The role of cytoskeletal tropomyosins in skeletal muscle and muscle disease
Nicole Vlahovich
This thesis is submitted in fulfilment of the requirements for the degree of Doctor of Philosophy
The Muscle Development Unit Children’s Medical Research Institute And The School of Natural Sciences University of Western Sydney
April 2007
Acknowledgements
Firstly I would like to thank Dr Edna Hardeman for providing me with the opportunity to be a member of the MDU at CMRI and a part of the muscle Tm project. I very much appreciated her help and support as a supervisor throughout my time as a PhD student, thankyou for all the amazing opportunities.
I would like to thank Dr Anthony Kee for all his help as a co-supervisor and also Professor Peter Gunning, the Tm guru, for much advice and an amazing amount of enthusiasm, which always made me feel like my results were groundbreaking! Thankyou to Dr Galina Schevzov for a great amount of help on the project and a calming nature that was very much appreciated. Also to Emma Kettle whose amazing work on the immunogold EM was invaluable and her friendship cherished. To our collaborators: Rob Parton and Delia Hernandez from IMB QLD, Kathy North and Bili Ilkovski at the NGU at the Children’s Hospital Westmead and David James and Greg Cooney at the Garvan Institute for all your help and advice. Also to Ross Boadle from Westmead Hospital, who allowed me to use the fantastic EM facility and for many helpful discussions.
To all of the MDU, past and present, whose technical advice and friendship over the years has got me though this PhD particularly Lini, Enoch, Mai-Anh and Majid who taught me my precious techniques and Nicole, along with Emma, whose company on coffee breaks was so helpful in times of stress. Also to all of the CMRI, I could not have asked for a better place to complete a PhD; fantastic staff, amazing facilities and a great atmosphere. I am grateful for all the opportunities I have been presented with. Thankyou to the animal house staff for looking after my mice, especially Ben Tuckfield and Shelley Dimech and also Tina Borovina from the ORU, CHW. And thankyou to the admin girls for lunchtime relaxation where I could get away from science for 45 mins, it was a mental health saviour.
A big thankyou to Dr Mark Jones, his advice and encouragement steered me into a PhD in the first place and allowed me to think outside the box. His advice, right from his first class at UWS when I was in second year has been instrumental to my career.
Finally to my family and friends who have made this whole experience possible. I want to thank my extremely supportive family: Carmel, Elie, Jeff, Eileen and Susy. Their efforts have kept me sane (at least partially) and I appreciate how much of my stress they have put up with over the last three years (well more like 10 really!). And to Greg Mitchell, from day one he has been supportive, patient and the best friend I could ask for. Thankyou for believing in me.
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Declaration: The work presented in this thesis is, to the best of my knowledge and belief, original except as acknowledged in the text. I hereby declare that I have not submitted this material, either in full or in part, for a degree at this or any other institution.
______Nicole Vlahovich
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Table of Contents
Section One: General Introduction 1 Chapter One: Literature review and research objectives 2 1.1 Cytoskeletal filament systems 2 1.1.1 Microtubules 2 1.1.2 Intermediate filaments 5 1.1.3 Actin microfilaments 8 Myosin motor proteins 12 Tropomyosins 14 1.2 Muscle cytoarchitecture 19 1.2.1 Filamentous proteins of the sarcomere 21 1.2.2 Cytoskeletal structures in muscle costameres and the Z-LAC 25 Costameres 25 Z-line Associated Cytoskeleton (Z-LAC) 27 1.2.3 The sarcoplasmic reticulum and the T-tubule system 29 1.2.4 Neuromuscular junction (NMJ) and actin 30 1.3 Significant functions of skeletal muscle 32 1.3.1 Muscle contraction 32 1.3.2 The transport of glucose in skeletal muscle 33 1.4 Muscle fibre formation 37 1.4.1 Muscle development 36 1.4.2 The regeneration of muscle fibres 41 1.5 Muscle Disease 42 1.5.1 Muscular dystrophies 43 1.5.2 Congenital myopathies with affected filaments 48 Nemaline myopathy (NM) 48 Actin myopathy (AM) 50 1.6 Research Objectives 52 Section Two: The roles of cytoskeletal tropomyosins in muscle 54 Chapter Two: Cytoskeletal tropomyosins form functionally distinct 55 filaments in skeletal muscle 2.1 Introduction 55 2.2 Materials and Methods 57 2.2.1 Specific materials 57 2.2.2 Animal strains 57 2.2.3 Primary antibodies 57 2.2.4 Secondary antibodies 58 2.2.5 Preparation of tissue samples for western analysis of protein 58 2.2.6 Western blotting analysis 59 2.2.7 Preparation of tissue samples for cryomicrotomy 60 2.2.8 Preparation of tissue samples for semi-thin cryomicrotomy 60 2.2.9 Immuno-staining of muscle sections 61 2.2.10 Immuno-gold labeling and electron microscopy (EM) analysis 61 2.2.11 Muscle fibre isolation and analysis 62 2.2.12 Isolation of membrane components 62 2.2.13 Processing of isolated membranes for protein analysis 63
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2.2.14 Processing of isolated triads fro EM 64 2.2.15 Processing of isolated triads for EM and immuno-labelling 64 2.3 Results 66 2.3.1 Tms are differentially expressed in skeletal muscles 66 2.3.2 Tm isoforms define filaments associated with organelles in muscle fibres 68 2.3.3 Tm4 and Tm5NM1 define discrete actin filament populations at the 72 Z-LAC 2.3.4 Tm4 is associated with the sarcoplasmic reticulum 77 2.4 Discussion 81 2.4.1 Various cytoskeletal Tm isoforms are expressed in skeletal muscle 81 2.4.2 Tm5NM1 and Tm4 define distinct membrane associated structures 82 adjacent to the Z-line in muscle fibres Chapter Three: Tropomyosin 4 indicates repair/remodeling in skeletal 85 muscle disease 3.1 Introduction 85 3.2 Materials and Methods 88 3.2.1 Specific materials 88 3.2.2 Animal strains 88 3.2.3 Human muscle samples 88 3.2.4 Primary antibodies 89 3.2.5 Secondary antibodies 89 3.2.6 Western blotting of human muscle samples 89 3.2.7 Protein preparations to enrich for Tms 89 3.2.8 Immunohistochemistry of human muscle biopsy samples 89 3.2.9 Notexin induced muscle regeneration 90 3.2.10 Mouse hindlimb immobilization 90 3.3 Results 91 3.3.1 Cytoskeletal Tm4 defines two cytoskeletal compartments in normal 91 skeletal muscle 3.3.2 Longitudinal structures defined by Tm4 are evident during myofibrillar 95 assembly and remodeling 3.3.3 Tm4 is an indicator of muscle disease 99 3.4 Discussion 105 3.4.1 Tm4-defined longitudinal filaments reflect the processes of skeletal 105 muscle regeneration and repair 3.4.2 A Tm4/actin cytoskeleton plays a role in the repair of skeletal muscle 106 fibres Chapter Four: The altered expression of Tm5NM1 in skeletal muscle 108 affects membrane morphology and metabolic pathways 4.1 Introduction 108 4.2 Materials and Methods 111 4.2.1 Specific materials 111 4.2.2 Animal strains 111 4.2.3 Primary antibodies 112 4.2.4 Secondary antibodies 112 4.2.5 Oligonucleotides used for RT-PCR 112 4.2.6 Ruthenium Red staining of isolated muscle fibres 112 4.2.7 RNA extraction from muscles for microarray analysis 113 4.2.8 Affymetrix gene chip analysis 113 4.2.9 RNA extraction from muscles from muscles for RT-PCR 114
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4.2.10 Transcription of RNA to cDNA 114 4.2.11 Agarose gel electrophoresis 114 4.2.12 Preparation of GAPDH standards for quantitative PCR 115 4.2.13 Quantitative real time PCR 115 4.3 Results 117 4.3.1 Ablation and over-expression of Tm5NM1 does not impact on levels or 117 localisation of other Tm isoforms 4.3.2 A lack of Tm5NM1 in skeletal muscle causes abnormalities in T-tubule 126 and caveolae morphology 4.3.3 Tm5NM1 knockout and transgenic mice have alterations in gene 130 expression in soleus muscle 4.4 Discussion 138 4.4.1 Tm isoforms from different genes are independently regulated 138 4.4.2 Tm5NM1 plays a role in the organisation of membrane morphology and 139 cellular metabolism Chapter Five: Tropomyosin 5NM1 is involved in glucose transport and 143 adipose tissue proliferation 5.1 Introduction 143 5.2 Materials and Methods 146 5.2.1 Specific materials 146 5.2.2 Animal strains 146 5.2.3 Primary antibodies 146 5.2.4 Secondary antibodies 146 5.2.5 Solubilisation of muscle and adipose tissue in RIPA buffer 146 5.2.6 In vitro analysis of glucose uptake in adipose tissue 147 5.2.7 Wortmannin inhibition of glucose uptake 147 5.2.8 Glucose tolerance testing 148 5.2.9 Analysis of fat pad mass 147 5.3 Results 149 5.3.1 Tm5NM1 co-localises with proteins involved in glucose uptake 149 5.3.2 De-regulation of Tm5NM1 causes changes in glucose uptake and glucose 151 tolerance and knockout and transgenic mice 5.3.3 Tm5/52 transgenic mice have increased body fat 159 5.4 Discussion 161 5.4.1 Tm5NM1-defined actin filaments play a role in insulin-mediated glucose 161 uptake 5.4.2 Tm5NM1 impacts on adipose tissue 163 Section Three: General discussion and future directions 164 Chapter Six: General Discussion 165 6.1 Cytoskeletal Tms segregate to form functionally distinct 167 compartments in skeletal muscle 6.1.1 Tm-defined filament populations segregate with organelles and 167 membrane structures 6.1.2 Tm isoforms are involved in the specification of γ-actin filaments in 168 skeletal muscle 6.1.3 Tm5NM1 plays a unique non-essential role defining γ-actin filaments in 171 association with the T-tubules and sarcolemma 6.1.4 Cytoskeletal Tm filaments are proposed to associate with other actin 174 binding proteins in skeletal muscle
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6.2 A role for Tm4 in the regeneration and repair of muscle tissue 178 6.2.1 A role for Tm4 in organisation of muscle structure during 179 regeneration/repair 6.2.2 A role for Tm4 in muscle disease 181 6.3 A role for Tm5NM1 and other cytoskeletal Tms in intracellular 182 transport 6.3.1 Cytoskeletal Tms are involved in the transport of vesicles to the 182 membrane 6.3.1 Implications of aberrant Tm expression in diseases related to vesicle 184 trafficking 6.4 A role for cytoskeletal Tms in membrane stability and dynamics 186 6.4.1 Cytoskeletal Tms play a role in membrane stabilisation in skeletal muscle 186 6.4.2 A role for cytoskeletal Tms in membrane dynamics associated with 188 vesicle fusion 6.5 A role for Tm5NM1 in the regulation of adipose tissue 189 6.5.1 Tm5NM1 impacts on adipogenicity and PPAR-γ levels 189 6.5.2 A possible role for Tm5NM1 for the treatment of obesity 190 6.6 Future directions and development 191 6.6.1 Generation of Tm4-null mice 192 6.6.2 Further investigation into the role of Tm5NM1 at the T-tubules 193 6.6.3 Further analysis into the role of Tm5NM1 in the translocation of GLUT4 194 6.6.4 Analysis of cytoskeletal Tms in the process of adipodicity 195 6.7 Concluding remarks 196 Reference List 198 Appendix A 225 Appendix B 229 Appendix C 233
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List of Figures
1.1 The microtubule filament contains organised arrays of 3 tubulin monomers. 1.2 Intermediate filaments assemble from single proteins 6 1.3 The actin filament is decorated with protein complexes. 9 1.4 Tropomyosin isoforms are derived from four genes 15 1.5 The cytoarchitecture of the muscle cell 21 1.6 The sarcomere contains ordered arrays of thick and thin 22 filaments 1.7 The costamere links the myofibrils to the sarcolemma 26 1.8 Tropomyosins define the Z-line associated cytoskeleton 28 (Z-LAC) 1.9 The synapse between motor neuron and the muscle fibre is 31 known as the neuromuscular junction (NMJ) 1.10 Tropomyosin and the troponin complex regulates myosin 33 binding to the thin filament 1.11 GLUT4 molecules cycle between the plasma membrane 35 and intracellular storage vesicles 1.12 Myofibrils are built in three steps 40 1.13 Satellite cells drive muscle regeneration 42 1.14 Dystrophic muscle contains regenerating fibres 43 1.15 Muscle samples from nemaline patients contain 49 filamentous accumulations 2.1 Cytoskeletal Tms are expressed in muscle 67 2.2 Cytoskeletal Tms recognised by 9d and αfast9d define a 70 compartment at the sarcolemma 2.3 Tm4 defines filaments at the MTJ and NMJ 71 2.4 Tm5NM1 and Tm4 are components of the Z-LAC 74 2.5 Tm5NM1 colocalises with the T-tubules whereas Tm4 75 does not 2.6 Both Tm5NM1 and Tm4 define -actin filaments at the Z- 76 LAC 2.7 Tm4 colocalises with the terminal cisternae of the SR 79 2.8 Tm4 is closely associated with the SR 80 3.1 Tm4 is expressed in mouse and human skeletal muscles 92 3.2 Tm4 defines novel structures in different mouse muscles 94 3.3 -actin colocalises with Tm4 in both Z-LAC and 96 longitudinal structures 3.4 Tm4 protein levels increase in regenerating and stretched 97 muscle 3.5 Localisation of Tm4 in structures changes during muscle 98 regeneration 3.6 Stretch immobilisation of hindlimb muscles induces the 100 formation of Tm4-defined longitudinal structures 3.7 Tm4 defines longitudinal filaments in mouse models of 101 nemaline myopathy and muscular dystrophy
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3.8 The level of Tm4 protein is elevated in myopathic muscle 103 3.9 Tm4 is present in Z-LAC and longitudinal structures in 104 myopathic human muscle 4.1 Tm5NM1 protein is absent in the skeletal muscle of 9d/89 118 mice and up-regulated in Tm5/52 mice 4.2 Ablation of Tm5NM1 in skeletal muscle has no impact on 120 Tm4 or sarcomeric Tm isoforms 4.3 Increased expression of Tm5NM1 in skeletal muscle has 121 no impact on Tm4 or sarcomeric Tm isoforms 4.4 Exogenous Tm5NM1 localises to the Z-LAC 123 4.5 Tm4 localisation at the Z-LAC is not altered with removal 124 or increased expression of Tm5NM1 4.6 Tm4 localisation at the MTJ is not altered with removal or 125 increased expression of Tm5NM1 4.7 Tm4 does not localise to the T-tubules in the absence of 128 Tm5NM1 4.8 Tm5NM1-null skeletal muscle has defects in T-tubule and 129 caveolae morphology 4.9 Knockout and over-expression of Tm5NM1 is detectable 131 by microarray 4.10 The ablation or up-regulation of Tm5NM1 in soleus 133 muscle impacts on gene products involved in cellular metabolism and transport 4.11 RNA encoding PPAR- is significantly up-regulated in 136 Tm5/52 soleus muscle 4.12 PPAR- protein levels are elevated in Tm5/52 skeletal 137 muscle 5.1 Tm5NM1 colocalises with GLUT4 and syntaxin-4 in 150 skeletal muscle 5.2 Glucose uptake is impaired in Tm5NM1-null adipose 152 tissue 5.3 Glucose clearance is enhanced in Tm5/52 male mice at an 154 early age 5.4 Glucose clearance is enhanced in older female Tm5/52 155 mice 5.5 GLUT4 levels are unchanged in skeletal muscle from 157 Tm5/52 and 9d/89 mice 5.6 GLUT4 levels are unchanged in adipose tissue from 158 Tm5/52 and 9d/89 mice 5.7 Tm5/52 mice have increased abdominal adipose tissue 160 6.1 Cytoskeletal Tms form discrete compartments in skeletal 168 muscle
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List of Abbreviations
ACh Acetylcholine NMJ Neuromuscular junction AM Actin myopathy PBS Phosphate buffered saline APS Ammonium persulfate PBST Phosphate-buffered saline + ATP Adenosine triphosphate Triton X100 PCR Polymerase chain reaction BMD Becker muscular dystrophy PDGF platelet-derived growth bp Base pair factor BSA Bovine serum albumin PI-3K phosphatidylinositol cDNA Complementary DNA 3-kinase CMRI Children’s Medical Research Institute PPAR- peroxisome proliferator CRU Calcium release unit activated receptor-gamma RNA Ribonucleic acid DGC Dystrophin-glycoprotein complex RT-PCR Reverse transcriptase PCR DHPR Dihydropyradine receptor RyR Ryanodine receptor DMD Duchenne muscular dystrophy SEM Standard error of the mean DNA Deoxyribonucleic acid SR Sarcoplasmic reticulum ECM Extracellular matrix TA Tibealis anterior ECU Extensor carpi ulnaris TBS Tris-buffered saline ED Embryonic day tg Transgenic EDL Extensor longus digitorum Tm Tropomyosin EOM Extraocular muscle TTBS Tween20 + tris-buffered saline FDB Flexor digitorum brevis T-tubules Transverse tubules FDP Flexor digitorum profundus wt Wild-type FHC Familial hypertrophic cardiomyopathy Z-LAC Z-line associated cytoskeleton GLUT4 Glucose transport molecule 4 μm Micrometers H&E Haemotoxylin and eosin
IC Intercostal kDa Kilo-Daltons ko Knock-out LD Latissimus dorsi MDU Muscle development unit mdx X-chromosome linked muscular dystrophy MEFs Mouse embryonic fibroblasts MTJ Myotendinous junction Mw Molecular weight nm Nanometers NM Nemaline myopathy x
Manuscripts and Abstracts
Manuscripts
• Sanoudou D, Corbett MA, Han M, Ghoddusi M, Nguyen M-AT, Vlahovich N,
Hardeman EC, Beggs AH (2006). Skeletal muscle repair in a mouse model of
nemaline myopathy. Human Molecular Genetics.
• Schevzov G, Vrhovski B, Vlahovich N, Sudarsan R, Hook J, Joya J, Lemckert
F, Puttur F, Lin J, Hardeman E, Wieczorek D, O’Neill G, Gunning P.
Divergent regulation of the sarcomere and the cytoskeleton. Submitted to
Journal of Biological Chemistry
• Vlahovich N, Schevzov G, Nair-Shaliker V, Ilkovski B, Artap ST, Kee AJ,
North KN, Gunning PW, Hardeman EC. Tropomyosin 4 indicates
repair/remodeling in skeletal muscle disease. Submitted to Cell Motility and
Cytoskeleton.
Conference Abstracts
• N Vlahovich, E Kettle, G Schevzov, V Nair-Shalliker, B Ilkovski, D
Hernandex-Deviez, R Parton, A Kee, K North, P Gunning, E Hardeman.
Tropomyosin 4 defines novel filament systems in normal and diseased muscle.
American Society for Cell Biology Meeting, San Diego, USA. December 10-13,
2006.
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• N Vlahovich, A Kee, E Hardeman, M R Jones. The role of tropomyosins in
muscle and muscle disease- a project seminar. University of Western Sydney
Innovation Conference, Penrith, NSW June 6-8, 2006. First Prize Winner.
• N Vlahovich, J Joya, R Parton, A Kee, G Schevzov, B Vrhovski, P Gunning
and E Hardeman. The non-muscle tropomyosin Tm4 defines novel
microfilament systems in developing myofibres and mature muscle. Hunter
Cellular Biology Meeting, Hunter Valley, NSW, Australia. March 22-24, 2006.
• N Vlahovich, S Artap, A Kee, G Schevzov, P Gunning and E Hardeman. Non-
Sarcomeric Tropomyosin Isoforms Define Novel Filament Compartments in
Skeletal Muscle Fibers. American Society for Cell Biology Meeting, San
Francisco, USA. December 10-14, 2005.
• N Vlahovich, S Elmir, G Schevzov, A Kee, P Gunning, E Hardeman. Non-
muscle tropomyosins define functionally distinct microfilament populations in
developing skeletal muscle. International Society for Developmental Biology,
Sydney, NSW, Australia. September 3-7, 2005.
• N Vlahovich, A Kee, P Gunning, E Hardeman. Tropomyosin isoforms sort to
functionally distinct compartments in skeletal muscle. Hunter Cellular Biology
Meeting, Hunter Valley, NSW, Australia. April 6-8, 2005.
• N Vlahovich, A Kee, P Gunning and E Hardeman. Tropomyosin isoforms sort
to functionally distinct compartments in skeletal muscle. Annual Conference for
the Organization and Expression of the Genome, Lorne, Vic, Australia February
13-17, 2005.
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Abstract
Cells contain an elaborate cytoskeleton which plays a major role in a variety of cellular functions including: maintenance of cell shape and dimension, providing mechanical strength, cell motility, cytokinesis during mitosis and meiosis and intracellular transport. The cell cytoskeleton is made up of three types of protein filaments: the microtubules, the intermediate filaments and the actin cytoskeleton. These components interact with each other to allow the cell to function correctly. When functioning incorrectly, disruptions to many cellular pathway have been observed with mutations in various cytoskeletal proteins causing an assortment of human disease phenotypes.
Characterization of these filament systems in different cell types is essential to the understanding of basic cellular processes and disease causation. The studies in this thesis are concerned with examining specific cytoskeletal tropomyosin-defined actin filament systems in skeletal muscle.
The diversity of the actin filament system relies, in part, on the family of actin binding proteins, the tropomyosins (Tms). There are in excess of forty Tm isoforms found in mammals which are derived from four genes: α, , and δTm. The role of the muscle- specific Tms in striated muscle is well understood, with sarcomeric Tm isoforms functioning as part of the thin filament where it regulates actin-myosin interactions and hence muscle contraction. However, relatively little known about the roles of the many cytoskeletal Tm isoforms.
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Cytoskeletal Tms have been shown to compartmentalise to form functionally distinct
filaments in a range of cell types including neurons (Bryce et al., 2003), fibroblasts
(Percival et al., 2000) and epithelial cells (Dalby-Payne et al., 2003). Recently it has
been shown that cytoskeletal Tm, Tm5NM1 defines a cytoskeletal structure in skeletal muscle called the Z-line associated cytoskeleton (Z-LAC) (Kee et al., 2004). The disruption of this structure by over-expression of an exogenous Tm in transgenic mice results in a muscular dystrophy phenotype, indicating that the Z-LAC plays an important role in maintenance of muscle structure (Kee et al., 2004).
In this study, specific cytoskeletal Tms are further investigated in the context of skeletal
muscle. Here, we examine the expression, localisation and potential function of
cytoskeletal Tm isoforms, focussing on Tm4 (derived from the δ- gene) and Tm5NM1
(derived from the -gene). By western blotting and immuno-staining mouse skeletal muscle, we show that cytoskeletal Tms are expressed in a range of muscles and define separate populations of filaments. These filaments are found in association with a number of muscle structures including the myotendinous junction, neuromuscular junction, the sarcolemma, the t-tubules and the sarcoplasmic reticulum. Of particular interest, Tm4 and Tm5NM1 define cytoskeletal elements in association with the saroplasmic reticulum and T-tubules, respectively, with a separation of less than 90 nm between distinct filamentous populations. The segregation of Tm isoforms indicates a role for Tms in the specification of actin filament function at these cellular regions.
Examination of muscle during development, regeneration and disease revealed that
Tm4 defines a novel cytoskeletal filament system that is orientated perpendicular to the sarcomeric apparatus. Tm4 is up-regulated in both muscular dystrophy and nemaline
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myopathy and also during induced regeneration and focal repair in mouse muscle.
Transition of the Tm4-defined filaments from a predominsnatly longitudinal to a
predominantly Z-LAC orientation is observed during the course of muscle regeneration.
This study shows that Tm4 is a marker of regeneration and repair, in response to
disease, injury and stress in skeletal muscle.
Analysis of Tm5NM1 over-expressing (Tm5/52) and null (9d89) mice revealed that
compensation between Tm genes does not occur in skeletal muscle. We found that the
levels of cytoskeletal Tms derived from the δ-gene are not altered to compensate for the loss or gain of Tm5NM1 and that the localisation of Tm4 is unchanged in skeletal muscle of these mice. Also, excess Tm5NM1 is sorted correctly, localising to the Z-
LAC. This data correlates with evidence from previous investigations which indicates that Tm isoforms are not redundant and are functionally distinct (Gunning et al., 2005).
