopathy. Nat Genet 1998; 20(1): 92‐5.
306. Vielhaber S, Jakubiczka S, Gaul C, Schoenfeld MA, Debska‐Vielhaber G, Zierz S, Heinze HJ, Niessen HG, Kaufmann J. Brain 1H magnetic resonance spectroscopic differences in myotonic dystrophy type 2 and type 1. Muscle Nerve 2006; 34(2): 145‐52.
307. Vihola A, Bassez G, Meola G, Zhang S, Haapasalo H, Paetau A, Mancinelli E, Rouche A, Hogrel JY, Laforet P, Maisonobe T, Pellissier JF, Krahe R, Eymard B, Udd B. Histopathological differences of myotonic dystrophy type 1 (DM1) and PROMM/DM2. Neurology 2003; 60(11): 1854‐7.
308. Vlasova IA, Tahoe NM, Fan D, Larsson O, Rattenbacher B, Sternjohn JR, Vasdewani J, Karypis G, Reilly CS, Bitterman PB, Bohjanen PR. Conserved GU‐rich elements mediate mRNA decay by binding to CUG‐binding protein 1. Mol Cell 2008; 29(2): 263‐70.
309. Voet NB, van der Kooi EL, Riphagen II, Lindeman E, van Engelen BG, Geurts AC. Strength training and aerobic exercise training for muscle disease. Cochrane Database Syst Rev 2010; (1): CD003907.
310. Wang GS, Kearney DL, De Biasi M, Taffet G, Cooper TA. Elevation of RNA‐binding protein CUGBP1 is an early event in an inducible heart‐specific mouse model of myotonic dystrophy. J Clin Invest 2007; 117(10): 2802‐11.
311. Wang GS, Kuyumcu‐Martinez MN, Sarma S, Mathur N, Wehrens XH, Cooper TA. PKC inhibition ameliorates the cardiac phenotype in a mouse model of myotonic dystrophy type 1. J Clin Invest 2009; 119(12): 3797‐806.
312. Warden CH, Krisans SK, Purcell‐Huynh D, Leete LM, Daluiski A, Diep A, Taylor BA, Lusis AJ. Mouse cellular nucleic acid binding proteins: A highly conserved family identified by genetic mapping and sequencing. Genomics 1994; 24(1): 14‐9.
313. Warf MB, Nakamori M, Matthys CM, Thornton CA, Berglund JA. Pentamidine reverses the splicing defects associated with myotonic dystrophy. Proc Natl Acad Sci U S A 2009; 106(44): 18551‐6.
314. Watson JD, Crick FH. Genetical implications of the structure of deoxyribonucleic acid. Nature 1953; 171(4361): 964‐7.
315. Weber YG, Roebling R, Kassubek J, Hoffmann S, Rosenbohm A, Wolf M, Steinbach P, Jurkat‐Rott K, Walter H, Reske SN, Lehmann‐Horn F, Mottaghy FM, Lerche H. Comparative analysis of brain structure, metabolism, and cognition in myotonic dystrophy 1 and 2. Neurology 2010; 74(14): 1108‐17.
316. Webster C, Silberstein L, Hays AP, Blau HM. Fast muscle fibers are preferentially affected in Duchenne muscular dystrophy. Cell 1988; 52(4): 503‐13.
317. Wei L, Hanna AD, Beard NA, Dulhunty AF. Unique isoform‐specific properties of calsequestrin in the heart and skeletal muscle. Cell Calcium 2009; 45(5): 474‐84.
318. Wei L, Gallant EM, Dulhunty AF, Beard NA. Junctin and triadin each activate skeletal ryanodine receptors but junctin alone mediates functional interactions with calsequestrin. Int J Biochem Cell Biol 2009; 41(11): 2214‐24.
319. Weingarten TN, Hofer RE, Milone M, Sprung J. Anesthesia and myotonic dystrophy type 2: A case series. Can J Anaesth 2010; 57(3): 248‐55.
320. Wheeler TM, Lueck JD, Swanson MS, Dirksen RT, Thornton CA. Correction of ClC‐1 splicing eliminates chloride channelopathy and myotonia in mouse models of myotonic dystrophy. J Clin Invest 2007; 117(12): 3952‐7.
321. Wheeler TM, Sobczak K, Lueck JD, Osborne RJ, Lin X, Dirksen RT, Thornton CA. Reversal of RNA dominance by displacement of protein sequestered on triplet repeat RNA. Science 2009; 325(5938): 336‐9.
| 125
322. Wieser T, Bonsch D, Eger K, Schulte‐Mattler W, Zierz S. A family with PROMM not linked to the recently mapped PROMM locus DM2. Neuromuscul Disord 2000; 10(2): 141‐3.
323. Wiltgen M, Tilz GP. DNA microarray analysis: Principles and clinical impact. Hematology 2007; 12(4): 271‐87.
324. Winter A, Bornemann A. NCAM, vimentin and neonatal myosin heavy chain expression in human muscle diseases. Neuropathol Appl Neurobiol 1999; 25(5): 417‐24.
325. Wohrle D, Kennerknecht I, Wolf M, Enders H, Schwemmle S, Steinbach P. Heterogeneity of DM kinase repeat expansion in different fetal tissues and further expansion during cell proliferation in vitro: Evidence for a casual involvement of methyl‐directed DNA mismatch repair in triplet repeat stability. Hum Mol Genet 1995; 4(7): 1147‐53.
326. Wong LJ, Ashizawa T, Monckton DG, Caskey CT, Richards CS. Somatic heterogeneity of the CTG repeat in myotonic dystrophy is age and size dependent. Am J Hum Genet 1995; 56(1): 114‐22.
327. Wozniak AC, Kong J, Bock E, Pilipowicz O, Anderson JE. Signaling satellite‐cell activation in skeletal muscle: Markers, models, stretch, and potential alternate pathways. Muscle Nerve 2005; 31(3): 283‐300.
328. Wu QL, Jha PK, Raychowdhury MK, Du Y, Leavis PC, Sarkar S. Isolation and characterization of human fast skeletal beta troponin T cDNA: Comparative sequence analysis of isoforms and insight into the evolution of members of a multigene family. DNA Cell Biol 1994; 13(3): 217‐33.
329. Yotova V, Labuda D, Zietkiewicz E, Gehl D, Lovell A, Lefebvre JF, Bourgeois S, Lemieux‐Blanchard E, Labuda M, Vezina H, Houde L, Tremblay M, Toupance B, Heyer E, Hudson TJ, Laberge C. Anatomy of a founder effect: Myotonic dystrophy in northeastern Quebec. Hum Genet 2005; 117(2‐3): 177‐87.
330. Zarain‐Herzberg A, MacLennan DH, Periasamy M. Characterization of rabbit cardiac sarco(endo)plasmic reticulum Ca2(+)‐ATPase gene. J Biol Chem 1990; 265(8): 4670‐7.
331. Zhang L, Lee JE, Wilusz J, Wilusz CJ. The RNA‐binding protein CUGBP1 regulates stability of tumor necrosis factor mRNA in muscle cells: Implications for myotonic dystrophy. J Biol Chem 2008; 283(33): 22457‐63.