Transgenic and null mice have also allowed the further elucidation of cytoskeletal Tm function in skeletal muscle. Analyses of these mice suggest a role for Tm5NM1 in glucose regulation in both skeletal muscle and adipose tissue. Tm5NM1 is found to co- localise with members of the glucose transport pathway such as GLUT4 in muscle fibres and analysis of both transgenic and null mice has shown an alteration to glucose uptake in adipose tissue. Taken together these data indicate that Tm5NM1 may play a role in the translocation of the glucose transport molecule GLUT4. In addition to this
Tm5NM1 may play a role in adipose tissue regulation, since over-expressing mice found to have increased white adipose tissue and an up-regulation of a transcriptional regulator of fat-cell formation, PPAR- .
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Section One: General Introduction.
Chapter One: Literature Review Research Objectives
Chapter One
1.1 Cytoskeletal Filament Systems
The cytoskeleton is an elaborate network of filaments responsible for a number of
functions in cells including motility, cell division, establishing cell shape and providing
mechanical strength. Three protein filament systems make up the cytoskeleton;
microtubules, intermediate filaments and microfilaments (also known as the actin
cytoskeleton). Dynamic interactions occur between the three systems to control the
organisation of cell structure.
1.1.1 Microtubules
Tubulin heterodimers comprised of α and tubulin assemble to form microtubules
(Figure 1.1). Microtubules play a major role in organising internal structures within the
cell and facilitate transport, particularly over long range, via motor proteins that run
along the length of the filament (Alberts et al., 1998). Microtubules are dynamic protein structures that undergo growth and reduction. Typically the organisation of
these filaments occurs at the microtubule organising centre (MTOC), a structure close
by the nucleus containing -tubulin and additional protein complexes that nucleate
microtubules at the minus end (Mayer and Jurgens, 2002). The microtubule minus ends
are located in the MTOC, with the filaments extending towards the plasma membrane
and growing at the plus ends (Gadde and Heald, 2004;Becker and Cassimeris, 2005).
Microtubules have a tendency to switch between stages of growth and shortening,
which is referred to as dynamic instability (Janson et al., 2003).
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Chapter One: Literature Review
Figure 1.1 The microtubule filament contains organised arrays of tubulin monomers.
The α, -tubulin heterodimer is the basic unit of the microtubule. These heterodimers are joined end to end to form a protofilament with alternating α and subunits. An helical arrangement of 13 protofilaments forms the cylinder of the microtubule.
Adapted from (Alberts et al., 1998)
The role of the microtubule cytoskeleton in cell division is well-documented.
Microtubules form the bipolar mitotic spindle, which transmits the chromosomes to the daughter cells during mitosis (Alberts et al., 1998). The microtubule nucleation site, the centrosome, plays a major role in the process of mitotic spindle formation serving as a template for nucleation and controlling polarity by anchoring the minus ends and allowing the plus ends to extend outwards (Gadde and Heald, 2004). During metaphase, microtubules facilitate movement by maintaining attachment with the chromosome then growing or shrinking (Howard and Hyman, 2003).
The axon and axonal terminus of neurons rely on the supply of vesicles, proteins, organelles and signalling molecules from the cell body. The microtubules are used in the process of intracellular transports down the axon and are organised in the axon with
Page 3 Chapter One: Literature Review
the plus end toward the axonal terminus and the minus end facing the cell body (Guzik
and Goldstein, 2004). This process relies on two classes of motor proteins that traverse the filament, kinesin and dynein. These proteins provide bi-directional movement with kinesin moving toward the plus end of the microtubule and dynein toward the minus end (Myers et al., 2006). These proteins are responsible for the long-range transport of proteins, lipids, vesicles and organelles from the cell body to the neurite (Guzik and
Goldstein, 2004). The kinesin family of proteins is diverse, with over 40 kinesin genes
found in the mammalian genome (Miki et al., 2005). Kinesins are composed of two
heavy and two light chains, which form a structure consisting of a motor domain that
binds the microtubule to facilitate movement, a central domain which forms dimers and
a tail that binds to cargo proteins (Chevalier-Larsen and Holzbaur, 2006). The motor
domain is well-conserved, however the tail domains have shown diversity among
family members and this reflects the diverse cellular functions of kinesins (Miki et al.,
2005). Dyneins are large complex molecules, composed of two heavy chains that
dimerise and form the motor domain. The ‘stalks’ of the heavy chains also dimerise and
point away from the microtubule to attach to cargo (Goldstein, 2003;Chevalier-Larsen
and Holzbaur, 2006). For the majority of its functions, dynein requires a co-protein,
dynactin, which binds both dynein and the microtubule to form a complex that
increases the efficiency of transport (Chevalier-Larsen and Holzbaur, 2006).
Defects in microtubule-facilitated intracellular transport have been shown to cause
disease. Charcot Marie-Tooth Syndrome Type 2A has been found to be associated with
a haplo-insufficiency of a kinesin (Zhao et al., 2001). This disease is characterised by
weakness and atrophy of distal muscle and mild sensory loss as well as depressed or
absent deep tendon reflexes. It is proposed that this peripheral neuropathy is caused by
Page 4 Chapter One: Literature Review reduced transport of synaptic vesicle precursors down axonal shafts (Zhao et al.,
2001;Goldstein, 2003;Hirokawa and Takemura, 2003). Another kinesin has been linked to hereditary spastic paraplegia, which is characterised by lower limb spasticity and weakness (Reid et al., 2002;Hirokawa and Takemura, 2003) The disruption of microtubule-facilitated axonal transport has been proposed to perturb axonal flow leading to axonal degradation in these patients (Burgunder and Hunziker, 2003).
Mutations in dynein have been shown to cause primary ciliary dyskinesia, characterised by chronic respiratory infections caused by defective mucociliary clearance due to immotile or dysfunctional respiratory cilia (Hornef et al., 2006). Defects in dynein– mediated transport in patients also leads to defects in sperm leading to reduced fertility in some males (Hornef et al., 2006).
1.1.2 Intermediate Filaments
Intermediate filaments, so named as their diameter (10 nm) is between microtubules (23 nm) and microfilaments (6 nm), play an important role in the structural support of the cell (Figure 1.2) (Fuchs and Cleveland, 1998;Coulombe and Wong, 2004). In humans, greater than 60 genes encode intermediate filament proteins which are expressed differentially in all cell types and are quite diverse in their amino acid content.
However, intermediate filament proteins share a common structure, two α-helical rods which assemble into a coiled-coil rod, with well-conserved ends that allow the rods to polymerise into filaments (Fuchs and Cleveland, 1998). These proteins are grouped into five categories of filaments, four of which reside in the cytoplasm of cells and one which localises to the nucleus (Helfand et al., 2004). Included in the intermediate filament family are the keratins, lamins, neurofilament proteins (NFs), vimentin, desmin, synemin and nestin (Coulombe and Wong, 2004;Helfand et al., 2004;Omary et
Page 5 Chapter One: Literature Review
al., 2004). Originally thought to be a static cytoskeletal system, these proteins, and
others, have been found to form dynamic networks and be involved in a wide range of
motile function (Helfand et al., 2004).
Figure 1.2 Intermediate filaments assemble from single proteins
Intermediate filament proteins assemble into dimers that are polar in nature. Dimers associate to form stable tetramers, which align to form protofilaments. These protofilaments are then linked end-to-end and two of these protofilaments are linked to form as protofibrils. Between four and six protofibrils are then linked to form the intermediate filament (Fuchs and Weber, 1994).
Adapted from (Fuchs and Cleveland, 1998)
Intermediate filaments perform essential functions in the cell, one being a role in mechanical support. An example of this is the role of the keratin family in keratinocytes, where filamentous networks extend throughout the cell making up approximately 70% of total protein (Kirfel et al., 2003;Coulombe and Wong, 2004).
The keratin networks extend between sites of cell-cell contact and also points of attachment with the basal lamina forming a structurally strong arrangement, optimised to provide maximum mechanical support for the keratinocyte (Kirfel et al., 2003).
Page 6 Chapter One: Literature Review
Intermediate filaments also function in organisation of organelle location. The desmin cytoskeleton has been proposed to be involved in positioning the mitochondria in skeletal muscle cells. It has been demonstrated that mitochondria are predominantly associated with the intermediate filament system in a number of cell types (Rappaport et al., 1998). Studies have shown that desmin intermediate filaments form a scaffold in muscle that surrounds Z-discs and extends from one Z-line to another potentially associating with other organelles including mitochondria (Capetanaki, 2002). Desmin- null mice provided functional evidence that desmin is involved in the proper organisation of mitochondria in muscle. Mitochondrial distribution, morphology and function was altered in the skeletal muscle of null mice (Milner et al., 2000).
Intermediate filaments have also been implicated in cell movement and migration. The role of the vimentin network in lymphocytes is an example of this type of function.
Under normal conditions, vimentin is organised into a cage-like orientation at the cell periphery (Eckes et al., 1998). Upon induction of chemotaxis, vimentin filaments move
rapidly to the perinuclear region allowing flexibility for movement of the cell during
extravasation (Eckes et al., 1998;Nieminen et al., 2006).
Mutations in and ablations of intermediate filament proteins lead to a variety of human
disorders [reviewed in (Omary et al., 2004)]. Keratin-related disorders result from
mutations in a number of the keratin genes and cause a range of clinical phenotypes that
primarily involve epithelial cells and keratinocytes such as alopecia, hyperkeratosis,
skin lesions and ulcerative colitis (Kirfel et al., 2003). Mutations in the intermediate
filament proteins lamins A and C cause Emery-Dreifuss muscular dystrophy and limb
girdle muscular dystrophies characterised by clinical features such as muscle weakness
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and cardiomyopathy (Decostre et al., 2005;Capell and Collins, 2006). Mutations in lamin genes also lead to a premature aging syndrome known as Hutchinson-Gilford progeria syndrome, characterised by symptoms such as alopecia and premature atherosclerosis (Capell and Collins, 2006). Desmin mutations also result in myopathic phenotypes including dilated cardiomyopathy type II and desmin-related myopathy
(Paulin et al., 2004).
1.1.3 Actin Microfilaments
Actin is a globular protein of approximately 42 kDa, which polymerises to form dynamic filaments. Actin filaments form two twisted α-helices that associate with a wide range of regulatory proteins including tropomyosins and troponins (Figure 1.3).
These filaments are ubiquitous and implicated in a wide-range of cellular functions including cytokinesis and cell motility. In association with tropomyosin (Tm) and the troponin complex they comprise the thin filaments in muscle (Pollard and Cooper,
1986). The dynamic state of actin contributes to the function of this filament system in a range of cellular processes (Wehrle-Haller and Imhof, 2003). Actin is nucleated and then rapidly assembled into filaments with monomers adding to the fast-growing barbed end of the filament, while the pointed, or slow-growing, end of the filament loses actin monomers. Like microtubules, the presence of these ends creates a polarity
on the actin filament (Staiger and Blanchoin, 2006).
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Figure 1.3 The actin filament is decorated with protein complexes.
Actin filaments are composed of monomers. Tropomyosin (Tm) filaments bind the actin groove, with one Tm filament spanning seven actin subunits. Troponin complexes composed of three proteins (TnT, TnC and TnI) bind actin and Tm and are involved in calcium regulation. Adapted from (Gordon et al., 2000)
The human genome contains six known genes that code for actin isoforms, four of
which are expressed exclusively in muscle. The names of the isoforms reflect the
muscle type in which they are predominantly expressed in the adult: α-skeletal, α- cardiac, α-vascular smooth and γ-enteric smooth. The remaining two actin isoforms β-
actin and γ-actin. are cytoskeletal proteins expressed in a wide range of cells [reviewed
in (Rubenstein, 1990)]. The actin isoforms have been shown to perform distinct
functions. Gene knockout analysis of the muscle actins, α-skeletal (Crawford et al.,
2002), α-cardiac (Kumar et al., 1997) and α-vascular smooth (Schildmeyer et al.,
2000) actins, have demonstrated that the sorting of actin isoforms is important in muscle. Removal of the α-cardiac gene resulted in embryonic lethality in the majority
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of mice (Kumar et al., 1997). Expression of ectopic actin, γ-enteric smooth, to
compensate for the lack of α-cardiac actin expression in the hearts of mice allowed
survival of mice to adulthood, however cardiac contractility was compromised. This
indicates that the actin composition is important to specific contractile functions in
muscle (Kumar et al., 1997). In contrast the, cytoskeletal actins have not been as well-
studied. In myoblast cell culture alterations in the expression of cytoskeletal actins
leads to changes in the cell morphology (Schevzov et al., 1992). Elevated expression of
-actin promotes an increase in cell spreading while increasing -actin decreased surface area of myobalsts (Schevzov et al., 1992). These results show that the roles of the cytoskeletal actins are functionally distinct. The ablation of the β-actin gene in mice results in embryonic lethality at embryonic day (ED) 9.5 (Shawlot et al., 1998). A point
mutation present in the -actin gene was identified in patients that caused an amino acid
change from an arginine to a tryptophan at position 183. This mutation caused
developmental malformations and a dystonia syndrome (Procaccio et al., 2006).
Lymphoblast cells derived from patients exhibited alterations in actin depolymerisation
in response to latrunculin B (Procaccio et al., 2006). The ubiquitous removal of γ-actin
is predicted to have a similar effect to the -actin knockout; however, the specific
deletion of γ-actin in skeletal muscles leads to a progressive myopathy (Sonnemann et
al., 2006).
Actin binding proteins are important for the regulation of actin filament organisation
and dynamics. These proteins bind to monomers or directly to filaments to control
polymerisation and disassembly. Proteins such as ADF/cofilin bind to actin monomers
with high affinity and promote the disassembly of filamentous actin from the pointed
end, while gelsolin plays a similar role and binds to barbed ends to sever actin filaments
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(Carlier and Pantaloni, 1997;Small and Resch, 2005). The branching of actin is
controlled by the binding of a protein complex known as the Arp2/3 complex. Arp2/3 is
composed of seven subunits, is thought to mimic an actin dimer and function as a
template for a daughter filament, which branches from the original filament (Goley and
Welch, 2006). Tropomodulin caps the ends of the actin filament and blocks elongation and depolymerisation to create a more stable filament (Weber et al., 1994;Fischer and
Fowler, 2003). This molecule attaches to the actin-Tm complex, binding to the pointed
end of actin filaments and the amino terminus of Tm (Vera et al., 2000;Fujisawa et al.,
2001).
The cytoskeletal actins have been studied extensively in migrating cells due to the
highly regulated filament dynamics required in this region. Studies into this process,
include examining movements of developing neurons and protrusion of neurites,
fibroblast migration and migration of immune cells during chemotaxis (Dent and
Gertler, 2003;Samstag et al., 2003). The leading edge of these cells has been the focus
of studies showing that actin dynamics play a vital role in the membrane protrusions
that drive cell movement (Pollard et al., 2000). Elongation of the barbed end of actin
drives the membrane protrusions such as lamellipodia, membrane ruffles and growth
cones which are necessary for mobility of the cell (Tilney et al., 1981;Pollard and
Borisy, 2003;Atilgan et al., 2005). In the leading edge, the dynamics of actin define two
separate regions: the lamellipodia and the lamella. The lamellipodia comprises the
leading edge of the cell, containing rapidly treadmilling actin, while the lamella is
located between the leading edge and the main cell body and contains long actin
filaments (Vallotton et al., 2004). These regions of the migrating cell are also distinct in
the actin-binding proteins present. The leading edge contains proteins important for
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dynamic actin filaments: Arp2/3 complex, ADF/cofilin and capping proteins. The lamella consists of proteins that stabilise the filaments such as the myosins (Svitkina and Borisy, 1999;Bailly et al., 1999;Gupton et al., 2005;Hotulainen and Lappalainen,
2006). Tropomyosin isoforms are present in both the lamella and the lamellipodia, however the isoforms differ between the regions (Schevzov et al., 2005a).
The actin binding proteins define the structure and function of the actin filament
(Pollard et al., 2000). Two of these families of proteins, myosins and tropomyosins, are described here in detail.
Myosin motor proteins
The myosin family of proteins move along the actin filament facilitated by ATP hydrolysis. This family of proteins are grouped into at least 20 structurally and functionally distinct classes, 12 of which are found in humans (Berg et al.,
2001;Krendel and Mooseker, 2005). These classes can be divided into the conventional myosins (class II) and the unconventional myosins (Suter et al., 2000). Conventional myosins, including those that polymerise to form the basis of the thick filament in striated muscle, are assembled from two heavy and four light chains to form a functional motor protein (Warrick and Spudich, 1987). The C-terminals of the heavy chains form the tail of the molecule, while the N-terminals form the head region which binds actin. The head region consists of two functional heads and also contains the light chains, which form a neck region that functions as a lever during contraction (Craig,
1994). The structure of the unconventional myosins are diverse; some do not polymerise to form filaments while others are single-headed (Dantzig et al., 2006).
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Muscle myosins comprise the thick filaments and are involved in contraction of the
sarcomere (discussed 1.2.1 and 1.3.1). Non-muscle myosins are involved in a range of cellular processes including cell migration, intracellular transport and organelle orientation. Myosin X has been implicated in the formation of filopodia in migrating cells and the recruitment of actin to this region. This myosin is proposed to recruit integrins to the filopodial tip and may elongate actin and link it to these matrix-bound proteins (Zhang et al., 2004). Similarly, non-muscle myosin IIB is associated with cell migration, playing a role in coordination of migration and directionality (Krendel and
Mooseker, 2005). Myosin IIB knockout fibroblasts were found to contain multiple, unstable lamellipodia and while the null cells were able to migrate, they exhibited defects in maintenance of the direction of migration (Lo et al., 2004). This non-muscle isoform is also implicated in retraction of the trailing edge of migrating cells (Maupin et al., 1994;Krendel and Mooseker, 2005).
Mutations in certain myosins have been shown to cause disease. Mutations in cardiac - myosin heavy chain cause some types of familial hypertrophic cardiomyopathy (FHC), which is defined by the presence of a hypertrophied left ventricle. This disease is the most common, identifiable cause of sudden cardiac death in young people (Tardiff,
2005). Defects in an unconventional myosin, myosin-VIIa, cause Usher syndrome type-
I, the most common cause of inherited deafness-blindness in humans (El Amraoui and
Petit, 2005). The faulty myosin protein causes primary defects in the sensory cells of the inner ear and the retina. In the ear, myosin-VIIa is involved in the maintenance of the hair bundles of the sensory cells. Mice expressing a mutated form of myosin-VIIa have disorganised hair bundles and abnormal stereocilia leading to deafness. These
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mice do not show signs of retinal degeneration, but do exhibit mild optical phenotypes
(Petit, 2001;El Amraoui and Petit, 2005).
Tropomyosin
The tropomyosin (Tm) family of actin-binding proteins plays an important role in the function of microfilaments. The Tm family consists of more than 40 isoforms generated by alternate RNA splicing from four genes: the α-, -, - and δ-Tm genes. The phenomena of alternative splicing was first identified in the α-Tm gene, which was shown to express striated muscle, smooth muscle and cytoskeletal isoforms (Wieczorek et al., 1988). Tm dimers bind the α-helical groove of the actin filament in a head-to-tail
orientation and are important components of the actin microfilament cytoskeleton
(Perry, 2001). In skeletal muscle, Tms function as a component of the contractile
apparatus, regulating the actin-myosin interaction to facilitate sarcomeric contraction
[reviewed in (Clark et al., 2002)] (detailed section 1.3.1). Muscle-specific isoforms,
which reside in the thin filament, are generated from three of the four genes (α, and )
and contain the exon 9a from their respective genes (Figure 1.4). The Tm cytokeletal
isoforms function in a variety of different cellular pathways in a range of cell and tissue
types and are developmentally regulated (Gunning et al., 2005). In non-muscle cells the
roles of Tm isoforms are not as well-understood.
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Figure 1.4 Tropomyosin isoforms are derived from four genes.
Greater than forty isoforms are generated from the four Tm genes, some of which are shown in this diagram. The three striated muscle isoforms are generated from the α, and genes (αTmfast, Tm and αTmslow respectively). All other isoforms produced are cytoskeletal Tms. Adapted from (Gunning et al., 2005).
The Tm isoforms have been shown to perform essential functions and the genes are not
redundant [reviewed in (Gunning et al., 2007)]. Gene knockout studies have shown that, when eliminated, both the α and genes cause embryonic lethality (Rethinasamy et al., 1998;Robbins, 1998;Hook et al., 2004). However, not all Tm isoforms are essential. The specific deletion of the 9c exon-containing isoforms from the gene in mice has produced no overt phenotype (Vrhovski et al., 2004). The upregulation of 9a
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containing isoforms to maintain a constant total Tm concentration was observed,
showing that some Tm isoforms may be compensated for by alternative isoforms from
the same gene (Vrhovski et al., 2004).
It has been shown that Tm isoforms are functionally distinct and sort to specific
compartments within the cell (Gunning et al., 2005;Gunning et al., 2007). Studies have shown that Tms protect actin filaments in an isoform specific manner from the severing action of gelsolin (Ishikawa et al., 1989a) and depolymerisation by ADF/cofilin (Ono and Ono, 2002;Bryce et al., 2003). Tropomyosin isoforms have also been shown to
differentially regulate myosin enzymology and mechanochemistry (Fanning et al.,
1994) and also the sorting of myosin motors (Bryce et al., 2003). This
compartmentalisation function of Tm isoforms is best described in neuronal cells
[reviewed in (Gunning et al., 2005)]. During neuronal development specific isoforms
sort to the axonal shaft and growth cone. Upon differentiation additional isoforms are
expressed and Tms re-localise to form distinct compartments in the axon, soma,
dendrite and presynaptic terminal (Weinberger et al., 1993;Had et al., 1993;Hannan et al., 1995;Schevzov et al., 1997;Hannan et al., 1998). This compartmentalisation is not restricted to neuronal cells. During the G1 phase of the fibroblast cell cycle, Tm isoforms align to stress fibres at different stages; whereas, Tm5NM-2 remains associated with the Golgi (Percival et al., 2004) and additional isoforms containing the
-9a exon localise to a perinuclear compartment (Schevzov et al., 2005b). Similar
phenomena have been described in epithelial cells, where Tm5a and 5b localise to the
apical surface of cultured cells, Tm2 and Tm3 locate to the basolateral membrane and
isoforms generated from the gene are found in the cytoplasm (Dalby-Payne et al.,
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2003). Sorting of functionally distinct Tm isoforms provides a mechanism for the
spatial regulation of actin filament function.
The Tm isoforms then convey specific properties to the actin filament, as the various isoforms have different affinities for actin binding proteins. An example of this phenomenon is the interaction of different Tm isoforms with myosin motors. Increased
expression of Tm5NM1 in neuronal cells was found to increase the recruitment of
myosin IIB to the growth cone area, while the over-expression of TmBr3 decreased the
activity of myosin in this region (Bryce et al., 2003). In addition to this, the Tm
isoforms and actin binding proteins ADF/cofilin have been shown to compete for actin
binding (Bernstein and Bamburg, 1982). Bryce and colleagues (2003) demonstrated
that increased expression of Tm5NM1 resulted in the elimination of ADF/cofilin from
the growth cone region where the Tm isoform localised. In contrast to this, the
increased expression of TmBr3 recruited the depolymerisation factor to the
lamellipodium. This study also demonstrated that ADF and TmBr3 exist on the same
filament.
The Tm isoforms provide diversity for actin filaments in a range of processes. The
known functions of the Tm proteins is not yet complete and further investigation into
the various isoforms and their unique properties will elucidate the mechanisms by
which the actin cytoskeleton performs its diverse functions in all cell types (Gunning et
al., 2007).
The sorting of Tm isoforms to specific cellular regions is regulated intrinsically. This
sorting has been described as a ‘molecular sink model’, whereby Tm isoforms
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accumulate at specific sites on the actin filament, depending on the structural integrity
at that site (Gunning et al., 2005). The disruption of actin filaments by exposure of cells to depolymerisation drugs abolishes the sorting of Tms in the growth cones of neuronal cells, while drug washout re-establishes the sorting (Schevzov et al., 1997). This indicates that the state of the actin filament, for example in actin monomers, short filaments or long and bundled filaments, influences Tm binding. The sorting of Tm isoforms to specific regions of the cell is dependent on the composition of the actin filament structures. The specificity of isoforms for certain structures is shown by the over-expression of Tm5NM1 in transgenic mice. In isolated neurons, Tm5NM1 was found to sort to the growth cones, mimicking the organisation of the endogenous
Tm5NM1, suggesting the composition of the actin filaments in the region are most compatible with this Tm isoform (Schevzov et al., 2005a;Gunning et al., 2007). Factors
that may affect the localisation of a Tm to a specific cellular region include the actin
isoform available, the dynamic state of the actin filament, actin binding and capping
proteins available and the myosin isoforms (Gunning et al., 2007).
Mutations in Tms underpin specific human diseases including FHC, nemaline
myopathy, cancer and ulcerative colitis. Mutations in the α-Tm gene cause a rare form
of FHC which is clinically heterogenous (Jongbloed et al., 2003). Typically mutations
in α-Tm are milder than the FHC caused by mutations in the -myosin heavy chain,
(Coviello et al., 1997), however a novel mutation found in a Finnish family caused
severe FHC leading to death at an early age (Jongbloed et al., 2003). Mutations in the
α-Tm are also associated with dilated cardiomyopathy (Olson et al., 2001). Mutations
in Tm and αTmslow, encoded by the and -Tm genes, respectively, cause nemaline
myopathy (detailed 1.5.2). This disease is characterised by the presence of electron-
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dense accumulations of thin filament protein in skeletal muscle leading to muscle weakness (Laing and Nowak, 2005). The involvement of Tm5 in ulcerative coliltis, characterised by diarrhoea, rectal bleeding and at times, weight loss and fever, is not due to a mutation in the gene, but an autoantibody response to Tm5 protein (Das et al.,
1993;Taniguchi et al., 2001). It was found that 95% of patients had antibodies to Tm5 in their blood serum, suggesting that an autoimmune reaction to Tm5 in intestinal epithelial cells of the digestive system may be the cause of disease symptoms (Das et al., 1993). The involvement of Tm isoforms in cancer has also been documented.
Changes in the expression levels of Tms has been detected in cancerous cells [reviewed
in (Gunning et al., 2007)]. The expression of high molecular weight (HMW) Tms is
decreased in a number of tumour cell lines as well as in primary tumours from both
breast cancer and neuroblastoma (Franzen et al., 1996;Yager et al., 2003).
1.2 Muscle Cytoarchitecture
The skeletal muscle fibre is an elongated multi-nucleated cell approximately 10 to
100μM in diameter and up to several centimetres in length in humans (Figure 1.5).
Each muscle fibre is attached to the tendon at the myotendinous junction (MTJ), with
the tendon attaching the muscle to the bone. Muscle fibres are densely packed with
myofibrils, each myofibril containing arrays of contractile apparatus know as the
sarcomeres (Figure 1.5). Sarcomeres contain interdigitating myosin thick filaments and
actin thin filaments bordered by a Z-line, which defines the boundary of the sarcomere
and provides an anchoring point for both actin thin filaments and sarcomeric associated
proteins such as nebulin and titin (Littlefield and Fowler, 1998). Muscle contraction is
controlled by signals from nerves. Axons from the motor neuron attach to the muscle
fibre at the neuromuscular junction (NMJ). Signals from the neuron are potentiated
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along the sarcolemma, the plasma membrane of the muscle fibre, and travel down invaginations of the plasma membrane, known as transverse tubules (T-tubules). These structures penetrate the muscle fibre at the level of the Z-line. At the Z-line, T-tubules interact with the sarcoplasmic reticulum (SR), a membranous structure which surrounds each of the myofibrils and interacts with the T-tubules to form the triad junction which
function to propagate action potentials and facilitate contraction of the sarcomeres
(Alberts et al., 1998;Martini et al., 2001). Two regions of the SR exist in muscle.
Longitudinal SR extends along the length of the sarcomere where it functions to re- gather free calcium ions from the surrounding area. The horizontal terminal cisternae,
found at the Z-line releases calcium to bind the myofibrils and trigger contraction
(Craig, 1994;Alberts et al., 1998).
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Figure 1.5 The cytoarchitecture of the muscle cell.
Skeletal muscle is composed of elongated, multi-nucleated cells known as muscle fibres. Each fibre is bound by the plasma membrane, the sarcolemma, and contains myofibrils, composed of sarcomeres. The sarcomere is the contractile apparatus of the muscle consisting of interdigitating thick and thin filaments bordered by the Z-line. The sliding of these filaments over one another is the basis of muscle contraction. Invaginations of the sarcolemma at the level of the Z-line, known as the transverse tubules (T-tubules), allow action potentials to propagate along the membrane and penetrate the cell and activate the sarcoplasmic reticulum (SR). The membranous SR surrounds each myofibril and is responsible for calcium storage and release.
[Adapted from (Davies and Nowak, 2006)]
1.2.1 Filamentous proteins of the sarcomere.
Sarcomeric structures contain parallel bands of actin thin filaments interdigitating with
myosin thick filaments that overlap at the A-band to facilitate contraction (Figure 1.6).
The actin molecule is the most abundant protein in striated muscle and with Tm and troponin complexes, forms the basis of the thin filament (Craig, 1994;Clark et al.,
2002). In the skeletal muscle contractile apparatus, α-skeletal actin is the predominant
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actin isoform present. Similarly, of the many Tm isoforms, only three are found in the
thin filament: αTmfast, αTmslow and Tm. The Tms function in both contraction and also in the stabilization of the thin filament, increasing stiffness and inhibiting the fragmentation of actin (Wegner, 1982). The troponin complex consists of three proteins
(TnC, TnI and TnT) which function with Tm to facilitate contraction (discussed in
1.3.1). The polarized filament is capped at the pointed end by tropomodulin (Fowler et al., 1993), which interacts with Tm, and at the barbed end by CapZ, which is situated in
the Z-line (Papa et al., 1999). These capping proteins ensure the filament length is
unchanged, blocking actin elongation and shortening.
The thin filaments are anchored at the Z-line, which defines the boundaries of the
sarcomere and appears as an electron dense structure at the centre of the I-band (Figure
1.6). Also known as Z-discs, these structures cross-link the thin filaments into a three
dimensional lattice (Clark et al., 2002). The major protein component of the Z-line is α-
actinin which assembles into homodimers and binds to actin, cross-linking and
organising the thin filaments. A number of other proteins make up the scaffold of the Z-
line such as the actin capping protein Cap Z and -filamin which interact with both
actin and α-actinin (Papa et al., 1999;van der Ven et al., 2000;Faulkner et al., 2001). Z-
lines from adjacent sarcomeres are linked by desmin filaments forming the ordered
array of sarcomeres found in the myofibrils (Granger and Lazarides, 1978). Desmin
also links the Z-line to cytoskeletal structures attached to the sarcolemma (Capetanaki
and Milner, 1998). The Z-line also has been implicated as a point of signal transduction
in muscle, with a number of signalling molecules localising to, and interacting with the
structural proteins of the Z-line [reviewed (Frank et al., 2006)].
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Myosin thick filaments are the motor proteins of the sarcomere (Figure 1.6). Each muscle myosin is a hexamer, composed of two heavy chains and four light chains
(Craig, 1994). Multiple isoforms of myosin heavy chains (MHC) are found in muscles conferring different contractile properties on the sarcomere. The MHCII isoforms are found in fast skeletal muscle fibres, while the MHCI isoform is found in the slow muscles. Additional isoforms are found in developing muscle (Bottinelli et al.,
1999;Reggiani et al., 2000). Myosin motor proteins assemble to form the thick filament, containing the m-band; the central zone of the myosin filaments composed of the myosin tails (Agarkova et al., 2003). Proteins associated with the thick filaments have been identified as important components of the sarcomere, implicated in assembly of the thick filaments and regulation of contraction (Clark et al., 2002). Two structurally similar proteins, myomesin and M-protein, bind to the central zone of the myosin thick filaments and to titin, participating in the cytoskeletal structure of the myofibril (Obermann et al., 1997;Agarkova et al., 2003). It is proposed that myomesin may link the myosin, playing a role similar to that of α-actinin in the Z-line, and function as a molecular spring in this region (Schoenauer et al., 2005).
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T-tubule SR Mitochondria
Figure 1.6 The sarcomere contains ordered arrays of thick and thin filaments.
One sarcomere is defined as the segment of contractile apparatus between two neighbouring Z-lines. Actin filaments, bound to the Z-line, comprise the I-band, while the area of overlap between thin and thick filaments is called the A-band. The centre of the sarcomere, the m-line, contains the tails of the myosin thick filaments. Mitochondria, T-tubules and SR membranes are found in the region between adjacent sarcomeres.
Schematic adapted from (Craig, 1994) and EM image kindly provided by Emma Kettle
Two giant proteins, titin and nebulin, are also important members of the sarcomere.
Titin, the third most abundant protein in muscle after actin and myosin, spans half the sarcomere and at approximately four mega-Daltons is the largest protein found in nature as yet (Tskhovrebova and Trinick, 2003). At the N-terminus, the titin molecule
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interacts with the Z-line, binding with α-actinin and another protein T-cap, while the C-
terminus region of titin is found at the m-line (Obermann et al., 1997;Gregorio et al.,
1998;Young et al., 1998). Nebulin extends along the length of the thin filament in the sarcomere, with the C-terminus interacting with CapZ in the Z-line and the N-terminal region extending towards the m-line (Wang and Wright, 1988;Witt et al., 2006). It has
been proposed that these proteins have roles in organisation and maintenance of the
sarcomeric structure (Horowits et al., 1986;Clark et al., 2002). It is suggested that nebulin serves as a ‘molecular ruler’, regulating thin filament length and also z-disc
structure, demonstrated using nebulin-deficient mice (Witt et al., 2006). Titin is also
implicated in the regulation of sarcomeric structure, found to be important for Z-line
structure and formation and also in the organization of thick filaments at the A-band
(Clark et al., 2002;Agarkova et al., 2003). In addition to this, the elastic properties of
titin indicate that this protein is involved in adaptations to mechanical stress
(Tskhovrebova and Trinick, 2003).
The proteins that comprise the sarcomere are vital to the integrity of the muscle.
Mutations in these proteins cause a range of myopathies with pathological features evident in both skeletal and cardiac muscle (discussed further in section 1.5).
1.2.2 Cytoskeletal structures in muscle- costameres and the Z-LAC
In all cells, the cytoskeleton plays an important role in many aspects of cell function.
Skeletal muscle is no exception and with the extreme mechanical stresses placed on structures within skeletal muscle, the cytoskeleton plays a major role in maintaining the regular structure of the sarcomeric apparatus.
Page 25 Chapter One: Literature Review
Costameres
The sub-sarcolemmal cytoskeletal elements known as costameres were first described in 1983 (Pardo et al., 1983) as an ‘orthogonal lattice associated with the sarcolemma of skeletal muscle cells’. These structures lie beneath the sarcolemmal surface providing a
link between the sarcomeric apparatus and the ECM via the membrane (Figure 1.7)
(Rybakova et al., 2000), aligning the plasma membrane with the sarcomeres and protecting the sarcolemma from contraction induced damage (Bloch et al., 2002).
Costameres also function to transmit contractile forces from the sarcomeric apparatus to the ECM (Bloch et al., 2002).
Figure 1.7 The costamere links the myofibrils to the sarcolemma.
Costameres consist of a number of proteins including - actin and the DGC complex and form a link between the contractile apparatus and the muscle plasma membrane.
Adapted from (Ervasti, 2003)
Costameres consist of an extensive number of proteins that form a large network and
include structural proteins and focal adhesion and signalling molecules. Dystrophin is
found concentrated in costameric regions along with its associated proteins, known as
the dystrophin-glycoprotein complex (DGC), which connects to the basal lamina via
laminin (Dickson et al., 1992). The cytoskeletal actin isoform, -actin, is found in the
costamere. Here it is believed to bind dystrophin to link the membrane-bound DGC to
the sarcomeric apparatus (Ervasti and Campbell, 1993;Rybakova et al., 2000).
Dystrophin also links -actin to the membrane. The presence of -actin at the costameric region is abolished in the dystrophin-deficient muscular dystrophy mouse
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model, mdx (Rybakova et al., 2000). -actin is also colocalises with vinculin in this
region and is postulated to bind a number of other proteins such as spectrin, synemin,
plectin and α-actinin in both the Z-line and in regions close to the membrane
(Rybakova et al., 2000;Ervasti, 2003). Intermediate filaments also play a role at the
costamere with proteins such as desmin (Capetanaki and Milner, 1998;O'Neill et al.,
2002), synemin and paranemin (Bellin et al., 2001;Bloch et al., 2002) distributed at the
level of the Z-line and the linker molecule plectin found to tether the desmin
intermediate filaments to the dystrophin complexes (Hijikata et al., 2003).
Animal models lacking costameric protein components have demonstrated that
costameric structures play a significant role in the preservation of muscle structure
(Rybakova et al., 2000;O'Neill et al., 2002;Hanft et al., 2006). The removal of a
number of these proteins including dystrophin, desmin and α-dystrobrevin (a member
of the DGC) in skeletal muscle leads to a dystrophic phenotype with compromised
membrane structure (Ervasti, 2003). Loss of one protein was also found to have effects
on the presence of other interacting proteins; the loss of dystrophin in the costamere led
to a loss of the association of -actin with the membrane (Rybakova et al., 2000). This
demonstrates that ablation of one protein can alter the function of the costameric
network (Ervasti, 2003).
The Z-line associated cytoskeleton (Z-LAC)
The Z-line associated cytoskeleton (Z-LAC) was first described by Kee et al. (2004) as
‘a novel structural compartment in muscle’. It was demonstrated that cytoskeletal Tms
from the gene, such as Tm5NM-1, localise to compartments in skeletal muscle fibres such as the sarcolemma and the Z-LAC (Kee et al., 2004). These Tms were found to
Page 27 Chapter One: Literature Review co-localise with the cytoskeletal -actin at these sites, suggesting a role for this filament system in linking myofibrils and costameres (Figure 1.8). Transgenic mice over- expressing Tm3 (not normally expressed in wild type skeletal muscle) demonstrated a mild dystrophic phenotype with the exogenous Tm localising to, and possibly disrupting, Z-LAC structures. This indicated the importance of Tm-defined cytoskeletal structures external to the sarcomere in skeletal muscle, suggesting that alterations or mutations in these Tm isoforms may cause muscle disease
Figure 1.8 Tropomyosins define the Z-line associated cytoskeleton (ZLAC).
Cytoskeletal Tms and actins are found adjacent to the Z-line in skeletal muscle. An isoform from the -gene, Tm5NM1 (light blue), is specifically found in this region colocalising with -actin. In addition to this -actin is also found at the costameric region linking the myofibrils to the DGC at the membrane.
Adapted from (Kee et al., 2004).
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1.2.3 The sarcoplasmic reticulum (SR) and T-tubule system
Muscle contraction is initiated by electrical stimulation in the surface membrane and T-
tubules, followed by the release of calcium ions from the SR. The T-tubule system
consists of invaginations of the sarcolemma at the level of the Z-line while the SR
membranes are not continuous with the sarcolemma and extend longitudinally along the
sarcomere ending at the terminal cisternae at the Z-line (Franzini-Armstrong, 1994).
The junction between these two membrane systems is known as the triad and forms the
calcium release unit (CRU) (Flucher et al., 1993;Takekura et al., 2001;Paolini et al.,
2004). The CRUs consist of voltage-gated Ca2+ channels, the dihydropyradine receptor
(DHPR) of the T-tubules, and the Ca2+ release channels, the ryanodine receptors
(RyRs) of the SR (Paolini et al., 2004). The DHPRs and RyRs are structurally linked and regulate one another, with the DHPR functioning as the voltage sensor signalling to the RyR to release Ca2+ units (Paolini et al., 2004). The longitudinal elements of the SR then re-gather the free Ca2+ units which are returned to the terminal cisternae (Flucher et al., 1993).
The accurate formation of the T-tubules and SR systems as well as preservation of the structure of these elements is extremely important to muscle function (Takekura et al.,
2001). The T-tubule system is subjected to extreme mechanical forces during contraction and is supported by the cytoskeletal protein scaffold. It is suggested by
Kostin et al. (1998) that at least three independent systems exist to link the cytoskeleton to the membrane and consequently the extracellular matrix (ECM) at the T-tubule: the vinculin, talin, integrin complex, the DGC and the spectrin-based membrane skeleton
(Kostin et al., 1998). Studies indicate that this scaffold resembles the cytoskeletal scaffold of the costameres (discussed in 1.2.2) (Ervasti and Campbell, 1993;Kostin et
Page 29 Chapter One: Literature Review
al., 1998). Similarly, it is hypothesized that the precise location of the terminal SR at
the level of the Z-line involves the actin cytoskeleton (Porter et al., 2005). Studies have
demonstrated the protein small ankyrin 1 (sAnk1) is enriched in the SR and is
important in targeting proteins to this region and it is postulated that this molecule links
the SR membrane to the actin-based cytoskeleton (Porter et al., 2005).
1.2.4 Neuromuscular junction (NMJ) and Actin
Skeletal muscle cells are stimulated by motor neurons at a chemical synapse known as the neuromuscular junction (NMJ). The axon of the motor neuron, whose cell body lies in the spinal cord or brain stem, is branched with each ending attaching to a single muscle fibre to form a NMJ. The motor neuron terminal is specialised for neurotransmitter release, containing a number of synaptic vesicles containing acetylcholine (ACh) (Sanes and Lichtman, 1999). The muscle membrane at the NMJ is specialised with deep folds and indentations in the sarcolemma, a surface rich with
ACh receptors and clusters of subsarcolemmal nuclei (Figure 1.9) (Hall and Sanes,
1993). Between the neuron and the muscle fibre is a 50 nm wide synaptic cleft containing the basal lamina, which extends into the junctional folds (Hall and Sanes,
1993). A third cell type is involved in the structure of the NMJ, the Schwann cell. Three to five Schwann cells cap the axon terminals and are involved in modulating synaptic transmission and playing a role in nerve terminal growth and maintenance (Hughes et al., 2006).
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Sarcomeric apparatus
Synaptic vesicles
Pre-synaptic terminal
Post-synaptic terminal
Figure 1.9 The synapse between motor neuron and the muscle fibre is known as the neuromuscular junction (NMJ)
At the NMJ the sarcolemma is specialised, containing deep folds and creases to increase surface area. Synaptic vesicles contained within the neuron are filled with acetylcholine (Ach) molecules, which when released will bind to ACh receptors, found in abundance on the muscle cell membrane. Schematic adapted from (Martini et al., 2001) and EM image kindly provided by Emma Kettle
A number of cytoskeletal proteins are concentrated at the NMJ including vinculin, talin,
α-actinin, actin, filamin, desmin and Tm2 (Bloch and Hall, 1983;Sealock et al.,
1986;Marazzi et al., 1989;Askanas et al., 1990). Cytoskeletal-associating proteins such
as dystrophin and utrophin are also found concentrated in this region (Nguyen et al.,
1991;Hall and Sanes, 1993). It has also been proposed that the actin cytoskeleton is
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involved in the assembly of the NMJ during development and filamentous actin
anchors signalling molecules to the NMJ region (Dai et al., 2000).
1.3 Significant Functions of Skeletal Muscle
The role of skeletal muscle in the body is to produce force and provide motion for
mobility, posture and breathing. Skeletal muscle is also largely responsible for the
uptake and metabolism of glucose.
1.3.1 Muscle Contraction
The sliding filament theory was developed in the 1960s and describes the mechanism of
skeletal muscle contraction as the movement of the thick and thin filaments across one
another (Craig, 1994). Briefly, the cycle of contraction begins with the release of
neurotransmitter which binds the acetylcholine receptors (AChR) at the neuromuscular
junction, which changes the membrane potential of the sarcolemma. This signal
transmits down the T-tubules causing the release of calcium ions from the adjacent
sarcoplasmic reticulum. The calcium ions (Ca2+) bind to the troponin complex causing a change in conformation of this complex and the orientation of Tm filament on the thin filament exposing active sites on the actin filament. Myosin heads bind to active sites and in doing so hydrolyse ATP, causing a shift in the position of myosin on the thin filament towards the plus end of actin. This process is extremely rapid with the contraction of muscle occurring within approximately one tenth of a second (Alberts et al., 1998;Martini et al., 2001).
The position of Tm and troponin subunits, T, C and I (TnT, TnC, TnI), on the actin filament regulates the binding of myosin (Figure 1.10). The thin filament operates in
Page 32 Chapter One: Literature Review three states: the ‘blocked’ state in the absence of Ca2+, where the myosin heads are unable to bind actin due to the site of Tm, the ‘closed’ state, where myosin weakly binds actin and the ‘open’ state where myosin actively binds the thin filament
(McKillop and Geeves, 1993;Gordon et al., 2000). The binding of Ca2+ to TnC causes the Tm filament to move more closely to the groove of the actin filament to cause the open state, allowing the binding of myosin to actin and the generation of force (Van
Dijk et al., 2002).
Figure 1.10 Tropomyosin and the troponin complex regulates myosin binding to the thin filament
A. In the absence of calcium, TnC is in a ‘shut’ position and the TnI binds actin, blocking the myosin binding site B. In the presence of calcium, TnC binds two calcium ions causing TnC to bind TnI, causing conformational changes in the actin/Tm filament, allowing the myosin to bind.
Adapted from (Smith and Geeves, 2003).
1.3.2 The transport of glucose in skeletal muscle
Glucose is an essential energy source for all eukaryotic cells and the uptake of glucose into tissue is critical for the maintenance of glucose homeostasis. Skeletal muscle plays a major role in the uptake of glucose. This process is initiated by the release of insulin by the pancreas in response to increased levels of glucose in the blood stream. Insulin binds to receptors on the cell membrane leading to the translocation of the membrane- bound molecule GLUT4. GLUT4, which facilitates the movement of glucose through the membrane, is stored primarily in intracellular storage vesicles and continually cycles between these sites and the cell surface (Foster et al., 2001). The pathways by
Page 33 Chapter One: Literature Review
which insulin-stimulated glucose uptake takes place are complicated and as yet, not
completely understood. The phosphatidylinositol 3-kinase (PI-3K) pathway has been implicated in the translocation of GLUT4 vesicles to the plasma membrane. This pathway begins with the binding of insulin to its receptor on the external surface of the cell membrane, which alters the receptor conformation causing a signalling cascade leading to the movement of vesicles toward the membrane [reviewed in (Bryant et al.,
2002;Chang et al., 2004)]. Once vesicles reach their destination they must dock with
the membrane (Figure 1.11). SNARE (soluble N-ethylmaleimide-sensitive factor
attachment protein receptor) proteins on both the vesicle and plasma membrane
facilitate the docking of GLUT4-containing vesicles. The vesicles contain the SNARE
protein VAMP2, which interacts with its binding partner syntaxin-4 which is situated in
the plasma membrane and, aided by accessory proteins, fuses GLUT4 vesicles to the
plasma membrane (Olson et al., 1997). GLUT4 translocation from intracellular storage
sites can occur in an insulin-independent manner. Muscle contraction can trigger
GLUT4 trafficking as well as through platelet-derived growth factor (PDGF) activation
of the PI-3K pathway.
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Figure 1.11 GLUT4 molecules cycle between the plasma membrane and intracellular storage vesicles.
Upon stimulation by insulin, a signalling cascade initiates the mobilisation of GLUT4 containing vesicles from an intracellular storage site to the plasma membrane. Here these vesicles, which contain a VAMP2 molecule on their surface, dock with the plasma membrane by association with the proteins syntaxin 4 and SNAP 23. After membrane fusion the GLUT4 protein is now found on the cell surface. Following the process of glucose uptake GLUT4 is then internalised.
Adapted from (Dugani and Klip, 2005)
Studies of GLUT4 in skeletal muscle identified internal storage compartments
containing GLUT4 vesicles (Rodnick et al., 1992;Ploug et al., 1998). It was found that
at basal state the GLUT4 localised to internal structures and was excluded from
membrane regions. In contrast to adipose tissue cells where the levels of GLUT4 at the
membrane of unstimulated cells is significantly higher (Rodnick et al., 1992). After stimulation, GLUT4 was found at both the plasma membrane and at the T-tubules
(Ploug et al., 1998). Recently, Lauritzen et al. (2006) identified the T-tubules to be of
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great importance in insulin signalling. Using imaging techniques of skeletal muscles in
live mice Lauritzen and colleagues concluded that the T-tubules play a major role in
extending insulin stimulation deep within the muscle fibre which is densely packed
with myofibrillar proteins. It is also suggested that defects of the T-tubule system may
lead to insulin resistance in muscle indicating a link to type II diabetes (Lauritzen et al.,
2006).
Cytoskeletal filaments are proposed to be involved in the insulin signalling pathway, the transport of GLUT4 vesicles from the cytosol to the plasma membrane and in the docking of vesicles to the membrane. Microtubules are implicated in anchoring
GLUT4-containing vesicles to their storage pool, with disruption of microtubules in adipocytes resulting in dispersion of the GLUT4 throughout the cell (Fletcher et al.,
2000;Liu et al., 2003). Huang et al. (2005) showed that disruption of microtubules in adipocytes rendered them unable to respond to addition of insulin. Instead the cells were forced to use an insulin-independent pathway, using platelet-derived growth factor
(PDGF) to stimulate translocation of GLUT4 vesicles (Huang et al., 2005). However, studies in skeletal muscle have shown that microtubules are not required for insulin- stimulated or contraction-induced glucose transportation, but may play a role in the positioning of GLUT4 within the cell or in the return of GLUT4 from the plasma membrane to the perinuclear region (Ai et al., 2003).
The role of actin in glucose transport has been well documented [reviewed in
(Tsakiridis et al., 1999;Kanzaki, 2006a)]. Studies in both adipocytes and myoblast cultures has shown that the remodelling of actin is required for the insulin-stimulated translocation of GLUT4 (Wang et al., 1998;Tong et al., 2001;Torok et al., 2004). The
Page 36 Chapter One: Literature Review insulin-dependent translocation of GLUT4 vesicles is disrupted by actin depolymerisation drugs cytochalasin D and latrunculin (Tsakiridis et al., 1994;Wang et al., 1998). It has been suggested that there is a functional link between actin and the microtubule cytoskeleton in insulin-mediated transport of GLUT4 (Olson et al., 2003).
The disruption of the actin cytoskeleton by latrunculin B prevented the insulin- dependent changes of microtubules in cultured adipocytes, however a molecular link is yet to be identified (Olson et al., 2003). It also has been found that disruption of the microtubules resulted in alterations in the signalling cascade, but that GLUT4 vesicles were still able to translocate to the cell membrane (Huang et al., 2005).
1.4 Muscle Fibre Formation
In mammals, the formation of muscle fibres occurs in utero, with muscles developing and maturing postnatally. The post-mitotic, multinucleated muscle cells form from mononucleate myoblast cells, which fuse and grow to form the fibres found in mature skeletal muscle. Regeneration of muscle fibres following injury or disease mimics muscle development. This is achieved by a pool of quiescent, mononucleate cells residing beneath the basal lamina of the muscle fibres, known as satellite cells. These cells can replicate and fuse into existing muscle fibres to repair damage and also regenerate entire muscle fibres. These processes are detailed here.
1.4.1 Muscle development
The first muscle fibres develop in the embryo from somite-derived cells at approximately ED 11-14 in the mouse limb and prior to 12 weeks gestation in the human embryo (Yang and Makita, 1996;Buckingham et al., 2003). Myoblasts, mononucleated muscle precursor cells, fuse to form myotubes, immature muscle fibres.
Page 37 Chapter One: Literature Review
There are two temporal waves of myotube formation resulting in the formation of primary and secondary myotubes (Franzini-Armstrong and Fischman, 1994;Schnorrer and Dickson, 2004). Primary myotubes develop first and groups of these are surrounded by a basement membrane-like structure. The formation of secondary myotubes follow approximately two to three days later. They form adjacent to primary myotubes under the basal lamina (Franzini-Armstrong and Fischman, 1994).
The development of the SR and T-tubules takes place over several weeks both in utero
and post-birth (Flucher et al., 1992). This process can be broken into a series of three
discreet steps: the separate differentiation of the membrane compartments of the T-
tubules and SR, the formation of the triad/CRU and the association of the membrane
compartments with the myofibrils (Flucher et al., 1994). The endoplasmic reticulum
(ER) of the myotube develops initially. The membranous system of the ER differentiates into the SR by the expression of SR-specific proteins such as the SERCAs and calsequestrin (Flucher et al., 1993). These SR-specific proteins are present from approximately ED 17 in the mouse and are visible as discreet bands near the Z-line
(Takekura et al., 2001). The T-tubules develop from small invaginations of the plasma membrane and are detected at approximately ED 15. These invaginations continue to develop penetrating deeper into the cell where they form the T-tubules and are found to run in a longitudinal orientation (Takekura et al., 2001). The transition of the T-tubules to a transverse orientation does not occur until after birth (Franzini-Armstrong, 1991).
T-tubules and SR membranes form junctions at ED 16 and these CRUs are also initially located in a longitudinal orientation (Franzini-Armstrong, 1991;Franzini-Armstrong,
1994). The final stage of development is the organisation of the CRUs and T-tubules into a transverse orientation relative to the myofibril (Franzini-Armstrong, 1994). At
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this stage there is an increase in longitudinal SR and an increase in pump and channel
proteins in the membrane (Franzini-Armstrong, 1994). Here the development of the T-
tubules and SR is coordinated and final maturation of the longitudinal SR and triad
positioning is not achieved until three weeks of age (Franzini-Armstrong,
1991;Franzini-Armstrong, 1994;Takekura et al., 2001). The T-tubules and SR are
considered fully developed when they are positioned at the border between the A and I
bands (Flucher et al., 1993).
The development of the contractile apparatus occurs throughout the process of myotube
development and is known as myofibrillogenesis (Figure 1.12) (Sanger et al., 2006).
Myofibrillogenesis describes the process of the organisation of the muscle
cytoarchitecture, the details of which are not well understood. The mechanisms of
sarcomere formation have been studied and described primarily by Sanger et al. (Rhee et al., 1994;Sanger et al., 2002;Wang et al., 2005;Sanger et al., 2006). Sanger et al. propose a three step model: premyofibrils to nascent myofibrils to mature myofibrils.
Premyofibrils represent mini-sarcomeres with smaller, closely-spaced Z-bodies
(immature Z-lines) and non-muscle isoforms of sarcomeric proteins. At this stage, non- muscle myosin II interdigitates with actin filaments which contain muscle isoforms of tropomyosin and troponin. The premyofibrils begin to align to form nascent myofibrils in which muscle myosins replace non-muscle myosins and titin is included. Z-bodies then align to form Z-lines, and the thick filaments also align and the mature myofibril is formed (Sanger et al., 2006).
Page 39 Chapter One: Literature Review
Figure 1.12 Myofibrils are built in three steps
Premyofibrils develop initially in the myotubes, containing non-muscle myosin isoforms and small dense bodies containing α-actinin. These develop into nascent myofibrils with the replacement of non-muscle myosins with muscle specific isoforms. Z-bodies also condense and begin to align and titin is found bound to α-actinin. These nascent myofibrils then align completely and the Z-lines develop to create the mature myofibril.
Adapted from (Sanger et al., 2006)
It is proposed that cytoskeletal -actin plays a role in myofibrillogenesis (Lloyd et al.,
2004). However, it has been demonstrated in a gene knockout mouse that expresses no
-actin in skeletal muscle, that -actin is not necessary for muscle development
(Sonnemann et al., 2006). Early in development, myoblasts contain stress fibre-like
structures which are composed of non-muscle actins ( and ) and associated proteins
including myosin and non-muscle α-actinin (Dlugosz et al., 1984). Following
differentiation into myotubes, -actin re-localises to the cell membrane where it binds
signalling protein N-RAP and sequesters α-actinin to form Z-bodies (Lloyd et al.,
2004). It is also proposed that -actin facilitates the organisation of the Z-bodies into
Z-lines (Lloyd et al., 2004). In mature muscle, -actin is found at the costameres and
the Z-LAC (Rybakova et al., 2000;Kee et al., 2004).
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1.4.2 The regeneration of muscle fibres
The process of muscle regeneration occurs following muscle injury or during disease and during focal repair or stretch (Bischoff, 1994). In healthy human muscle, local injury occurs occasionally, but in certain diseases, such as the dystrophies, muscle fibres are more susceptible to damage, particularly to the plasma membrane (Carlson and Faulkner, 1983). Although the muscle fibre itself is considered post-mitotic, the regenerative capacity of muscle is ensured due to a population of cells known as the satellite cells that lie beneath the basal lamina of each myofibre (Bischoff, 1994;Charge and Rudnicki, 2004;Ehrhardt and Morgan, 2005). The process of muscle regeneration typically occurs following trauma that damages the sarcolemma (Figure 1.13). The muscle then undergoes necrosis, with macrophages infiltrating the basal lamina to remove debris. Satellite cells then proliferate, exit the cell cycle and fuse to form multinucleated myotubes (Figure 1.13) (Charge and Rudnicki, 2004). These myotubes then fuse with the remaining myofibre to form a regenerated muscle cell with centralised nuclei, which is a hallmark of a regenerated fibre (Charge and Rudnicki,
2004;Jarvinen et al., 2005;Ehrhardt and Morgan, 2005). Satellite cells also respond to mild injury, such as stretch induced damage, in a process known as focal regeneration
(Bischoff, 1994;Jarvinen et al., 2005). Here, the satellite cells in close proximity to the injury are activated and the damage is repaired. Unlike the complete regeneration of the fibre, centralised nuclei may not be detected due to the isolation of the injury to a small region of the fibre (Aarimaa et al., 2004).
Page 41 Chapter One: Literature Review
Figure 1.13 Satellite cells drive muscle regeneration.
Muscle damage causes satellite cell activation and proliferation (B). These cells invade the muscle fibres and align to form centralised nuclei (C) and begin to transcribe proteins to repair the sarcomeric apparatus and other muscle fibre structures (D). The nuclei will then remain centralised, which is a hallmark of a regenerated muscle fibre.
Adapted from (Bischoff, 1994)
Muscle regeneration, both mild and severe, can be modelled in animal muscle. Severe
muscle degeneration and regeneration can be induced by the injection of a myotoxic
agent directly into the muscle. Notexin, a myotoxin agent derived from tiger snake
venom, elicits a specific response in muscle causing the fibres to degenerate completely
leaving only the basement membrane after three days (Plant et al., 2006). Following this the regenerative processes of muscle can be studied as the muscle regenerates from mononucleated cells to a relatively mature muscle at approximately 10-20 days post- injection (Plant et al., 2006). Processes such as immobilisation of the limbs and
crushing or laceration of muscle also cause varying degrees of muscle damage,
resulting in repair (Booth, 1977;Carlson and Faulkner, 1983;Goldspink, 1999;Hill et
al., 2003). These techniques are found to mimic human injury and allow the study of
the processes involved in repair.
1.5 Muscle Disease
Muscle diseases, or myopathies, can be extremely debilitating and arise from defects in
the muscle fibre. The prognosis for patients with myopathic disorders varies from mild,
Page 42 Chapter One: Literature Review
with normal lifespan and little to no disability, to severe, affecting the muscles required
for breathing and food intake, often leading to premature death. Two groups of diseases are described here: muscular dystrophies and congenital myopathies with affected filaments.
1.5.1 Muscular Dystrophies
Muscular dystrophies are a group of genetically diverse muscle diseases characterised
by the degeneration and regeneration of muscle fibres causing progressive muscle
weakness (Weir, 2000). Dystrophic muscle is characterised pathologically by a
variation in fibre size, fibrosis, necrosis and centrally located nuclei (Figure 1.14)
(Dalkilic and Kunkel, 2003). The inheritance of these disorders is varied and can be X-
linked recessive (ie. Duchenne or Becker muscular dystrophy), autosomal dominant
(Emery Dreiffus muscular dystrophy) or autosomal recessive (limb-girdle muscular
dystrophy type 2) (Durbeej and Campbell, 2002).
*
*
Figure 1.14 Dystrophic muscle contains regenerating fibres.
Biopsy samples from control (A) and dystrophic (B) muscle stained with
haemotoxylin and eosin (H&E) show the typical features of dystrophic muscle. Nuclei (stained purple) are situated at the periphery of fibres in the control muscle,
however, dystrophic muscle has centrally located nuclei (arrows B). Dystrophic muscle also contains areas of necrosis (asterixis, B) and small regenerating fibres
(arrowheads, B).
Adapted from (Davies and Nowak, 2006) Page 43 Chapter One: Literature Review
Dystrophies arise from deficiencies and defects in a range of proteins including proteins
located in the ECM, transmembrane and membrane-associated proteins, nuclear
membrane proteins and cytoplasmic proteins associated with the sarcomere (Rando,
2001). The most common form of muscular dystrophy is the X-linked recessive
Duchenne muscular dystrophy (DMD) caused by mutations in the gene encoding
dystrophin. Mutations in this gene also cause the milder Becker muscular dystrophy
(BMD). Cells deficient in dystrophin are particularly prone to membrane damage as
this protein links the membrane to the contractile apparatus (Ervasti and Campbell,
1993). An increase in serum creatine kinase levels is indicative of sarcolemmal damage
and leaky membranes (Engel et al., 1994). This enzyme, normally contained within muscle fibres, is released into the blood stream through damaged muscle membranes and can be measured in dystrophic patients by a simple blood test (Engel et al., 1994).
Muscles with damaged membranes degenerate and are forced to regenerate. Due to repeated rounds of degeneration and regeneration, the proliferative capacity of satellite cells is diminished and the muscle is unable to continue regeneration. Following this muscles are invaded by fibrosis and connective tissue (Blau et al., 1983). Patients suffering from DMD experience progressive muscle wasting and weakness, generally resulting in death by the third decade of life (Engel et al., 1994). Mutations have also been found in genes encoding other proteins from the DGC, as well as proteins from the nuclear membrane, the sarcomere and proteins that reside in the sarcoplasm (Table
1.5.1).
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Table 1.1 Examples of genes causing various muscular dystrophies
Disease Gene Product DMD Dystrophin BMD Dystrophin Emery-Dreifuss muscular dystrophy Emerin Lamin A/C Limb-girdle muscular dystrophy Titin Myotilin Lamin A/C Caveolin-3 Calpain-3 Dysferlin Sarcoglycans Congenital muscular dystrophy Fukutin-related protein Fukutin Integrin α7 Collagen type VI
(Spence et al., 2002;Dalkilic and Kunkel, 2003)
A number of mouse models are available to study muscular dystrophy pathogenesis and
treatment options (Durbeej and Campbell, 2002). The most commonly used mouse
model is the mdx mouse that arose from a spontaneous mutation in a C57BL/10 mouse.
The disease phenotype arises from a mutation in exon 23 of the dystrophin gene
causing a premature stop codon and hence a dystrophin deficiency (Sicinski et al.,
1989). As with the human disease, the serum creatine kinase levels are increased in the mouse, but unlike DMD, clinical muscle weakness is not observed and the lifespan of mdx mice is not grossly reduced (Blake et al., 2002). However, pathologically the muscle appears dystrophic with evidence of degeneration/regeneration including necrotic and centrally nucleated fibres. A crisis period, consisting of a significant degeneration event, occurs at approximately 3-4 weeks of age. A decline in the degeneration/regeneration process is seen after this time (DiMario et al., 1991;Grady et al., 1997b). A more dramatic phenotype is seen in mice that are null for both dystrophin
Page 45 Chapter One: Literature Review
and utrophin, a protein structurally similar to dystrophin found at the NMJ (Grady et al., 1997a). Mice developed weight loss, decreased muscle strength, continuous
degeneration/regeneration and spinal deformities, with these symptoms leading to
premature death most likely by respiratory failure (Grady et al., 1997b). This model is
said to more closely resemble human DMD (Grady et al., 1997b). Studies of limb-
girdle muscular dystrophy have been aided by mice null for α-, - or δ-sarcoglycans and
the naturally occurring dysferlin deficient mouse, the SJL mouse (Weller et al.,
1997;Vafiadaki et al., 2001;Watchko et al., 2002). These mice exhibit varied
pathologies and differences in age of onset; a phenomena also observed in human
patients affected by limb-girdle muscular dystrophies (Watchko et al., 2002).
Currently, there is no cure for muscular dystrophy. Until a cure is developed, the
majority of therapeutic approaches focus on supportive care for patients while closely
monitoring cardiac and pulmonary status (Strober, 2006). Recent research has been
focused on four main strategies: viral-mediated gene therapy, non-viral gene therapy,
donor cell transplantation and autologous cell transplantation (Judge and Chamberlain,
2005;Quenneville and Tremblay, 2006;Rando, 2007;Price et al., 2007).
DMD is a good candidate for treatment by gene therapy as it is a monogenic disease,
caused only by mutations to the dystrophin gene (Foster et al., 2006). Gene therapy for
DMD has been hampered due to the limited capacity of viral vectors carry the large
dystrophin gene (2.3 Mbp). To overcome this, a functional miniaturised form of
dystrophin was developed, leading to the expression of a smaller but functional protein.
Research into viral-mediated gene therapy has explored the use of adeno-associated
viral vectors to deliver the mini-dystrophin to muscle cells (Gregorevic et al.,
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2006;Wang et al., 2007). The introduction of virus into the muscle cell incorporates the mini-dystrophin into the genome to express dystrophin, correcting the disease phenotype. Viral delivery of antisense oligonucleotides against exon splice junctions that skip the defective exon in the dystrophin gene has also been investigated as a treatment of DMD (Denti et al., 2006). Using this method, antisense oligos are
packaged in adeno-associated viral vectors and delivered systemically via the tail vein.
This approach has shown success in the mdx mouse model, rescuing dystrophin
expression in a range of muscles and showing significant functional recovery (Denti et
al., 2006). Caution must be exhibited with viral incorporation into the genome of
patients. Potential adverse effects include the induction of immune response and the
gene incorporation into sites containing oncogenes leading to lymphoma and leukaemia
(Liu and Muruve, 2003;Porteus et al., 2006).
Transplantation of donor muscle precursor cells has been investigated as a potential
therapy. In this approach, healthy myoblast cells are obtained from a donor, expanded
in cell culture and transplanted into patient muscles where they fuse with muscle fibres
(Skuk et al., 2004). However, challenges also exist with this type of therapy. Problems
include poor donor cell survival, inability of donor cells to migrate from the point of
introduction and immune rejection of and inflammatory response to donor cells (Urish
et al., 2005). The delivery of autologous cells, those harvested from the patients, is a
more attractive option. Using an autologous cell transplantation approach, harvested
cells from the patient are transduced with viral vectors, such as lentivirus, prior to
transplantation back into the patient, avoiding donor rejection (Li et al.,
2005;Quenneville et al., 2007). The search for an ideal cell population for
transplantation is ongoing, with researchers looking to identify cells that are able to
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form muscle fibres, are easily transducible and can be delivered systemically
(Dellavalle et al., 2007). Muscle precursor cells derived from satellite cell populations have shown been shown to be difficult to isolate in sufficient numbers. Problems have arisen with the potency of muscle precursor cells which was found to decrease upon expansion of cells in cell culture to required amounts for injection (Montarras et al.,
2005). Cells isolated from blood vessels and bone marrow have been shown to hold promise for cell based therapies as they can be delivered systemically and are able to cross the blood vessel barrier into the muscle tissue (Dezawa et al., 2005;Sampaolesi et al., 2006;Dellavalle et al., 2007).
1.5.2 Congenital Myopathies with Affected Filaments
Congenital myopathies include a subset of muscle diseases such as nemaline myopathy
(NM), actin myopathy and intranuclear rod myopathy (Clarkson et al., 2004). This group of muscles diseases is clinically heterogenous and is characterised by progressive skeletal muscle weakness. These diseases are caused by mutations in the thin filament proteins.
Nemaline Myopathy (NM)
Nemaline patients are clinically characterised by muscle weakness and hypotonia.
Muscle weakness is either static or slowly progressive and a type I (slow) fibre
predominance is usually observed. Pathologically, NM skeletal muscle is characterised
by the presence of nemaline bodies or ‘rods’. These rods are detectable at the light
microscopy (LM) level as dense accumulations of proteins and by electron microscopy
(EM) are found to originate from the Z-line and contain thin filament proteins (Figure
1.15) (Corbett et al., 2001;Ilkovski et al., 2001). NM has been described as a disease of
Page 48 Chapter One: Literature Review
the sarcomere. Mutations that cause NM have been identified in five genes encoding
thin filament proteins: nebulin, α-Tmslow, -Tm, troponin-T and skeletal α-actin (Laing
and Nowak, 2005). Nemaline rods have been found to contain a number of thin
filament proteins including α-actinin, filamentous actin, myotilin and filamin
(Wallgren-Pettersson et al., 1995;Ilkovski et al., 2001;Schroder et al., 2003).
Figure 1.15 Muscle samples from nemaline patients contain filamentous accumulations.
Light (A) and electron microscopy (B) show protein accumulations, known as rods, and disruptions of sarcomeric structure (arrows) of patients containing mutations in both skeletal actin (A) and α-Tmslow (B). Gomori-trichrome staining of patient muscle biopsies in cross section allows visualisation of filament accumulations at the level of the light microscope for diagnostic purposes while electron microscopy allows more detailed analysis of electron-dense nemaline rods.
Adapted from (Corbett et al., 2001;Ilkovski et al., 2001)
We have generated a mouse model for NM with a mutation in the gene encoding α-
Tmslow at position nine where a methionine is replaced by an arginine designated
TPM3(Met9Arg) (Corbett et al., 2001). Mutations in the α-Tmslow gene are a rare cause
of NM in humans. The Met9Arg mutation was first discovered as an autosomal
dominant mutation in an Australian family with a late onset, slowly progressive form of
NM (Laing et al., 1995). Since this initial observation, other mutations have been
Page 49 Chapter One: Literature Review
observed in the α-Tmslow gene. These patients exhibit differences in pathology, with disease phenotypes varying in severity and age of onset (Wattanasirichaigoon et al.,
2002). The mouse model of the TPM3 (Met9Arg) mutation was found to phenocopy the human condition with the same mutation. The mice exhibited late-onset muscle weakness, detection of rods at varying extents in all muscles and increased type I fibres
(Corbett et al., 2001). This mouse model was found to be extremely useful in examining the affect of exercise on NM muscles (Joya et al., 2004;Nair-Shalliker et al.,
2004). The TPM3(Met9Arg) mouse model was used to determine that, unlike in muscular dystrophies, exercise does not aggravate NM-affected muscle (Nair-Shalliker et al., 2004) and can resolve the severe muscle weakness that is elicited in patients with this disease who experience prolonged periods of immobilisation (Joya et al., 2004).
Unlike the muscular dystrophies, membrane damage and degeneration are not historically recognised features of NM. However, recent microarray analysis of both human (Sanoudou et al., 2003) and TPM3(Met9Arg) mouse (Sanoudou et al., 2006) muscle identified the presence of markers for activated satellite cells and immature fibres indicating the presence of a regenerative process. Examination of NM ultrastructure also revealed evidence for a repair process in NM mouse muscle with affected fibres containing centralised nuclei, but no evidence of degeneration or necrosis (Sanoudou et al., 2006).
Actin Myopathy (AM)
Actin myopathy (AM) arises from mutations in the ACTA1 gene, which encodes α- skeletal actin. Patients present as floppy babies, with distressed respiration requiring ventilation often progressing to death within the first year of life. This disease is
Page 50 Chapter One: Literature Review characterised pathologically by excess thin filament deposits in skeletal muscle. Patient muscle biopsies contain actin aggregates, which are sometimes found to fill entire transversely-sectioned muscle fibre (Sparrow et al., 2003;Clarkson et al., 2004;Goebel and Muller, 2006).
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1.6 Research Objectives
Previous research in the Muscle Development Unit of the Children’s Medical Research
Institute had identified a novel filament structure defined by cytoskeletal Tm,
Tm5NM1, in skeletal muscle. This novel structure was named the Z-LAC (Kee et al.,
2004). The disruption of this structure leads to a disease phenotype in skeletal muscle, however, the specific role of cytoskeletal Tms in the Z-LAC and any other region of skeletal muscle has not yet been elucidated.
The aim of this project is to identify Tm-defined filamentous populations in skeletal muscle and study the role/s of these in normal and diseased skeletal muscle.
In order to do this, a number of specific analyses were undertaken:
• Western blotting techniques with exon-specific antibodies (Schevzov et al.,
2005b) were used to identify the cytoskeletal Tm isoforms present in muscle.
• Immunohistochemical analysis of muscle sections and isolated muscle fibres
were carried out to identify the localisation of Tm isoforms and their associated
organelles or membrane systems.
• Tm4 abundance and localisation was analysed using a combination of
techniques in muscle undergoing regeneration and repair, such as muscles
exhibiting disease phenotypes, to elucidate the role of Tm-defined filaments in
muscle disease.
• Transgenic (Bryce et al., 2003) and knockout mice (Vrhovski et al., in
preparation) were generated to study the roles of Tm5NM1. The skeletal
Page 52 Chapter One: Literature Review
muscle of these mice were analysed to give insights into regulation of
cytoskeletal Tms in muscle.
• Microarray analysis was performed on Tm5NM1 transgenic and knockout
mouse skeletal muscle to identify potential physiological roles of Tm5NM1.
• The potential role of Tm5NM1 in glucose metabolism was explored by
conducting glucose uptake and tolerance testing on transgenic and knockout
mice and analysis of adipose tissue.
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Section Two: The Roles of Cytoskeletal Tropomyosins in Muscle.
Chapter Two: Cytoskeletal tropomyosins form functionally distinct filaments in skeletal muscle
Chapter Three: Tropomyosin 4 is an indicator of repair/remodelling in skeletal muscle disease
Chapter Four: The altered expression of tropomyosin 5NM1 in skeletal muscle affects membrane morphology and metabolic pathways
Chapter Five: Tropomyosin 5NM1 is involved in glucose transport and adipose tissue proliferation
Chapter Two Cytoskeletal tropomyosins form functionally distinct filaments in skeletal muscle
2.1 Introduction
The specific Tm isoform bound to actin alters the dynamics of the filament, modulating the actin-binding proteins that associate with the filament. Tm isoforms have been shown to protect from the severing actions of gelsolin (Ishikawa et al., 1989a) and
ADF/cofilin (Ono and Ono, 2002;Bryce et al., 2003) and also affects binding of myosin motor proteins (Bryce et al., 2003). Sorting of functionally distinct Tm isoforms provides a mechanism for the spatial regulation of different actin filament populations
(Gunning et al., 2005;Gunning et al., 2007). This phenomena is best described in neuronal cells undergoing development, where Tms sort to specific compartments in the immature neuron with localisation altered following differentiation (Gunning et al.,
1998a). Compartmentalisation of Tms has also been observed in fibroblasts and epithelial cells (Percival et al., 2000;Dalby-Payne et al., 2003). In skeletal muscle, the sarcomeric Tms, αTmfast, Tm and αTmslow, sort to the thin filament in the sarcomere and facilitate the interactions between actin and myosin to drive muscle contraction
(Craig, 1994). The specific arrangement of the actin/Tm filaments is essential for proper contractile function.
Like all cells, the skeletal muscle fibre contains a cytoskeleton. The most well-defined cytoskeletal structure in muscle is the costamere. Costameric structures consist of an arrangement of proteins that connect the myofibrils to the sarcolemma playing a role in
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Chapter Two: Cytoskeletal Tms in skeletal muscle the maintenance of sarcomeric structure and membrane stability (Rybakova et al.,
2000). Cytoskeletal -actin, attached to the Z-line, connects to the dystrophin glycoprotein complex (DGC) which is embedded in the membrane and links to the
ECM (Bloch et al., 2002). The disruption of the costameres can result in muscle disease. A well-known example of a costameric disruption is Duchenne’s muscular dystrophy (DMD), a debilitating muscle disease caused by a mutation in dystrophin.
DMD patients experience muscle degeneration in response to disrupted sarcolemmal membranes, highlighting the importance of the costameres in maintaining membrane stability (Bloch et al., 2002;Ervasti, 2003).
Our laboratory, the Muscle Development Unit at the CMRI, described a population of cytoskeletal filaments in skeletal muscle defined by cytoskeletal Tm isoforms. This cytoskeleton is named in relation to their location; the Z-line associated cytoskeleton
(Z-LAC) (Kee et al., 2004). Transgenic mice over-expressing an exogenous Tm, Tm3, in skeletal muscle showed that Tm3 targeted to the Z-LAC region in skeletal muscle
(Kee et al., 2004). The incorporation of this inappropriate Tm caused disruption of the
Z-LAC which lead to a late onset dystrophic phenotype (Kee et al., 2004). This finding led to the hypothesis that other cytoskeletal Tms may be present in skeletal muscle and that Tms may play a role in the diversification of the actin cytoskeleton in muscle.
Here we show that Tms generated from all four Tm genes are present in skeletal muscle and define functionally distinct filament populations in relation to specific organelles.
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Chapter Two: Cytoskeletal Tms in skeletal muscle
2.2 Materials and Methods
2.2.1 Specific Materials
Specific materials included normal goat serum (Jackson, West Grove, PA, USA), PBS
(Gibco, Invitrogen, Carlsbad, CA, USA).
2.2.2 Animal strains
FVB/N mice 2 to 4-months of age were used for all animal experiments. All studies using mice were performed in accordance with the ethical guidelines of the Animal
Welfare Committee, National Health and Medical Research Council, Australia with
approval from the CMRI Animal Car and Ethics Committee.
2.2.3 Primary Antibodies
The following primary antibodies were used in this study:
Name Protein Recognised Antibody Type Source δ9d Tm4 Rabbit Polyclonal (Schevzov et al., 2005b) 9d Tm5NM1, Tm5NM2 Sheep Polyclonal (Schevzov et al., 2005b) αfast9d Exon 9d containing Sheep Polyclonal (Schevzov et al., 2005b) products from α and Tm genes -actin -actin Sheep Polyclonal (Schevzov et al., 2005b) 311 Tms containing exon Mouse Monoclonal Sigma (St Louis, MO, USA) 1a from all genes CH1 Sarcomeric Tm Mouse Monoclonal Sigma NCL-Dys Dystrophin Mouse Hybridoma Novocastra (Norwell, MA, USA) EA-53 α-actinin Mouse Monoclonal Sigma 5B2 α-actinin Rabbit Polyclonal (North and Beggs, 1996) MAB47 Dihydropyridine Mouse Monoclonal Chemicon International receptor (T-tubules) (Temecula, CA, USA) Calsequestrin Calsequestrin Mouse Monoclonal BD Biosciences (Franklin clone 51 Lakes, NJ, USA)
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Chapter Two: Cytoskeletal Tms in skeletal muscle
2.2.4 Secondary antibodies
The following secondary antibodies were used in this study:
Name Description Company Goat anti-mouse HRP- For western blot detection of BioRad (Hercules, conjugated mouse derived antibodies CA, USA) Goat anti-rabbit HRP- For western blot detection of BioRad conjugated rabbit derived antibodies Donkey anti-sheep For western blot detection of BioRad HRP-conjugated sheep derived antibodies Goat anti-mouse 594 Alexa-fluor 594 red Molecular Probes, fluorescent protein detection Invitrogen (Mt of mouse antibodies Waverly, VIC, Australia) Goat anti-rabbit 488 Alexa-fluor 594 green Alexa-Fluor, fluorescent protein detection Invitrogen of rabbit antibodies Donkey anti-sheep 488 Alexa-fluor 488 green Alexa-Fluor, fluorescent protein detection Invitrogen of sheep antibodies Donkey anti-sheep 594 Alexa-fluor 594 red Alexa-Fluor, fluorescent protein detection Invitrogen of sheep antibodies Goat anti-mouse 20nm For immuno-gold labelling British BioCell gold-conjugated of mouse derived antibodies International, (Cardiff, UK) F(ab)2 Goat anti-rabbit For immuno-gold labelling British BioCell 5nm gold-conjugated of rabbit derived antibodies International
2.2.5 Preparation of tissue samples for western analysis of protein
Muscle samples were weighed using an analytical balance and Total-Pro Solubilising
Buffer (20mM 1,4-Dithio-DL-threitol [DTT], 10mM Tris [pH 7.6], 2%SDS) added at
20 times volume to muscle weight. Samples were boiled at 95° for 10 minutes then crushed using a plastic pestle. The samples were again boiled at 95° for five minutes,
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Chapter Two: Cytoskeletal Tms in skeletal muscle
crushed again then re-boiled for five minutes to ensure complete solubilisation of
muscle sample. Samples were then centrifuged at 18,000g for five minutes and
supernatant transferred into fresh tubes. This solution was analysed according to
manufacturers recommendations to estimate the total protein concentration using the
BCA™ Protein Assay Kit (Pierce, Rockford IL, USA). Loading buffer (Laemmli [Bio-
Rad]) was added to the supernatant at a final concentration of 2x and stored at -20°C
until required.
2.2.6 Western Blotting analysis
20μg of total protein was run per sample (with the exception of EOM where 10 μg was loaded due to small volumes yielded from protein extraction). Samples were run on a
12.5% SDS PAGE gel under running buffer (20mM Tris, 190mM glycine, 1% SDS) for approximately 2 hours at 100V. Following separation of proteins by polyacrylamide gel electrophoresis, proteins were transferred to PVDF membrane (previously treated with
100% methanol) at 90 volts for 2 hours at 4°C under transfer buffer (10% methanol,
25mM Tris, and 192mM glycine). Membranes were blocked using 5% skim milk powder in TTBS (0.15M Tris, 0.2M NaCl pH 7.5 and 0.05% Tween 20) for 1 hour at room temperature or overnight at 4°C. Membranes were then washed briefly in TTBS, then incubated with the primary antibody diluted in TTBS for 2 hours at room temperature. The membranes were again washed three times for 10 minutes each then incubated with the secondary antibody for 1 hour at room temperature. The membrane was then washed three times for 20 minutes each and incubated with chemiluminescent detection solution SuperSignal Pico Substrate (Pierce) for one minute. The membrane
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Chapter Two: Cytoskeletal Tms in skeletal muscle
was then exposed to Fuji RX film in a chemiluminescent cassette and developed using
a Konica SRX-101A medical film processor. When required, band intensity was calculated by densitometry using a GS-800 calibrated densitometer (BioRad).
2.2.7 Preparation of tissue samples for cryomicrotomy
Skeletal muscles were dissected from mice. Hind- and forelimb muscles were stretched and held in place during fixation. Muscles were fixed by incubation in 2% PFA for 20-
30 minutes. Tissues were then immersed in tissue-tek solution and snap-frozen in liquid nitrogen-chilled isopentane. Tissues were stored in liquid nitrogen until required.
Transverse sections (10 μm) were cut using a HM500OM cryomicrotome (Carl Zeiss
MicroImaging, Inc., Oberkochen, Germany). Sections were placed on Starfrost poly-O-
lysine coated slides (Knittle, Germany).
2.2.8 Preparation of tissue samples for semi-thin cryomicrotomy
Immediately after dissection, mouse hind- and forelimb muscles were stretched and
held in place and all muscles were fixed in 2% PFA for 20-30 min and processed for
cryoultramicrotomy according to a modification of Griffiths et al. (1984). After
fixation, muscles were cut into strips approximately 4–5-mm long x 1-mm wide and
transferred to 1.8M sucrose/20% PVP for overnight infusion at 4°C. Muscle strips were
trimmed further, mounted on cryopins, and snap frozen in liquid nitrogen. Semi-thin
(0.5–0.8μm) sections were cut at -60°C using an Ultracut UCT ultramicrotome (Leica,
Wetzler, Germany) equipped with an EM FCS cryochamber (Leica). Sections were
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Chapter Two: Cytoskeletal Tms in skeletal muscle suspended in 2.3M sucrose, allowed to thaw, and placed on Starfrost poly-O-lysine coated slides.
2.2.9 Immuno-staining of muscle sections
Slides were blocked for one hour at room temperature or overnight at 4°C in fish gelatine blocking solution (0.2% fish gelatine, 2% BSA in PBS containing 0.05%
Triton-X 100 [PBST]). Following blocking, slides were incubated with the primary antibody diluted in PBST for two hours at room temperature or one hour at 37ºC. Slides were washed three times with PBST then incubated with fluorescent labelled secondary antibody appropriate to the primary antibody, diluted in PBST for one hour in the absence of light. Slides were again washed three times in PBST then mounted with a coverslip and one drop of Vecta-shield (Vector Laboratories, Burlingame, CA, USA).
Slides were sealed using nail polish and stored at 4°C in the absence of light. Slides were viewed using fluorescent microscopy, with different fluorescent proteins or secondary antibodies identified using specific light filters. Photographs were taken using digital photography and any adjustments made, such as addition of scale bars, contrast adjustments or superimposing of photographs, performed using software SPOT
Version 4.0.1 (Diagnostic Instruments, Sterling Heights, MI, USA).
2.2.10 Immuno-gold labeling and electron microscopy (EM) analysis
Immediately after dissection, EDL muscles were fixed in 4% PFA/0.1% glutaraldehyde/phosphate buffer pH 7.2 for 1 hr. Tissue was then incubated in 1.84M sucrose/20% polyvinylpyrrolidone for overnight infusion at 4°C and snap frozen in
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Chapter Two: Cytoskeletal Tms in skeletal muscle
liquid nitrogen. Sections were cut at 70nm at -110°C using an Ultracut UCT ultramicrotome equipped with an EM FCS cryochamber and mounted on gilded nickel
grids with a formvar/piloform film and carbon coating. Grids were blocked for 15 min
in 50mM glycine followed by 30 mins in 5% normal goat serum. Primary antibodies
diluted in immuno-incubation buffer (0.25% BSA-c/12.5mM sodium azide/PBS pH
7.4) were incubated overnight at 4°C and grids were washed in immuno-incubation
buffer. Grids were then incubated in gold conjugated secondary antibodies, F(ab)2 goat
anti rabbit 5nm gold and goat anti mouse 20nm gold diluted in immuno-incubation
buffer for two hours at room temperature, washed in immuno-incubation buffer and
then fixed in 2% glutaraldehyde/PBS for five minutes. Grids were washed in PBS and
distilled H2O then embedded for 15 minutes in 1% methylcellulose/3% aqueous uranyl
acetate diluted 9:1. Immunolabelled tissue was examined on a Philips CM10 TEM and
negatives taken.
2.2.11 Muscle fibre isolation and analysis
Isolated muscle fibres from flexor digitorum brevis (FDB) muscle were isolated and
cultured as previously described (Hernandez-Deviez et al., 2006). Isolated fibres were
cultured for 12h and then fixed in 3% PFA for 20 min, treated with 1% Triton X-100
for 5 min and blocked in PBS containing 3% BSA for 30 min. Primary antibody
incubations were carried out at 37°C for 60 min, or overnight at 4°C at appropriate
dilutions in PBS containing 3% BSA. Samples were washed with PBS and incubated
with secondary antibodies. After washing, treated fibres were mounted and viewed
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Chapter Two: Cytoskeletal Tms in skeletal muscle
using a Zeiss LSM 510 META confocal microscope system (Carl Zeiss). Single optical
sections were captured using a Plan apochromatic 63X 1.4 NA oil immersion objective.
2.2.12 Isolation of membrane components
A modification of the method described by Saito et al. (1984) was used substituting mouse muscle for rabbit and adjusting volumes accordingly. Muscles were dissected from the hindlimbs of mice and homogenized in 0.3M sucrose, 5mM imidazole-HCL pH 7.4 (homogenization buffer). Homogenates were spun at 5,000 gmax (6,000 rpm) in a
Beckman JA 17 rotor for 10 min. Pellets were resuspended and homogenized in
homogenization buffer. The homogenates were centrifuged again at 5,000 gmax for 10 min. The supernatant was taken and spun at 110,000 gmax in a Beckman SW41 rotor for
90 min. The microsomal pellet was resuspended in homogenization medium and layered onto a sucrose step gradient consisting of 45% sucrose, 38% sucrose, 32% sucrose and 27% sucrose all buffered with 5mM imidazole-HCL pH 7.4. The gradients were centrifuged for 14 hours in a Beckman SW41 rotor at 75,000 gmax (21,000 rpm).
The membrane fractions at the interfaces between the gradient steps were collected and diluted approximately two fold in 5mM imidazole-HCL pH 7.4 and centrifuged in a
Beckman TLA 100.2 rotor for two hours at 20,000 gmax (21,000 rpm). The pellets were
then processed for either protein analysis or electron microscopy.
2.2.13 Processing of isolated membranes for protein analysis
Pellets were solubilised in approximately 200µL of Laemmli buffer containing - mercaptoethanol. Samples were boiled at 90°C for 5 minutes and then loaded onto a
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Chapter Two: Cytoskeletal Tms in skeletal muscle
12.5% PAGE gel and western blotting was then performed (as per method 2.2.6) to
analyse Tm4 and calsequestrin in the sucrose gradient fractions
2.2.14 Processing of isolated triads EM
Pellets containing triads were fixed using Karnovsky’s fixative (2.5%
gluteraldehyde/2.5% formaldehyde/0.008% sodium azide/2.092% MOPS buffer)
overnight at 4°C. Following fixation pellets were washed and 10% BSA was added and
incubated for 20 minutes at room temperature. Triads were pelleted and Karnovsky’s
fixative added and incubated overnight at 4°C to cross link the BSA forming solid
blocks. Blocks were then cut into cubes. Blocks were incubated in 1% osmium
tetroxide for two hours at room temperature, and then dehydrated in ethanol at
concentrations of 50%, 70%, 95% and 100% for 15 minutes each. Dehydration was
continued with two changes of 100% ethanol for 20 minutes each followed by two
changes of 100% dry ethanol for 20 minutes each. Blocks were infiltrated with
acetone/resin in a 1:1 ratio for one hour and then further infiltrated with two changes of
100% resin for 15 minutes each at 70°C. Blocks were embedded in resin and
polymerized overnight at 70°C and stored at room temperature until sections were cut.
2.2.15 Processing of isolated triads for cryo-EM and immuno-labelling
Pellets containing triads were fixed in 4% PFA containing 0.1% gluteraldehyde for two
hours. The pellets were then washed in MOPS buffer (0.1 M 3-[N-Morpholino]
propane sulfonic acid pH 7.4) and centrifuged at 4000 gmax for two hours. Triads were re-suspended in 10% gelatine in PBS, incubated for 10 minutes at 37ºC, centrifuged
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Chapter Two: Cytoskeletal Tms in skeletal muscle again at 7500 rpm for 10 minutes and then left to set for 20 minutes at 4°C. Gelatine was cut into cubes and infused with 2.8M sucrose overnight at 4°C, then snap frozen in liquid nitrogen onto cryo-pins. Ultrathin (70nm) sections were cut at -110°C using an
Ultracut UCT ultramicrotome equipped with an EM FCS cryochamber. Sections were placed on gilded nickel grids with a formvar/piloform film and carbon coating and stained as per method 2.2.10.
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Chapter Two: Cytoskeletal Tms in skeletal muscle
2.3 Results
2.3.1 Tms are differentially expressed in skeletal muscles
It has previously been reported that mRNA transcripts encoding cytoskeletal Tms are
expressed in adult skeletal muscle (Gunning et al., 1990) and that the Tm5NM1 protein is evident in a range of mouse skeletal muscles (Kee et al., 2004). To examine the
expression of Tm proteins in specific muscle types, western blotting was performed on
a range of muscles (Figure 2.1). The antibody αfast9d recognises products containing the
9d exon from both the α and genes. In adult skeletal muscle Tm6 (40 kDa), Tm1 (38 kDa), Tm2 (36 kDa) and Tm5b (28 kDa) are found at varying levels between different muscles (Figure 2.1 A). Tm1 was found to be expressed strongly in the soleus, diaphragm and EOM, while the longissimus dorsi (LD) expresses relatively low amounts of this isoform. Both Tm6 and Tm2 are present in relatively low amounts in some muscles. Tm4 (30 kDa) is present in all muscles at varying levels (Figure 2.1 B) with unidentified bands of higher molecular weight also seen in some lanes. The muscle in which Tm4 is highly expressed are the same as those expressing Tm1 at high levels, including the soleus, diaphragm and EOM. Sarcomeric Tms (36 kDa) were expressed approximately evenly across all muscles (Figure 2.1 C). Coomassie stained bands indicated approximate even loading across all wells with the exception of the
EOM which contains half the total protein (Figure 2.1 D).
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Chapter Two: Cytoskeletal Tms in skeletal muscle
2.3.2 Tm isoforms define filaments associated with organelles in muscle fibres
Cytoskeletal Tms define distinct microfilament populations in nonmuscle cells
(Gunning et al., 1998b) and associate with distinct membrane systems (Dalby-Payne et
al., 2003;Percival et al., 2004). In skeletal muscle, it has been shown previously that
Tm5NM1 defines a novel Z-LAC compartment and is also found at the sarcolemma
(Kee et al., 2004). In order to further identify the structures that cytoskeletal Tms define in skeletal muscle, muscle sections were stained with Tm specific antibodies and markers to view various organelles within the muscle fibre. In mouse muscle, a number of filamentous compartments defined by cytoskeletal Tms were identified. The localisation of cytoskeletal Tms in transverse sections of soleus muscle was investigated using 9d, αfast9d and δ9d (Figure 2.2). An antibody to dystrophin was
used to identify the sarcolemma of muscle fibres (Figure 2.2 B, E and H). Using the 9d
antibody, Tm5NM1 is found at the periphery of the muscle fibre (Figure 2.2 A-C),
these results correspond with those previously published (Kee et al., 2004). Although
Tm5NM2 is also detected by this antibody, Tm5NM2 has not been detected in skeletal muscle using the antibody WS5/9d, which preferentially detects this isoform (Percival et al., 2004;Kee et al., 2004). Therefore, the isoform detected by the 9d antibody in skeletal muscle is Tm5NM1. Fluorescent labelling using the αfast9d antibody (Figure
2.2 D-F) revealed that Tms detected by this antibody are localised to the sarcolemma region as well as areas within the fibre. Strong fluorescence is detected at the blood vessels using the αfast9d antibody (Figure 2.2 D, arrow). Smooth muscle Tm, Tm6, recognised by the αfast9d antibody is most likely expressed in the vasculature of skeletal muscle (Vrhovski et al., 2005). The staining pattern of Tm4 (Figure 2.2 G-I), unlike the other cytoskeletal Tms, is located primarily within the sarcoplasm of muscle fibre, with some sarcolemmal staining evident. Tm4-specific antibody, δ9d, recognised filaments
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Chapter Two: Cytoskeletal Tms in skeletal muscle at the myotendinous and neuromuscular junctions (MTJ and NMJ, respectively) (Figure
2.3). At the MTJ Tm4 is located beneath the convoluted membrane (Figure 2.3 A, arrowhead), defining filaments in a network arrangement and is also found in cells that reside in the tendon itself (Figure 2.3 A-C). Tm4 localisation was examined at the NMJ
(Figure 2.3 D-F). The NMJ is viewed using bungarotoxin which binds to the ACh receptors in the membrane of the junction. Tm4 staining is seen beneath the membrane at the NMJ (Figure 2.3 D, arrowhead) and does not co-localise with ACh receptors, as indicated in the merged image (Figure 2.3 F).
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Chapter Two: Cytoskeletal Tms in skeletal muscle
2.3.3 Tm4 and Tm5NM1 define discrete actin filament populations at the Z-LAC
Transverse sections of skeletal muscle were fluorescently stained using antibodies to
Tm5NM1 and Tm4 to examine the location of these Tms in relation to the sarcomere
(Figure 2.4). Z-lines, which border the sarcomere, were detected using an antibody to
α-actinin (Figure 2.4 B and D). Tm5NM1 can be seen at the Z-LAC in longitudinal sections of stretched soleus muscle as fluorescent staining adjacent to the Z-lines
(Figure 2.4 A-C). Tm4 is also present at the Z-LAC, with staining seen adjacent to the
Z-lines (Figure 2.4 D-F). In order to determine if Tm5NM1 and Tm4 are present in the same structure double labelling was performed on muscle sections. Labelling of muscle sections with Tm4 and Tm5NM1 antibodies revealed that the filaments defined by
these two cytoskeletal Tms are not identical since the signal from TM5NM1 (red)
appeared between the lines defined by Tm4 (green) (Figure 2.5 A-D). In skeletal
muscle, the triads, consisting of the T-tubules and SR, are found adjacent to the Z-lines.
The staining pattern of Tms at the Z-LAC is similar to the localisation of the T-tubules.
An antibody to a well-characterized marker of the T-tubule system the dihydropyridine
receptor (DHPR) was used to investigate a potential relationship between the T-tubules
and Tm4 or Tm5NM1. In collaboration with Dr Delia Hernandez (IMB, Queensland)
single muscle fibres from the FDB muscle were isolated and incubated with antibodies to DHPR and either Tm5NM1 or Tm4. Tm5NM1 showed a striking co-localization with DHPR (Figure 2.5 E-G). In contrast, Tm4 did not co-localise with DHPR, but instead is present in the area between the 2 bands defined by DHPR labelling either side of the Z-line (Figure 2.5 H-J). In addition, the immuno-labelling of the isolated fibres demonstrated the presence of Tm4 and absence of Tm5NM1 in longitudinal structures
(further investigated in Chapter Three). These results confirm that these cytoskeletal
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Chapter Two: Cytoskeletal Tms in skeletal muscle
Tms define different structures and that the Tm5NM1-defined structures co-localise with the T-tubule system.
The actin backbone of the Tm-defined filaments was investigated to determine if
Tm5NM1 and Tm4 are bound to the same actin isoform at the Z-LAC. Tm5NM1 has been shown to co-localise with -actin filaments at the Z-LAC region (Kee et al.,
2004). Double labelling of Tm5NM1 and -actin (Figure 2.6 A-C) or Tm4 and -actin
(Figure 2.6 D-F) was performed to determine the actin backbone on which Tm4 resides
and compare staining between the Tm isoforms. Both Tm5NM1 and Tm4 co-localise
with -actin indicated by yellow staining in the merged images (Figure 2.6 C and F).
This result shows that the actin backbone for both Tm5NM1 and Tm4 is -actin, suggesting these Tm isoforms specify -actin for different roles within the muscle cell.
It is also possible that -actin, another cytoskeletal actin isoform, is present in this cytoskeletal filament system, however attempts to examine this isoform by immuno- staining did not reveal any staining in longitudinal skeletal muscle sections.
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2.3.4 Tm4 is associated with the sarcoplasmic reticulum (SR)
To gain further insight into Tm4 localisation, immunogold labelling and electron microscopy of frozen sections of mouse muscle was performed by Emma Kettle (MDU,
CMRI) (Figure 2.7). The triad can be seen in cryo-EM sections as negatively stained regions adjacent to the top and bottom ends of the Z-line. The terminal-SR membranes sit outside of the T-tubule, to make up the triad. The SR can be labelled using antibodies to calsequestrin, a protein situated in the terminal SR. Immuno-labelling of mouse EDL sections with Tm4 (Figure 2.7 A,D) showed the Tm4 localises to a distinct membrane element, suggestive of the sarcoplasmic reticulum (SR) in proximity to the
Z-line. To determine the nature of these membranes, sections were labelled for calsequestrin, a marker of the terminal cisternae of the SR (Figure 2.7 B,E). Co- labelling of the sections showed that Tm4 and calsequestrin localise to the same membranous structures (Figure 2.7 C,F). These results indicate that the Tm4-defined cytoskeleton associates with the terminal cisternae of the SR system. This association was further examined by muscle cell membrane fractionation. Membrane fractions were produced by ultracentrifugation of homogenates of pooled hindlimb muscle through a discontinuous sucrose gradient. Gradient fractions were analysed using western blotting techniques for the presence of calsequestrin, to identify the SR membrane fraction. Calsequestrin was expressed most strongly in fraction four (Figure
2.8 A). This fraction has been shown previously to contain the SR membrane (Saito et al., 1984). Detection of Tm4 using western blotting showed that it was present only in the SR fraction (fraction four) (Figure 2.8 A). The pellet of this fraction was processed for EM (Figure 2.8 B) and immuno-EM (Figure 2.8 C-E). Standard EM revealed the presence of SR vesicles in the preparation. Terminal SR appears as vesicles with electron dense centres (Figure 2.8 B arrow) while longitudinal SR forms a hollow
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Immuno-EM staining with Tm4 and calsequestrin (performed by Emma Kettle) revealed staining of terminal SR vesicles with both calsequestrin (Figure 2.8 C and E, large gold) and Tm4 (Figure 2.8 D and E, small gold).
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2.4 Discussion
2.4.1 Various cytoskeletal Tm isoforms are expressed in skeletal muscle
This is the first report that cytoskeletal Tms from the α-, - and δ-genes are expressed
in muscle at the level of the protein. Both Tm4 and Tm1 are expressed in skeletal at
levels comparable to expression in the brain. Immuno-labelling using the 9d antibody
on skeletal muscle transverse and longitudinal section agrees with previously published
data, that Tm5NM1 localises to the Z-LAC and regions of the sarcolemma (Kee et al.,
2004). Cytoskeletal Tms recognised by the 9d and αfast9d antibodies localise to the
sarcolemma and networks of Tm4-defined filaments are found beneath both the NMJ
and MTJ. The MTJ represents a major force transduction site in the muscle fibre
(Tidball and Daniel, 1986). The subsarcolemmal cytoskeleton at the MTJ consists of a
number of intermediate filament proteins including desmin, synemin and paranemin
(Carlsson et al., 2000). It has also been proposed that -actin plays a role at the MTJ.
Mice that contain muscles that do not express -actin exhibit a phenotype consistent with defects of the connectivity at the MTJ (Sonnemann et al., 2006). Actin stress fibres have also been localised to stress fibres within the fibroblasts of the tendon, which are shown here to contain Tm4 (Ralphs et al., 2002). The NMJ also contains a number of cytoskeletal elements including actin and intermediate filaments (Berthier and Blaineau, 1997;Ruiz-Canada and Budnik, 2006). Cytoskeletal Tms are shown to stabilise the actin filament by mechanisms including the protection of actin filaments from depolymerisation factors (Bernstein and Bamburg, 1982;Ishikawa et al.,
1989b;Ono and Ono, 2002). These properties are ideal for a cytoskeleton at the muscle membrane, NMJ and MTJ to provide stability and prevent membrane damage in these areas.
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The compartmentalisation of cytoskeletal Tms from the four Tm genes indicates that,
similar to neuronal cells, specialised cytoskeletal networks defined by Tms are evident
in muscle fibres (Gunning et al., 1998a;Gunning et al., 2005). Tm5NM1 and Tm4 were both shown in this study to be associated with membrane structures.
Compartmentalisation of Tm isoforms in association with the cellular membrane has been shown in a number of cell types. Tm5a/5b-defined stress fibres are found at the cell periphery and ruffled membranes of fibroblasts (Schevzov et al., 2005b) and also localise to the apical membranes of cultured epithelial cells (Dalby-Payne et al., 2003).
Tm2 and Tm3 are also found at the membrane in epithelial cells, however these Tms compartmentalise to the basolateral membrane (Dalby-Payne et al., 2003). Sorting of cytoskeletal Tms in neuroepithelial cells has been shown to provide specificity to actin filaments. Tm5NM1-defined filaments are rich in the myosin motor, myosin II, and depleted of ADF/cofilin. In contrast TmBr3-defined filaments contained ADF/cofilin and decreased myosin motor proteins (Bryce et al., 2003). The precise compartmentalisation and association with membrane structures involved in high mechanical stress indicates that the Tms may stabilise actin filaments for roles in these regions.
2.4.2 Tm5NM1 and Tm4 define distinct membrane-associated structures adjacent to the
Z-line in muscle fibres
In this study cytoskeletal Tms are shown to define at least two distinct filament compartments at the Z-LAC in skeletal muscle. Tm5NM1 co-localises with the T- tubules, whereas Tm4 co-localises with the closely apposed terminal sarcoplasmic reticulum. The role of the T-tubule system is to propagate action potentials to trigger calcium release from the sarcoplasmic reticulum in order to cause muscle contraction.
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This system is subjected to extreme mechanical forces during contraction and relaxation of the muscle fibre and is supported by an underlying cytoskeleton. This cytoskeletal structure has been shown to contain molecules such as the focal adhesion proteins vinculin and talin as well as membrane molecules such as dystrophin and spectrin (Kostin et al., 1998). The localisation of Tm5NM1 to the T-tubules suggests a role in stabilisation of membranes or transport in this region.
In stress fibres of neuroepithelial cells Tm5NM1 can displace ADF/cofilin, preventing the severing action of this molecule and has also been shown to accumulate on filaments containing myosin II motors (Bryce et al., 2003;Schevzov et al., 2005a).
These features suggest that a more stable filamentous structure is defined by Tm5NM1.
It would therefore be expected that myosin II motors may also be associated with these
T-tubule-associated filaments. Indeed, Takeda et al. (2000) reported that nonmuscle myosins II-A and II-B are present in the Z-lines of skeletal muscle fibers adjacent to the
Z-line in muscle fibres and it is possible that cytoskeletal Tm/actin filaments associate with these myosins. In fibroblasts and epithelial cells Tm isoforms are associated with the plasma membrane and are proposed to be involved in mechanical functions such as endocytosis of proteins at the apical membrane and the establishment of polarity
(Dalby-Payne et al., 2003;Schevzov et al., 2005b). Therefore, it is possible that
Tm5NM1 is functioning in a role similar to these at the T-tubule membrane, while Tm4 plays a role in the SR. The role of the cytoskeleton at the SR is not well-defined and it is not clear if trafficking events take place at this internal membrane system. However, it is suggested that an actin cytoskeleton is involved in positioning the terminal SR in its precise location at the level of the Z-line (Porter et al., 2005). It is then probable that
Tm4 plays a role in this process.
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We illustrate the presence of Tm5NM1 and Tm4 structures at the Z-line in close, but
distinct compartments. Tm5NM1 and Tm4 have the ability to form heterodimers in
vitro (Temm-Grove et al., 1996). However due to the clear separation of the Tm5NM1 and Tm4-defined structures, it is most likely that these filaments are composed of homodimers of each Tm isoform. In skeletal muscle fibres, -actin is present at the Z-
LAC and both Tm5NM1 and Tm4 are found to co-localise with this actin isoform in this region [this study and (Kee et al., 2004)]. This raises questions about the -actin filaments that bind these Tms in a specific and non-overlapping manner. It is hypothesized that Tms segregate to actin filaments in a ‘molecular sink’ manner whereby an isoform accumulates at an actin-based structure that has the greatest affinity for that isoform (Gunning et al., 2005). Tm isoforms binding to actin is dependant on the structural integrity of the actin filament and the availability of other actin binding proteins in that region. Therefore, we postulate that Tm4 and Tm5NM1 have different affinities for actin filaments adjacent to the Z-line that are decorated with different actin binding proteins.
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Chapter Three
Tropomyosin 4 indicates repair/remodelling in skeletal muscle disease
3.1 Introduction
Remodelling of the contractile apparatus in muscle fibres is a feature of muscle injury and disease. A role for actin has been established in the development and alignment of
the contractile apparatus (Sanger et al., 2002;Lloyd et al., 2004;Sanger et al., 2006).
The development of the contractile apparatus involves cytoskeletal isoforms of both
actin and myosin. Actin filaments and non-muscle myosin form premyofibrils initially
in muscle differentiation. These premyofibrils develop into nascent myofibrils with the
replacement of non-muscle myosin with muscle-specific myosin isoforms (Sanger et
al., 2002;Sanger et al., 2006). Nascent myofibrils then progresses to form mature
myofibrils with the alignment of Z-lines (Sanger et al., 2006). This process is detailed in section 1.4.1 of this thesis. Cytoskeletal -actin plays a role in the development of myofibrils. Early in myofibrillogenesis, myoblasts express -actin which defines stress
fibres associated with non-muscle myosins (Lloyd et al., 2004). Later in the
establishment of the contractile apparatus -actin structures are proposed to play a role in alignment of Z-lines (Lloyd et al., 2004). -actin filaments are also found in the
costameres of mature skeletal muscle. Costameres are structures that lie beneath the
sarcolemma and provide a link between the plasma membrane and the internal
sarcomeric apparatus (Rybakova et al., 2000).
The damage of muscle fibres by injury or disease results in the repair or regeneration of
the muscle (Jarvinen et al., 2005). The process of regeneration involves the activation
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of satellite cells, quiescent muscle precursor cells that reside between the basal lamina
and the sarcolemma (Ehrhardt and Morgan, 2005). These cells align and fuse to form a
myotube, the immature muscle fibre. Myotubes then undergo myofibrillogenesis to
establish the sarcomeric apparatus. Regenerated muscle fibres are characterised by
nuclei located within the centre of the fibre as opposed to mature fibres when nuclei are
located at the periphery (Franzini-Armstrong and Fischman, 1994). Focal repair of
muscle fibres following localised damage also involves activation of satellite cells. In
this instance, the repair does not involve the replacement of the entire fibre and
therefore, indicators of muscle repair such as centralised nuclei may not be readily
detected (Aarimaa et al., 2004). The processes involved in regeneration are not well-
detailed. The identification of proteins involved in regeneration and repair aids in the
understanding of this process, which may lead to new or improved treatments of
diseased and injured muscle.
Mutations in actin and Tm genes cause muscle diseases (Goebel and Muller, 2006).
Mutations in α-skeletal actin and the sarcomeric Tm isoforms, Tm and α-Tmslow
(encoded by TPM2 and TPM3 genes, respectively) cause nemaline myopathy, a disease characterized by the aggregation of thin filament proteins, termed rods, in skeletal muscle (Wattanasirichaigoon et al., 2002;Donner et al., 2002;Sparrow et al., 2003). In addition, α-skeletal actin mutations have also been found to cause actin myopathy and intra-nuclear rod myopathy (Sparrow et al., 2003)..
It has previously been reported that cytoskeletal Tm4 is present at significant levels in adult mouse muscles (Gunning et al., 1990). Cytoskeletal -actin filaments have been implicated in the development and alignment of the contractile apparatus (Lloyd et al.,
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2004). We have shown that Tm4-defines -actin filaments in skeletal muscle (Section
2.3.3). Here we examine Tm4 in the context of muscle regeneration and repair and also in muscle disease phenotypes.
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3.2 Materials and Methods
3.2.1 Specific materials
Specific materials included Ketamine (Ketavet 100, Delvet, Asquith, NSW, Australia)
and Xylazine (Ilium Zylazil-20, Troy Laboratories, Smithfield, NSW, Australia)
Notexin (Latoxan, France)
3.2.2 Animal strains
FVB/N mice, TPM3(Met9Arg) nemaline mice (Corbett et al., 2001) and mdx mice
(Tanabe et al., 1986) 2 to 4-months of age were used for all animal experiments. All
studies using mice were performed in accordance with the ethical guidelines of the
Animal Welfare Committee, National Health and Medical Research Council, Australia.
3.2.3 Human muscle samples
Human muscle tissue was taken by biopsy or at autopsy and stored in liquid nitrogen.
Samples were de-identified in accordance with The Children’s Hospital at Westmead
(CHW) ethical guidelines, and the use of the samples was approved by the CHW Ethics
Committee. Normal control biopsies: 19, 22, 25 and 36 weeks gestation, 8 days, 8 weeks, 5 years and 38 years of age (no muscle disease evident by pathology). Nemaline myopathy samples: Patient NM1 [autopsy sample, 55 year-old with TPM3(Met9Arg) mutation (Laing et al., 1995)], Patient NM2 [biopsy sample, 22 year-old with
ACTA1(V163M) mutation (Hutchinson et al., 2006)]. Duchenne muscular dystrophy biopsies: Patients DMD1 (5 year-old), DMD2 (9 year-old). Becker muscular dystrophy biopsies: Patients BMD1 (11 year-old), BMD2 (12 year-old).
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3.2.4 Primary antibodies
All primary antibodies used in this study are as described in Table 2.2.1
3.2.5 Secondary antibodies
All secondary antibodies used in this study are described in Table 2.2.2.
3.2.6 Western blotting of human muscle samples
Human biopsy samples were treated similarly to mouse samples (methods 2.2.5 and
2.2.6) with the exception that homogenization was performed on frozen muscle cryo- sections (10 sections, 8 microns thick, each measuring approx. 10 mm2).
3.2.7 Protein preparations to enrich for Tms
To enrich for Tms, tissue was sonicated in 20 volumes of 50mM Tris, pH 7.4, containing protease inhibitors (Complete EDTA Free, Roche, Indianapolis, USA) by sonication. The homogenate was boiled for 10 minutes and centrifuged at 18,000 gmax for five minutes to precipitate and remove proteins other than the heat stable and soluble Tms.
3.2.8 Immunohistochemistry of human muscle biopsy samples
Human muscle biopsy samples were sectioned in a cryostat (HM500OM; Carl Zeiss
MicroImaging, Inc., Oberkochen, Germany) at 7µm, fixed in cold 2% paraformaldehyde (PFA) for 2 min, and washed in PBS. Slides were then analysed by immunohistochemistry following method 2.2.9.
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3.2.9 Notexin induced muscle regeneration
Ketamine and Xylazine (100mg and 10mg/kg body weight, respectively) were used to anesthetize the mice. The hindlimbs were shaved and incisions made to expose the soleus or EDL muscles. 20μL Notexin at a concentration of 10µg/mL in PBS was injected directly into each muscle (Plant et al., 2006). Skin was sutured and mice left to recover for a period of 3, 5, 7 or 10 days. Three mice were anlaysed per timepoint.
Mice were sacrificed and the muscles collected for western or immunohistochemical analysis. Uninjected limb was used as control tissue. Muscle was completely broken down three days post-Notexin injection, with no intact muscle fibres evident and an abundance of mononucleated cells. After five days of recovery mononucleated cells were still evident; however, regeneration had begun as evidenced by many fibres containing centralised nuclei. At seven and ten days post-injection, mononucleated cells were reduced and increased interfibrillar spaces evident in comparison to uninjected muscles. Regeneration was almost complete at ten days post surgery (Plant et al.,
2006).
3.2.10 Mouse hindlimb immobilisation
The hindlimbs of three adult FVB/N mice were immobilised for five days as described
by Joya et al. (2004) such that the soleus muscle was lengthened and the EDL was
shortened. Following five days of immobilisation, soleus and EDL muscles were
collected for western blotting and immunohistochemistry.
Techniques including preparation of tissue for protein analysis, western blot analysis,
tissue preparation for cryomicrotomy and immuno-staining were also used in this
chapter. These methods are described in sections 2.2.5, 2.2.6, 2.2.7, 2.2.8 and 2.2.9.
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3.3 Results
3.3.1 Cytoskeletal Tm4 defines two cytoskeletal compartments in normal skeletal
muscle
Analysis of Tm4 protein expression in skeletal muscle (section 2.3.1) revealed that the
expression levels of this isoform varies between muscles. A larger set of mouse
muscles were examined with the aim of identifying a pattern between muscles. Tm4 is
expressed in all muscles examined, but at varying levels (Figure 3.1 A). In the mouse,
Tm4 is highly expressed in the soleus (Sol), flexor digitorum brevis (FDB), diaphragm
(Dia), intercostal (IC), laryngeal (Lar) and extraocular (EOM) muscles. The
longissimus dorsi (LD) in the back expresses a relatively lower amount of this Tm
along with hindlimb muscles such as the extensor digitorum longus (EDL) and tibialis
anterior (TA). A higher molecular weight 38kDa protein, recognized by the Tm4-
specific antibody and abundant in mouse embryonic fibroblasts (MEFs), also is present
in some muscles. Tm4 expression in human muscle development was also examined.
The level of Tm4 protein is significantly higher in fetal and infant muscle compared
with adult muscle (Figure 3.1 B). This correlates with the observed decrease in levels of
Tm4 mRNA during muscle development (Gunning et al., 1990). Taken together, these results reveal that Tm4 is differentially expressed between mouse muscles and during muscle maturation in humans.
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Cytoskeletal Tms define distinct microfilament populations in non-muscle cells
(Gunning et al., 1998b) and associate with distinct membrane systems (Dalby-Payne et al., 2003;Percival et al., 2004). We examined the distribution of Tm4 in sections of a variety of murine muscles (Figure 3.2). Tm4 is present predominantly in a Z-LAC orientation in the soleus (Figure 3.2 A-C), ECU, EDL and LD muscles. In the diaphragm, Tm4 is present both in Z-LAC and longitudinal filaments that are orientated perpendicular to the Z-line (Figure 3.2 D-F). EOM (Figure 3.2 G-I) and laryngeal muscles presented yet another immuno-labelling pattern with some fibres containing predominantly Z-LAC oriented structures and others longitudinal structures (Figure 3.2
G-I). A common feature of muscles that display Tm4-defined longitudinal structures is the presence of low level chronic repair, such as in the EOM which has continuous myofibre remodelling as evidenced by the up-regulation of activated satellite cell markers and developmental myosin heavy chain isoforms (Wieczorek et al.,
1985;Lucas and Hoh, 2003;McLoon et al., 2004).
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Tms exist in association with actin filaments. It has been shown by Kee et al. (2004)
and in section 2.3.3 that both Tm5NM1 and Tm4 co-localise with -actin at the Z-LAC region in muscle. Immuno-labelling of the soleus (Figure 3.3 A-C) and EOM (Figure
3.3 D-F) muscles shows that Tm4 associates with -actin filaments in both Z-LAC
(Figure 3.3 A-C) and longitudinal filaments (Figure 3.3 D-F). To investigate a role for
Tm4-containing -actin filaments in repair and regeneration in more detail we have
performed additional experiments on control and diseased mouse models as well as
paradigms for regeneration and repair.
3.3.2 Longitudinal structures defined by Tm4 are evident during myofibrillar assembly
and remodelling
To further explore whether Tm4-defined longitudinal filaments reflect repair in muscle,
Tm4 localisation was assessed in models for muscle regeneration and stretch. Notexin-
induced damage results in rapid degeneration followed by regeneration (Mendler et al.,
1998). Degeneration/regeneration was induced in the soleus and EDL muscles and Tm4
structures were monitored as the muscle fibres formed and matured. Tm4 levels
increased at least three-fold during the time course of regeneration (Figure 3.4A).
Examination of the localisation of Tm4 by immunohistochemistry revealed that Tm4 is
present in longitudinal, but not Z-LAC structures five days after muscle injury (Figure
3.5 A-C). These longitudinal filaments persist at later stages of regeneration (7 and 10
days post notexin injection; Figure 3.5 D & G, arrows); however, Z-LAC structures are
also evident (arrowheads). This suggests that Tm4 is associated with myofibrillar
formation and alignment.
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Tm4 localisation was examined during stretch-induced myofibrillar remodelling and
repair using a stretch immobilisation protocol. The hindlimbs of mice were
immobilised such that the soleus and surrounding muscles were held in a stretched position while the EDL remained relaxed. Tm4 protein expression was induced upon stretch (Figure 3.4B). The localisation of Tm4 in the stretched soleus muscle (Figure
3.6 D-F) differs from unstretched muscle (Figure 3.6 A-C) in that the tight localisation
of Tm4 with Z-LAC is lost and some Tm4 longitudinal structures are evident (Figure
3.6 D, arrow). Tm4-defined longitudinal structures are not evident in control,
unstretched muscle. The increase in Tm4 protein and altered localisation in filamentous
structures during regeneration and in stretch is consistent with a role in building,
repairing and remodelling of myofibrils.
3.3.3 Tm4 is an indicator of muscle disease
Regeneration and repair are hallmarks of muscle diseases such as muscular dystrophy
and nemaline myopathy. To determine if the localisation of Tm4-defined filaments
reflects these processes in muscle disease as it does during muscle injury, we examined
the muscles of mouse models for these diseases. Tm4 was not found at the α-actinin- rich rod region in muscle from the nemaline mouse model, TPM3 (Met9Arg) (Figure
3.7 A-C, arrow); however, longitudinal filaments were evident in the affected fibres
(Figure 3.7 A and C, arrowhead). Longitudinal filaments were also present in the muscle fibres from the diaphragms of mdx mice (Figure 3.7 D and F, arrowhead). They were particularly prominent in fibres containing centrally located nuclei and developing sarcomeres as indicated by small, closely spaced Z-lines. We conclude that the amount of Tm4 protein and its subcellular organization is a sensitive indicator of the stage of maturity of muscle fibres and the existence of repair processes in muscle.
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Tm4 expression and localisation was examined in muscle biopsies from human patients
in which repair and regeneration are characteristic features of their disease (Watchko et al., 2002). Human muscle biopsy samples from patients with both dystrophic and
myopathic pathology were analysed to determine if the Tm4-defined cytoskeletal
system was present. Tm4 protein expression was elevated in both dystrophic and
nemaline muscle (Figure 3.8) The nemaline muscles from adults had higher levels of
Tm4 relative to contractile protein (sarcomeric Tm) than adult control muscles,
although less than that of the 5 year-old control sample (Figure 3.8). The dystrophic
samples (from 5 to 12 year-old children) were elevated over the 5 year-old control
muscle sample (Figure 3.8).
Nemaline and dystrophic biopsy samples were examined for the presence of Tm4-
containing structures by immunohistochemistry. Tm4 is present at the Z-LAC in
normal human muscle (Figure 3.9 A-C). However, in nemaline muscle, Tm4-
containing longitudinal structures were detected (Figure 3.9 D,F arrowhead). These
structures are present and are more prominent in dystrophic muscle than nemaline
muscle (Figure 3.9 G-I). The lack of well-defined Z-lines in Tm4 filament-rich regions
of dystrophic fibres suggests that these structures reflect sites of immature myofibrils.
We hypothesise that the relative abundance of Tm4 longitudinal structures in nemaline
vs. dystrophic muscle reflects the level of repair/regeneration occurring in these
diseases.
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3.4 Discussion
3.4.1 Tm4-defined longitudinal filaments reflect the processes of skeletal muscle
regeneration and repair.
The establishment of the contractile apparatus occurs in myofibres during development, in regeneration following muscle injury or disease and during focal repair and stretch.
We have observed novel longitudinal structures defined by Tm4 in muscle fibres
undergoing sarcomeric remodelling/repair/regeneration. Longitudinal structures are
also present in stretch-immobilised muscle, a condition where there is an increase in the
synthesis of sarcomeric proteins leading to remodelling of myofibrils [reviewed in
(Goldspink, 1999)]. These structures are particularly prominent in muscles undergoing
extensive regeneration such as in muscular dystrophies and are also seen to a lesser degree in nemaline myopathy, a disease with focal regeneration (Sanoudou et al.,
2003). We therefore conclude that Tm4-defined longitudinal structures are a feature of sarcomeric remodelling as well as repair and regeneration.
Myopathies and injuries entail focal repair as well as degeneration and regeneration in muscle (Watchko et al., 2002;Jarvinen et al., 2005). Recently, indicators of satellite cell
activation and repair were also found in the human muscle disease nemaline myopathy.
Microarray analysis of both human and mouse muscle samples from nemaline patients
or a mouse model the TPM3(Met9Arg) mice versus control muscle showed an up-
regulation of satellite cell markers suggesting repair is taking place in these muscles
(Sanoudou et al., 2003;Sanoudou et al., 2006). Here we show that Tm4 is up-regulated at the protein level in both muscular dystrophies and nemaline myopathy and this is associated with the existence of longitudinal filaments in the diseased muscles. Muscle
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samples from both Becker and Duchene muscular dystrophy patients contain a
significant number of regenerating fibres and display extensive longitudinal structures when immuno-labelled with the Tm4 antibody. Thus, Tm4 appears to be a very good and sensitive indicator of regeneration/repair/remodelling and is potentially diagnostic of muscle disease where regeneration/repair/remodelling is a feature.
3.4.2 A Tm4/actin cytoskeleton plays a role in the repair of skeletal muscle fibres.
The pre-myofibril model of myofibrillogenesis hypothesises that stress fibres in myobasts are transformed into pre- and then nascent myofibrils in the developing myotube which in turn form the mature myofibrils of the skeletal muscle (Dlugosz et
al., 1984;Lin et al., 1989;Sanger et al., 2002;Sanger et al., 2006). Non-muscle -actin is present in stress fibre-like filamentous structures in C2C12 cultures and has been implicated in the development of the contractile apparatus during the process of muscle differentiation, (Lloyd et al., 2004). In the present study, Tm4 was found to be present in regenerating myofibres and diseased muscle in longitudinal structures similar to those defined by -actin in C2C12 myotubes. We have also observed Tm4 in stress fibre-like structures in newly formed C2C12 myotubes (Schevzov, unpublished data).
These longitudinal structures become less prominent as regenerating myofibres mature and disappear in some muscles, concomitant with an increase in prominence of the
Tm4-defined Z-LAC structures. Tm4 therefore provides a marker for stress fibre-like structures in newly formed myotubes and may aid in visualizing myofibril assembly.
Hanft et al. (2006) proposed that -actin contributes to remodelling of the costameric cytoskeleton in the absence of dystrophin(Hanft et al., 2006). Western blotting of skeletal muscle from mdx mice showed a significant increase of -actin in the
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dystrophic muscle, observed by immunohistochemistry to be located at both the
periphery and within the cytoplasm of muscle fibres (Hanft et al., 2006). The increase of both Tm4 and -actin in dystrophic muscle and the co-localisation of Tm4 and - actin in muscles undergoing chronic repair, indicates that -actin filaments decorated by this Tm isoform may play a role in this compensatory remodelling process. The Tm4- defined longitudinal structures may provide a scaffold for the assembly of the contractile proteins as has been suggested for the γ-actin based filaments (Lloyd et al.,
2004). It is also possible that these structures serve a role in shuttling sarcomeric proteins to and from sites of repair/remodelling.
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Chapter Four
The altered expression of Tm5NM1 in skeletal muscle affects membrane morphology and metabolic pathways
4.1 Introduction
The four Tm genes produce in excess of 40 isoforms using both alternate splicing and
different promoters. These isoforms confer diversity to actin filaments by attracting or
repelling other actin binding proteins. These isoforms are found to localise to specific
intracellular regions, forming separate populations of functionally discrete filaments
[reviewed in (Gunning et al., 1998b;Gunning et al., 2005)]. In addition to this, actin
binding affinities have been shown to differ between different isoforms (Pittenger et al.,
1995) and Tms have been shown to occupy slightly different positions on the actin
filament (Lehman et al., 2000). Ablation of all isoforms from a specific Tm gene or
specific isoforms has shown that Tm genes are essential, but that individual cytoskeletal
isoforms are not (Gunning et al., 2005). The ablation of the gene products from the α-
gene causes embryonic lethality, as does the specific removal of αTmfast (Blanchard et
al., 1997;Rethinasamy et al., 1998). Similarly, the attempted generation of knockout
mice by ablation of the 1b exon, removing all non-sarcomeric products of the -gene,
has shown that low molecular weight isoforms from this gene are essential for
development and cell survival (Hook et al., 2004). However, not all isoforms from this
gene are required for cell function, possibly due to compensation by other Tm isoforms.
Ablationof the brain-specific 9c exon from the -gene in knockout mice results in no
overt phenotype; however, there is a compensatory up-regulation of exon 9a-containing
products in neural tissue (Vrhovski et al., 2004).
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The over-expression of Tm isoforms also has provided information about the roles of specific Tms. Increased expression of Tm5NM1 in neuronal culture and transgenic mice has shown that Tm5NM1 impacts on the composition of stress fibres by increasing the association with myosin II and repelling ADF/cofilin (Bryce et al.,
2003;Schevzov et al., 2005a). Mice expressing high levels of Tm3 have also provided information about Tm filament populations in both neuronal cells and skeletal muscle.
The expression of this inappropriate Tm in the brain resulted in alterations in neuronal cell shape including inhibition of neurite outgrowth and a decrease in the number and length of dendrites (Schevzov et al., 2005a). In skeletal muscle this exogenous isoform localised to a region usually populated by cytoskeletal Tms, the Z-line associated cytoskeleton (Z-LAC). Here this isoform appeared to disrupt normal function of this structure, resulting in a dystrophic phenotype in these mice (Kee et al., 2004). These findings indicate that the levels and sorting of individual Tm isoforms impacts on actin filament function in particular regions of the cell.
The low molecular weight products of the -Tm gene are essential to cell development and survival, however the specific roles of the individual isoforms from the -gene have not been elucidated (Hook et al., 2004;Gunning et al., 2007). To study the roles of
Tm5NM1, a low molecular weight isoform from the -gene two strains of mice were developed: the Tm5/52 mice which express human Tm5NM1 at high levels under the
-actin promoter (Bryce et al., 2003) and the 9d/89 mice which are Tm5NM1-null due to the ablation of the 9d exon of the -gene (Vrhovski et al., in preparation). The analysis of these mice gives clues as to the specific role of Tm5NM1 in skeletal muscle.
Analysis of the knockout mice revealed alterations in the structure of regions of the plasma membrane. T-tubules, invaginations of the sarcolemma that function in calcium
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4.2 Materials and Methods
4.2.1 Specific materials
Specific materials included Ruthenium Red (ProSciTech, Kirwan, QLD Australia),
SYBR Green (Eugene, Oregon, USA), Tri-Reagent (Sigma) and Platinum Taq DNA polymerase (Invitrogen, Mt Waverly, VIC, Australia)).
4.2.2 Animal strains
Tm5NM1 transgenic and knockout mice and littermate controls at 2-4 months of age were used for experiments. Genetically modified mice are described below:
Tm5/52 mice: Mice were developed to express human Tm5NM1 gene under the control of the -actin promoter on an FVB/N background as described in Bryce et al.
(2003).
9d/89 mice: Mice were generated as described in Vrhovski et al. (in preparation).
Briefly, a knock-out construct was designed to allow for the specific deletion of exon
9d-containing isoforms from the -gene plus a neo-cassette to allow for drug
selection. The knock-out construct was electroporated into Bruce 4 C57BL/6 ES cells
and clones selected by growth in Genticin (Invitrogen). Clones were screened for the
presence of the knock-out allele, expanded in culture and injected into 129/SvJ blastocysts which were transferred into pseudo-pregnant females. Male chimeras were selected by coat colour and bred with C57Bl/6 females. Heterozygous mice were bred with mice expressing Cre under the CMV promoter to delete the neo-cassette (Schwenk et al., 1995). Mice were then bred onto a C57Bl/6J (ARC) background (9d/89 mice) and genotypes confirmed by southern blot and PCR to ensure the removal of the 9d exon and the neo-cassette.
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All studies using mice were performed in accordance with the ethical guidelines of the
Animal Welfare Committee, National Health and Medical Research Council, Australia.
4.2.3 Primary antibodies
Table 4.2.1: Primary antibodies and their sources Name Protein Recognised Antibody Type Source Ab19481 PPAR- Rabbit Polyclonal Abcam (Cambridge, UK) LC1 Human Tm5NM1 Mouse monoclonal Kind gift from Jim Lin
All other antibodies used in this study are as described in Table 2.2.1
4.2.4 Secondary antibodies
All secondary antibodies used in this study are described in Table 2.2.2.
4.2.5 Oligonucleotidess used for RT-PCR
Table 4.2.2 Oligonucleotides used and their product size Gene Forward Primer Reverse Primer Expected (5’-3’) (5’-3’) Product Size Caveolin- 2 atgacgccracagccaccacga gcaaacaggatacccgcaatg 268 bp Caveolin-3 ccgagaccccaagaacattaac cgaacaggaagccccagagca 200 bp Cofilin cgcaagtcttcaacaccaga tgaacaccaggtcctccttc 244 bp Cortactin caagaacaccagacgctcaa gtccatctggacaccgaact 201 bp GAPDH gctggcattgctctcaatgacaac gggtgcagcgaactttattcatgg 314 bp Gelsolin cctgtgtcctgggacagttt ctcctccctcttcagacacg 187 bp Kcnma caccattaagtcgggctgat ctgtccattccaggaggtgt 214 bp Myosin 8 tgaccttgagctgacactgg acttgctgttccagcttggt 228 bp Myozenin gagctctggggagcatgtag caggaacttgaagggtggaa 226 bp Ppar- atgtgtcgccttcttgctct cggtgtctgtagtggcttga 229 bp Triadin ccgaagacattgtgacaacatt atcctcatcatcttcgtcacct 263 bp
4.2.6 Ruthenium red staining of isolated muscle fibres
Muscle fibres were isolated as per method 2.2.11 as described by Hernandez-Deviez et
al. (2006). Fibres were washed in cocodylate buffer (100mM pH 7.4) (Sigma) and
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were then fixed for one hour at room temperature in 2.5% Gluteraldehyde/100mM
cocodylate containing 1mg/mL ruthenium red to stain the membranes of the muscle
fibres. Fibres were then washed three times in cocodylate buffer for ten minutes/wash
and then osmicated for three hours at room temperature in 1% osmium/100mM
cocodylate containing 1mg/mL ruthenium red. Muscle fibres were then dehydrated and resin embedded as per method 2.2.14. Ultrathin sections were cut from blocks and
placed onto grids and stained with uranyl acetate and lead citrate. Grids were viewed
using a Jeol 1010 transmission electron microscope.
4.2.7 RNA extraction from muscles for microarray analysis.
Muscles were dissected from three mice per genotype and immediately frozen in liquid
nitrogen. Samples were stored at -80°C until required for RNA extraction. Muscles for
each genotype were pooled and then immersed in TriReagent and homogenised using a
Polytron homogeniser for approximately 30 seconds. To the homogenate, 500μL of chloroform was added and the mixture vortexed for approximately 10 seconds. Samples were then centrifuged at 4,000 gmax for 15 minutes at 4°C. The clear supernatant was removed and transferred to a fresh tube where an equal volume of 70% ethanol was added. RNA was then isolated using a Qiagen RNeasy kit (Qiagen). Briefly, the solution was added to RNeasy mini column centrifuged and flow through was discarded. Columns were washed using supplied buffers according to manufacturer’s instructions and RNA was eluted from the column in sterile RNase-free water.
4.2.8 Affymetrix Gene Chip Analysis
RNA isolated using method 4.2.7 was quantified by spectrophotometry using Cary spectrophotometer and 7 μg sent to the Australian Genome Research Faciltiy (AGRF)
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where fluorescent cDNA was transcribed and hybridised to a 430_2 Affymetrix gene
chip (Affymetrix, Santa Clara, CA, USA). Results were analysed using GCOS software
Version 1.2 (Affymetrix) and GeneSifter (VizX Labs, Seattle, WA, USA).
4.2.9 RNA extraction from muscles for RT-PCR
Muscles were stored at -80°C. Each muscle was immersed separately in TriReagent and
homogenised using a Polytron homogeniser for approximately 30 seconds. Chloroform
was added and the mixture vortexed, then centrifuged at 4,000 gmax for 15 minutes at
4°C. The clear supernatant was removed, an equal volume of cold isopropanol was
added and the tubes were incubated at -20ºC overnight. Following incubation RNA was
pelleted by centrifuged at 18,000 gmax for 15 minutes at 4ºC. The pellet was washed three times with 70% ethanol and then resuspended in sterile RNase-free water
4.2.10 Transcription of RNA to cDNA
RNA was transcribed to cDNA following manufacturer’s instructions from the
ImProm-IITM Reverse Transcription Kit (Promega). Briefly, 1 μg of RNA was incubated with 20 pmol of Oligo dT at 65ºC for five minutes, then placed on ice.
Reaction mix was added that contained ImProm-IITM Reverse Transcriptase, ImProm-
TM II reaction buffer, 2.4μL of 25mM MgCl2 and dNTPs (final concentration 0.5mM each) made up with sterile water. Reaction mixture was incubated at 37ºC for 2 hours following which the cDNA product was stored at -20ºC until required.
4.2.11 Agarose gel electrophoresis
DNA was mixed with loading buffer (0.021% Bromophenol Blue, 0.021% Xylene
Cyanol FF, 0.02 M EDTA, pH 8.0, and 5% Glycerol). DNA/loading buffer mix was run
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on 0.8-1.0% agarose/TBE (0.89M Tris, 0.02M EDTA-Na2, 0.89M Boric acid) gel with ethidium bromide at 100 volts for 1-2 hours. DNA was viewed under UV light.
4.2.12 Preparation of GAPDH Standards for Quantitative PCR
The GAPDH gene was amplified from cDNA by PCR using specific primers (Table
4.2.2). cDNA was denatured at 92ºC for one minute and then was amplified by 30 cycles of denaturation (30 seconds at 92ºC), annealing (30 seconds at 62ºC) and extension (30 seconds at 72ºC). PCR products were run on an agarose gel (method
4.2.11) and the bands were cut out for purification using the QIAquick gel extraction kit (Qiagen). Briefly, DNA bands were solubilised in buffer at 50ºC for 10 minutes and one volume of isopropanol was added. Sample was added to QIAquick spin column and cDNA was bound to the membrane. The cDNA was put through a series of wash steps and then eluted in water. The concentration of cDNA was determined using Cary
300 Bio UV-visible spectrophotometer (Varian, Palo Alto, CA, USA) and appropriate dilutions made for quantitative PCR.
4.2.13 Quantitative Real time PCR
All PCR reactions were performed in a 25 μL sample containing 1.5 μL of MgCl
(Invitrogen), 2.5 μL of PCR buffer (provided with Taq kit, Invitrogen), 0.5 μL 100 mM
dNTPs, 0.2 μL (1U) Platinum Taq DNA polymerase, forward and reverse primers (1
μL or 10 μM of each) with 1 μL of the appropriate template cDNA and 1 μL of SYBR
Green. Samples were made up to 25 μL with sterile water. Reactions were carried out
on a Rotor Gene RG-3000 (Corbett Research, Sydney, Australia) under the following
conditions: an initial 95oC denaturation step for 3 min followed by 35-40 cycles of
denaturation (95oC for 30 s), annealing (appropriate temperature for 1 min) and
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synthesis (72oC for 30 seconds). Unknown samples were plotted against the standard curve generated using standard amounts of GAPDH. Samples were normalised to the amount of GAPDH present and concentrations of products calculated in ng/μL
Tissue preparation for protein analysis, western blot analysis, cryomicrotomy and immuno-staining techniques were also used in this chapter. Methods were performed as described in 2.2.5, 2.2.6, 2.2.7, 2.2.8 and 2.2.9.
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4.3 Results
4.3.1. Ablation and over-expression of Tm5NM1 does not impact on levels or localisation of other Tm isoforms.
Mice that are null for the 9d exon of the -gene and therefore do not express Tm5NM1
(Vrhovski, in preparation) and also mice that over-express human Tm5NM1 (Bryce et al., 2003) were examined to analyse the specific role/s of this Tm in skeletal muscle.
Using the 9d antibody western blot analysis of muscle samples from 9d/89 knockout and Tm5/52 transgenic mice was carried out and results confirmed that protein levels of
Tm5NM1 were altered in these mice (Figure 4.1). Analysis of soleus muscles indicated that protein levels of Tm5NM1 are reduced approximately two-fold in the 9d/89 heterozygotes, that contain one copy of the 9d exon, and are completely ablated in the
Tm5NM1-null mice (Figure 4.1, A). Using densitometry, Tm5NM1 was found to be up-regulated in the Tm5/52 mice by approximately ten-fold (Figure 4.1, B). Exogenous
Tm5NM1 can be detected using the LC1 antibody which is specific to human
Tm5NM1 protein. Exogenous Tm5NM1 is detected only in Tm5/52 mice (Figure 4.1
C).
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Kee et al. (2004) reported previously that the over-expression of Tm3 in skeletal muscle has no detectable effect on the expression levels of sarcomeric and other non- sarcomeric Tms. To examine if this phenomena also occurred with reduction or over- expression of Tm5NM1, western blot analysis was performed on protein solubilised from 9d/89 (Figure 4.2) and Tm5/52 mice (Figure 4.3) using antibodies to Tm4 and the sarcomeric Tms. Analysis of Tm4 expression revealed that this isoform was not up- regulated to compensate for the lack of Tm5NM1 in muscle (Figure 4.2, A) or down- regulated in response to excess Tm5NM1 in the transgenic mice (Figure 4.3, B).
Analysis of the sarcomeric isoforms revealed that, like Tm4, sarcomeric Tms were unaffected by the ablation (Figure 4.2, B) or increased expression of Tm5NM1 (Figure
4.3, B).
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Over-expression of Tm5NM1 in B35 neuronal cells leads to an increase in stress fibres in these cells and excess Tm5NM1 was not incorporated into additional actin-based structures (Bryce et al., 2003). This was also seen in mouse embryonic fibroblast cells
from the Tm5/52 mouse (Schevzov, in preparation). To determine if the exogenous
Tm5NM1 in the Tm5/52 mice localises exclusively to the Z-LAC or other
compartments as well, soleus muscle sections were stained to detect the presence of
Tm5NM1 (Figure 4.4). LC1 was used to detect exogenous Tm5NM1 in Tm5/52
muscle. The exogenous Tm5NM1 was found to localise to the Z-LAC (Figure 4.4, D-F)
as control muscle (Figure 4.4 A-C), showing that Tm5NM1 specifically localises to this
region of the muscle fibre.
Bryce et al. (2003) also demonstrated that the over-expression of Tm5NM1 and over- expression TmBr3 altered the localisation of other Tms in the B35 neuronal cell line thus changing the composition of the stress fibres. To determine whether the localisation of Tm4 was altered at the Z-LAC in order to compensate for the lack of or increased amounts of Tm5NM1, semi-thin sections of soleus muscle were stained for the presence of Tm4 in relation to the Z-lines (Figure 4.5). The Z-LAC staining was similar in control (Figure 4.5, A-C), 9d/89 (Figure 4.5, D-F) and Tm5/52 (Figure 4.5,
G-I) muscle fibres. Similarly, Tm4 localisation at the myotendinous junction (MTJ) was not altered in the muscles of 9d/89 (Figure 4.6, D-F) or Tm5/52 mice (Figure 4.6,
G-I) in comparison to controls (Figure 4.6, A-C).
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It was previously shown that Tm5NM1 localises to the T-tubules while Tm4 does not
(Chapter 2). We hypothesised that it was possible for Tm4 to compensate for lack of
Tm5NM1 at the T-tubules in the 9d/89 mice. To examine this in more detail soleus muscle sections were fluorescently labelled using antibodies to Tm4 (Figure 4.7, A and
D) and DHPR to identify the T-tubules (Figure 4.7, B and E). Merged images show that
Tm4 does not co-localise with the T-tubules in the control or the 9d/89 mice. Therefore, if a cytoskeletal Tm does replace Tm5NM1 at the T-tubules it is not Tm4. This also shows Tm4 specifically targets to the SR region.
4.3.2. A lack of Tm5NM1 in skeletal muscle causes abnormalities in T-tubule and caveolae morphology
Since Tm5NM1 localises to the sarcolemma and T-tubules in skeletal muscle (Section
2.2.3), it is possible that in the absence of this cytoskeletal Tm there are alterations to the structure and/or morphology of these membrane structures. To examine the morphology of the T-tubules in the absence of Tm5NM1, single muscle fibres from
Tm5NM1-null mice were stained with ruthenium red (performed by Dr Robert Parton,
IMB, QLD, Australia) to examine this membrane system at the ultrastructural level
(Figure 4.8). Ruthenium red is taken up by the plasma membrane at the sarcolemma and into the T-tubules (Figure 4.8, A arrows) and also stains the small invaginations of the plasma membrane known as the caveolae (Figure 4.8, D arrow). In comparison to control muscle fibres (Figure 4.8 A and D), muscles from the 9d/89 mice showed reduced uptake of ruthenium red into the T-tubules (Figure 4.8, B arrowheads), which may be indicative of T-tubule dysfunction. The T-tubules of the knockout muscle are also more disorganised and appear to contain more longitudinal elements than the control muscle (Figure 4.8, C arrows). The organisation of the caveolae which are lipid
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structures at the sarcolemma, differs in the 9d/89 mice. Typically the caveolae appear as darkly stained circle-shaped structures positioned below the sarcolemma (Figure 4.8,
E arrowheads). However, in the muscles of the 9d/89 knockout mice these membrane invaginations clustered into rosette-like structures. This clustering also appears in wild-
type muscle (Figure 4.8, D, arrowhead); however, much less frequently. These findings
suggest that the loss of Tm5NM1 impacts on the T-tubule and caveolae morphology
and may play an important role in the organisation and function of these membranous
structures.
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4.3.3. Tm5NM1 knockout and transgenic mice have alterations in gene expression in
soleus muscles.
Microarray technology is a high-throughput method for detection of genome wide
changes in mRNA levels (Brown and Botstein, 1999). Thousands of discrete DNA
sequences can be assayed in one experiment, allowing the comparative quantitation of
gene expression between two samples. We have used this technology to examine gene
changes between the knockout and transgenic mice with their respective control strains
to gain insight into the pathways affected by changes in Tm5NM1 expression. Three
muscles from 9d/89 and Tm5/52 and their wild-type littermate mice were collected and total RNA extracted from each sample. Samples were then pooled and 7μg of RNA was sent to the Australian Genome Research Facility (Victoria, Australia) to perform microarray analysis using the Affymetrix 430 2.0 mouse chip. Gene changes were then analysed using software by Affymetrix and VizX labs.
The microarray analyses detected a 16-fold decrease in Tm5NM1 RNA in the 9d/89 soleus muscle (Figure 4.9 A) and a three-fold increase in the Tm5/52 mice (Figure 4.9
B). Using the GeneSifter software the lists of genes deregulated by two-fold or more were categorised to assess the biological significance of gene changes. The category of cellular process was used to create pie charts to explain the processes affected by the knockout or over-expression of Tm5NM1 (Figure 4.10). The tables that follow show that the deregulation of Tm5NM1 impacts strongly on processes of cellular metabolism, regulation of physiological processes and intracellular transport (Tables
4.3.1 and 4.3.2).
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Table 4.3.1: Changes observed in the 9d/89 mice in relation to cellular process
Genes Total Cellular physiological process Up Down Altered arrayed cellular metabolism 1783 734 1049 5536 regulation of cellular 1051 413 638 3082 physiological process transport 803 321 482 2315 cell organization and biogenesis 594 243 351 1861 cell proliferation 233 90 143 620 cell cycle 229 82 147 719 cell death 207 86 121 669 cell motility 138 51 87 365 cell homeostasis 81 31 50 227 cell division 69 27 42 211 cell growth 41 13 28 129 extracellular matrix organization 31 12 19 77 and biogenesis detection of stimulus 25 10 15 63 chromosome segregation 21 10 11 54 response to extracellular 9 4 5 23 stimulus autophagy 5 2 3 19 hormone-mediated signaling 3 2 1 11 cellular pigment accumulation 1 0 1 1 reproductive cellular 0 0 0 1 physiological process
Table 4.3.2 Changes observed in the Tm5/52 mice in relation to cellular process Genes Total Cellular physiological process Up Down Altered arrayed cellular metabolism 1630 885 745 5536 regulation of cellular 971 545 426 3082 physiological process transport 771 422 349 2315 cell organization and biogenesis 545 301 244 1861 cell cycle 232 125 107 719 cell proliferation 203 101 102 620 cell death 189 110 79 669 cell motility 136 79 57 365 cell homeostasis 85 46 39 227 cell division 69 38 31 211 extracellular matrix organization 31 22 9 77 and biogenesis cell growth 30 17 13 129 detection of stimulus 24 15 9 63 chromosome segregation 22 12 10 54 autophagy 4 2 2 19 hormone-mediated signaling 4 3 1 11 response to extracellular 3 2 1 23 stimulus cellular pigment accumulation 1 0 1 1 reproductive cellular 0 0 0 1 physiological process
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Interestingly, a number of changes were evident in the expression of genes involved in
pathways of actin cytoskeleton dynamics and axon guidance, two processes Tm5NM1
is known to play a role in (Bryce et al., 2003;Schevzov et al., 2005a). Of 192 genes arrayed involved in actin regulation, 76 and 65 of these had altered regulation in the
9d/89 and Tm5/52 mice, respectively. Significant amounts of gene changes were also observed in genes involved in the calcium signalling pathway, cell adhesion molecules and regulation of adhesion and tight junctions. Complete tables can be found in appendices A and B.
Genes were chosen for further analysis by quantitative RT-PCR based on the following criteria:
• Genes known to interact with Tm5NM1 and or actin filaments (cofilin, cortactin
and myosin-8).
• Genes recognised by more than one oligo (peroxisome proliferator-activated
receptor-gamma [PPAR- ], myozenin and potassium large conductance
calcium-activated channel, subfamily M, alpha member 1 [kcnma1]).
• Genes that may explain the T-tubule or caveolae phenotype recognised in the
9d/89 mice (caveolin-2 and triadin).
Primer pairs were designed to recognise the genes mentioned above as well as caveolin-
3 to complement the study (Table 4.2.2). Primers were also designed to the GAPDH gene to act as a control.
Quantitative real-time RT-PCR was performed to confirm changes in gene expression detected by microarray analysis. Three to five mice per strain were analysed for each gene and products were expressed as ng/μL in relation to the total RNA content,
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normalised to the amount of the ubiquitously expressed GAPDH. Only one gene was
statistically significantly deregulated as detected by real-time RT-PCR: PPAR- , a
regulator of genes involved in fatty acid production and transport, adipocyte
differentiation and formation and also glucose regulation (Lopez-Soriano et al., 2006).
Results of RT-PCR of all other genes are contained in appendix C. By RT-PCR the
mRNA of PPAR- was up-regulated 5.9-fold in Tm5/52 soleus muscle (P< 0.001)
(Figure 4.11, A). An up-regulation of PPAR- was detected by microarray analysis of
9d/89 soleus; however, this up-regulation was not found to be statistically significant using quantitative RT-PCR (Figure 4.11, B). In order to determine if PPAR- protein levels reflected the changes in level of mRNA, western blotting was performed. Soleus muscle from three mice from each genotype were analysed using an antibody to PPAR-
. At the protein level an up-regulation of PPAR- (50 kDa) was detected in the
Tm5/52 mice (Figure 4.12, A), but not in the 9d/89 mice (Figure 4.12, B).
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4.4 Discussion
4.4.1 Tm isoforms from different genes are independently regulated
Collectively, the low molecular weight Tm isoforms from the -gene are essential for embryonic development and survival (Hook et al., 2004). However, individually these
-Tm isoforms are not required for cell survival. This study and Vrhovski et al. (2004) show that ablation of the 9d or 9c exons, respectively, lead to the development of mice that survive to adulthood with no overt phenotypes. The lack of phenotypes may be explained by the increased expression of other Tm isoforms from the -gene.
Compensatory up-regulation of the brain specific 9c-containing gene products was observed in the neurons of Tm5NM1-null mice (Vrhovski et al., in preparation).
Likewise, an up-regulation of exon 9a-containing products was observed in the brains of mice null for 9c-containing products (Vrhovski et al., 2004). It is possible that gene products containing exon 9a are altered in skeletal muscle of the Tm5NM1 null mice.
However, such a change cannot be detected in skeletal muscle as sarcomeric Tm isoforms also contain the 9a exon and changes using exon-specific antibodies would be masked by the abundance of sarcomeric Tms. The over-expression of exogenous Tm3 in skeletal muscle does not lead to alterations in the levels or localisation of cytoskeletal or sarcomeric Tm isoforms (Kee et al., 2004). Results from the current study agree with previous outcomes, that under- or over-expression of Tm5NM1 does not change the level or location of Tm4 or the sarcomeric Tms. This suggests that the regulation of Tms may be restricted to products from the same gene and that sarcomeric and cytoskeletal Tms are differentially regulated.
The over-expression of cytoskeletal Tm isoforms in both cell culture (Bryce et al.,
2003) and mouse tissue (this thesis) has shown that ectopic protein sorts to the same
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region as the endogenous protein. In neuronal cells, Tm5NM1 targets to stress fibres
and regions of the growth cone. Exogenous Tm5NM1 also localises to these regions
(Bryce et al., 2003;Schevzov et al., 2005a). Over-expression of TmBr3 produced
similar results with wild-type and exogenous protein localising to the lamellipodia of
neuronal cells (Bryce et al., 2003). This is also true for the over-expressed Tm5NM1 in
skeletal muscle which compartmentalises to the Z-LAC, mimicking the localisation of
the endogenous protein. The specific localisation of the exogenous Tm5NM1 in
skeletal muscle supports the theory that Tms accumulate on actin filaments that have
the greatest affinity for that particular isoform (Gunning et al., 2005). In this study, we
show that Z-LAC-localised Tm4 is unable to compensate for a lack of Tm5NM1 at the
T-tubules. This shows that Tm4 localises specifically to actin filaments at the SR. The
specific association of Tm5NM1 with -actin filaments at the T-tubule and Tm4 with those at the terminal SR is most likely due to other actin-binding proteins, capping proteins and myosin motors available in that region (Gunning et al., 2005).
4.4.2 Tm5NM1 plays a role in the organisation of membrane morphology and cellular metabolism
In this study, through use of microarray techniques and examination of ultrastructure, we have revealed that the alteration in expression of Tm5NM1 impacts on membrane morphology and muscle metabolism.
At the plasma membrane, Tm isoforms have been shown to mediate actin filament dynamics, often providing stability to the filaments by protecting actin from the severing activities of factors such as ADF/cofilin and gelsolin (Ishikawa et al.,
1989b;Ono and Ono, 2002;Bryce et al., 2003). The exclusion of Tms from actin
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filaments contributes to the fast-paced actin turn-over that occurs in regions such as the
leading-edge of migrating cells (DesMarais et al., 2002). Cytoskeletal Tm filaments
are included in the more stable region of lamellipodia beneath the leading edge
(DesMarais et al., 2002). The alteration of the Tm isoform composition in this region
by inclusion of sarcomeric Tms into the leading edge of migrating cells has been shown
to alter the actin dynamics and hence migration and morphology of these cells (Gupton
et al., 2005). In osteoclasts, Tm5a and Tm4 to localise to regions in the attachment structures in the podosome, while Tm5NM1 is located to internal structures of these migrating cells (McMichael et al., 2006). In skeletal muscle, Tm5NM1 is found in association with both the sarcolemma and T-tubules, while Tm4 localises to the SR and the convoluted membranes of the MTJ and NMJ (Chapter 2, this thesis). Collectively, these studies suggest specific, independent roles for the Tm isoforms in association with membrane structures.
The removal of Tm5NM1 from the T-tubule membrane impacts on the ability of these structures to take up ruthenium red dye, suggesting a dysfunction in the membrane structure or possibly a collapse in the T-tubule membrane, blocking access to the cell exterior. The cytoskeletal network supporting the T-tubules contains filamentous proteins including vinculin, talin and other focal adhesion molecules (Kostin et al.,
1998). In addition to these proteins, we show here that Tm5NM1 filaments contribute to T-tubule membrane structure. The actin cytoskeleton has previously been implicated in the localisation of caveolae and are said to hold these structures in place at the membrane (van Deurs et al., 2003) and disruptions of actin by cytochalasin D has been found to cause loss of caveolae in cardiac myocytes (Head et al., 2006). Membrane abnormalities are also evident at the sarcolemma of Tm5NM1-null skeletal muscle, in
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the form of abnormal clustering of caveolae structures. These disruptions to the T-
tubule and caveolae structures of the knockout mouse muscle suggest a specific role for
Tm5NM1-defined actin filaments in skeletal muscle membrane organisation or
stability.
An up-regulation of PPAR- was detected by microarray and then subsequently
confirmed by quantitative RT-PCR and protein analysis. Nine other genes found to be
deregulated by microarray were followed up by RT-PCR analysis and results were not
consistant between the two analyses. This could be explained by the sample preparation
whereby the microarray analysis was performed on pooled muscle samples from three
mice, however the RT-PCR analysis was carried out on single cDNA samples. The up-
regulation of PPAR- in the skeletal muscle of Tm5/52 mice suggests that the increased
expression of Tm5NM1 impacts on metabolic pathways. PPAR- is activated by small
lipophilic compounds and acts as a transcription factor (Semple et al., 2006). It binds
specific promoters, modulating the expression of genes involved in processes such as
fatty acid transport through the plasma membrane, cytoplasmic fatty acid binding,
lipogenesis, control of lipid metabolism, mitochondrial transport and glucose
metabolism (Wahli et al., 1995;Lopez-Soriano et al., 2006). Over-expression of this
transcription factor reduces focal adhesions at the cell membrane of cancer cells (Chen
et al., 2006). PPAR- has been shown to play a role in the insulin-activated glucose
uptake pathway (Semple et al., 2006;Sharma and Staels, 2007)]. The insulin sensitising properties of PPAR- have lead to the development of drugs which activate this
molecule. These drugs are used in the treatment of type II diabetes to reduce insulin
resistance in the adipose tissue and skeletal muscle of these patients (Lehmann et al.,
1995;Olefsky and Saltiel, 2000).
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The actin cytoskeleton is involved in a number of processes modulated by PPAR- .
Actin filaments have roles in aspects of lipid metabolism, participating in the transport of lipid bodies and fatty acid metabolites (Fatima et al., 2005;Kwiatkowska et al.,
2005). Tm5NM1 may potentially play a role in this intracellular transport by stabilising actin filaments during this process. The actin cytoskeleton also plays an essential role in the insulin-stimulated glucose uptake pathway (Tong et al., 2001;Torok et al., 2004).
The ablation of actin filaments prevents the normal uptake of glucose into skeletal muscle and adipose tissue (McCarthy et al., 2006). The association of Tm5NM1 with the T-tubules also suggests that this Tm isoform may be involved in mechanisms relating to type II diabetes and lipid metabolism as the T-tubules play a significant role in hormonal and metabolic processes in muscle (Lauritzen et al., 2006). Based on this evidence, the up-regulation of Tm5NM1 and subsequent changes in actin filament properties may cause alterations in aspects of fatty acid metabolism or metabolic defects in the transgenic mice, requiring a concomitant increase in levels of PPAR- .
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Chapter Five
Tropomyosin 5NM1 is involved in glucose transport and adipose tissue proliferation
5.1 Introduction
The regulation of glucose uptake by insulin requires the translocation of the glucose transport molecule, GLUT4, to the plasma membrane and is critical for the maintenance of glucose homeostasis. The details of the pathways leading to GLUT4 translocation are not completely understood. An increase in blood glucose stimulates the release of insulin from the pancreas, which binds the insulin receptors on the muscle or adipose cell membrane (Chang et al., 2004). The receptor complex undergoes auto- phosphorylation, which in turn causes the phosphorylation of other intracellular substrates that interact with the signalling molecule PI-3K (Foster et al., 2001;Bryant et al., 2002). This molecule plays an essential role in glucose uptake, initiating a signalling cascade that activates molecules including Akt/PKB and Rab proteins that leads to the translocation of GLUT4 vesicles to the plasma membrane. Here, SNARE proteins found on both the vesicle and plasma membranes interact to dock and fuse vesicles, incorporating the GLUT4 molecule on the membrane (Chang et al., 2004). A second pathway, independent of insulin, also facilitates glucose uptake. Platelet-derived growth factor (PDGF), another activator of PI-3K, stimulates the translocation of
GLUT4 vesicles in an insulin-independent pathway (Wang et al., 1999;Whiteman et al., 2003). Processes such as muscle contraction and a rise in intracellular calcium also result in an increase in GLUT4 translocation (Thong et al., 2007).
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The process of GLUT4 trafficking is reliant on the cytoskeleton and both actin
filaments and microtubules have been shown to be involved (He et al., 2007). Ablation of microtubule structure in adipocytes results in the diffusion of GLUT4 throughout the cell and also a shift from the insulin-dependant pathway to the PDGF pathway (Fletcher et al., 2000;Huang et al., 2005). It has been shown that actin filaments play an important role in the translocation of GLUT4 to the plasma membrane and that a loss of these filaments impairs this trafficking (Tong et al., 2001;McCarthy et al.,
2006;Kanzaki, 2006a). Specifically, actin filaments are proposed to be involved in the
insulin-stimulated pathway of GLUT4 trafficking, but not the PDGF pathway (Torok et
al., 2004). The movement of GLUT4 is facilitated by a myosin motor, Myo1c, which is recruited to actin filaments to drive the motility of the GLUT4-containing vesicle to the membrane prior to docking and fusion (Bose et al., 2002).
The phenomenon of GLUT4 translocation is diminished in type 2 diabetes, resulting in
insulin resistance (James, 2005). The mechanism by which insulin directs the insertion
of GLUT4 proteins into the plasma membrane is not fully understood and the cellular
basis of insulin resistance remains poorly explained. Induction of insulin resistance by
glucose oxidising agents or long-term exposure to insulin and glucose reduces actin
remodelling (Tong et al., 2001;Jebailey et al., 2007). This suggests that actin filaments
and the remodelling of these filaments impacts on glucose metabolism and may
contribute to insulin resistance.
The involvement of actin filaments and myosin motors in glucose uptake suggests a
concomitant involvement of actin-binding proteins such as Tms. Due to its location at
the T-tubules and sarcolemma, that are sites of GLUT4 insertion into the sarcolemma in
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muscle (Lauritzen et al., 2006), Tm5NM1 was hypothesised to play a role in the trafficking of GLUT4 and subsequently in glucose uptake and metabolism. In this study, Tm5NM1 transgenic and null mice were used to analyse the role of this Tm in the process of glucose uptake.
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5.2 Materials and Methods
5.2.1 Specific materials
Specific materials included [3H] 2-Deoxyglucose (Perkin Elmer, Rowville, VIC,
Australia), BSA (Bovogen, Essendon, VIC, Australia), cytochalasin D (Sigma), wortmannin (Sigma), insulin (Sigma).
5.2.2 Animal strains
All animal strains used in this study are described in section 4.2.4
5.2.3 Primary antibodies
Table 5.2.1: Primary antibodies and their sources Name Protein Recognised Antibody Type Source R820 GLUT4 Rabbit Polyclonal (James et al., 1989) Syn-4 Syntaxin-4 Rabbit Polyclonal (Tellam et al., 1997)
Both antibodies were kind gifts from David James
All other antibodies used in this study are as described in Table 2.2.1
5.2.4 Secondary antibodies
All secondary antibodies used in this study are described in Table 2.2.2
5.2.5 Solubilisation of muscle and adipose tissue in RIPA buffer
Tissue samples were weighed using an analytical balance and RIPA Buffer (20mM Tris
pH 7.4, 150mM sodium chloride, 1% Nonident P-40, 0.5% sodium deoxycholate, 1mM
EDTA, 0.1% SDS) was added at 20 times volume to tissue weight. Tissue samples
were incubated on ice for 10 minutes and then crushed using a sterile plastic pestle.
This process was then repeated and samples were centrifuged at 13,000 rpm for five
minutes to remove insoluble matter. The protein concentration of the supernatant was
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analysed using the BCA™ Protein Assay Kit and loading buffer was added to a final
concentration of 2x and stored at -20° until required.
5.2.6 In vitro analysis of glucose uptake in adipose tissue
Gonadal adipose tissue was removed from mice and immediately immersed in Krebs
Ringer Phosphate buffer (KRP) (12.5 mM Hepes, 120 mM NaCl, 6 mM KCl, 1.2 mM
MgSO4, 1 mM CaCl2, 0.4 mM NaH2PO4, 0.6 mM Na2HPO4, 2 mM Na-Pyruvate, 2%
BSA, pH 7.4) warmed to 37 ºC. Non-adipose tissue was removed from the adipose
tissue and the tissue was cut into small (5-10mm) pieces (adipose explants) and washed
twice in KRP buffer. Adipose tissue was incubated at 37ºC with varying concentrations
of insulin for 15 minutes before the addition of [3H]2-Deoxyglucose at a final
concentration of 1 μM for a further 10 minutes. Negative controls were prepared by
adding cytochalasin D (Sigma) prior to addition of [3H]2-Deoxyglucose. Tissue was
then washed quickly in ice-cold PBS to stop the reaction, weighed and transferred to
scintillation vials. [3H]2-Deoxyglucose was measured using a 1200CA Tri-carb liquid
scintillation analyser (Packard-Canberra, Meriden CT, USA). Results were expressed
as pmol of glucose/min/gram of tissue.
5.2.7 Wortmannin inhibition of glucose uptake
Experiments were carried out as per section 5.2.6 with the following exception.
Adipose explants were incubated at 37ºC in 100nM wortmannin solubilised in DMSO
(0.5μL of 100μM stock added to 500μL reaction solution) for 15 minutes prior to the addition of insulin. To ensure inhibition was not due to addition of DMSO, an equal volume of DMSO was added to negative controls.
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5.2.8 Glucose tolerance testing
Glucose tolerance tests were performed as per Molero et al. (2004). Briefly, glucose
(Intravenous Infusion BP 50%, AstraZeneca, Sydney, Australia) (2 g/kg glucose) was
administered by intraperitoneal injection to mice that were fasted overnight. Blood
samples were obtained from the tail tip at the times indicated in the results. Glucose levels were measured using a glucometer (ACCU-CHEK GO; Roche Diagnostics
Corporation). Blood glucose levels were plotted as both glucose index/time and as well the area under the curve was used as an indicator of difference between the two mouse strains.
5.2.9 Analysis of fat pad mass
Inguinal (groin region), retroperitoneal (around kidneys), epididymal (testis), parametrial (uterus, etc) and brown (interscapula) fat pads were removed from the mouse. Adipose tissue was weighed using an analytical balance and data combined to compare abdominal (inguinal, retroperoteneal and epididymal/parametrial) and brown fat pad masses between mouse strains.
Techniques also used in this chapter included tissue preparation for protein analysis, western blot analysis, cryomicrotomy and immuno-staining techniques. Methods were described in sections 2.2.5, 2.2.6, 2.2.7, 2.2.8 and 2.2.9
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5.3 Results
5.3.1 Tm5NM1 co-localises with proteins involved in glucose uptake
The translocation of GLUT4-containing vesicles to the membrane to facilitate the
uptake of glucose occurs at both the sarcolemma and in the T-tubule system (Lauritzen
et al., 2006). The presence of Tm5NM1 at the sarcolemma (Section 2.3.2) and co- localisation of Tm5NM1 with proteins present in the T-tubule network (Section 2.3.3)
suggests that this Tm isoform may define actin filaments involved in glucose uptake in
skeletal muscle. The co-localisation of Tm5NM1 with two proteins in the glucose
uptake pathway, GLUT4 and syntaxin-4, was analysed by immunohistochemistry in
semi-thin sections from soleus muscle (Figure 5.1). Both GLUT4 (Figure 5.1 B) and
syntaxin-4 (Figure 5.1 E) appear to co-localise with Tm5NM1 (Figure 5.1 A, D). Co-
localisation between syntaxin-4 and Tm5NM1 was seen in the T-tubule region (Figure
5.1 D-F).
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5.3.2 Deregulation of Tm5NM1 causes changes in glucose uptake and glucose
tolerance in knockout and transgenic mice
To analyse the physiological relevance of the association between Tm5NM1 and
GLUT4-containing vesicles, glucose uptake and glucose tolerance tests were performed
on mice from the 9d/89 and Tm5/52 mouse lines. For glucose uptake analysis, adipose
tissue was removed from eight to ten Tm5NM1-null mice and wild-type counterparts at
10-11 weeks of age and processed for adipose explants. These explants were then
stimulated with insulin and tritium labelled deoxyglucose was added. The concentration
of radiolabelled glucose taken up into adipose explants was expressed as pmol per
minute per gram of tissue. The glucose uptake over insulin concentrations between a
basal level (no insulin) and 10μM was plotted and curves compared between wild-type
and 9d/89 mice (Figure 5.2 A). At the highest concentration of insulin (10μM), adipose tissue from 9d/89 had a reduced rate of glucose uptake in comparison to adipose tissue from wild-type tissue (P<0.05) (Figure 5.2 B). Wortmannin was used to inhibit the PI-
3K signalling cascade, allowing the measurement of glucose uptake in the absence of this pathway (Wojtaszewski et al., 1996). Wortmannin, at a final concentration of
100nM was added to adipose explants 15 minutes prior to the addition of insulin and then glucose uptake analysis was carried out as normal. Wortmannin inhibition was expressed as a percentage of normal glucose uptake (Figure 5.2 C). Wild-type and
Tm5NM1 adipose tissue glucose uptake was reduced by 66.4% and 66.5%, respectively. If Tm5NM1-defined filaments were involved in a pathway other than the
PI-3K pathway it would be expected that there would be an increased reduction in the
9d89 adipose tissue following wortmannin treatment. The similarity in reduction of uptake indicates that the removal of Tm5NM1 from actin filaments impacts on the PI-
3K pathway of insulin-stimulated glucose uptake.
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The glucose tolerance test assesses the clearance of glucose from the blood stream.
Glucose tolerance tests were performed on male (Figure 5.3) and female (Figure 5.4) mice to examine the glucose tolerance Tm5/52 mice and wild-type littermates at 3 and
5-6 months of age. Glucose was delivered to the mice by intraperitoneal injection and blood glucose levels were collected at intervals over a 90 minute period. These values were plotted as the concentration of glucose in mM over time (Figure 5.3, 5.4 A & C).
The area under the curve was determined to compare the glucose tolerance between strains as mM per minute (Figure 5.3, 5.4 B & D). Male mice over-expressing
Tm5NM1 clear glucose more efficiently from the blood stream at 3 months of age in comparison to the control littermates (Figure 5.3 A-B). The difference between wild- type and transgenic mice increases in older mice as the Tm5/52 glucose tolerance remains approximately the same (between 1500 and 1700 mM/min), yet the glucose clearance occurs more slowly with increased age in wild-type mice. Blood glucose levels of approximately 23mM at 90 minutes in three month old mice increased to approximately 28mM over the same period of time in five month old mice (Figure 5.3
C-D). There is no difference in the blood glucose levels between female Tm5/52 and wild-type littermates at 3 months of age (Figure 5.4 A-B); however, at 6 months of age mice over-expressing Tm5NM1 clear glucose from the body more quickly than wild- type mice (Figure 5.4 C-D). Taken together, the results of the glucose uptake and tolerance tests demonstrate that Tm5NM1 facilitates the uptake of glucose into cells and/or tissues. We observed reduced glucose uptake into adipose tissue when Tm5NM1 was absent and increased glucose clearance from the blood stream when Tm5NM1 was over-expressed.
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To ensure that the alterations in glucose uptake were not due to an alteration in GLUT4
levels, protein levels of GLUT4 were analysed by western blot. A GLUT4-specific
antibody was used to examine tissue from transgenic and knockout mice and their
respective controls. Protein was solubilised from soleus muscles (Figure 5.5) and
adipose tissue (Figure 5.6) and 10μg of protein was run on 12.5% PAGE gels and transferred to PVDF membrane for western analysis (Figure 5.5 A and 5.6 A). PAGE gels were stained with coomassie to confirm equal protein loading between samples
(Figure 5.5 A and 5.6 B). The intensity of the GLUT4 band was normalised to the level of total protein in each sample by densitometry. Normalised GLUT4 (54 kDa) levels were equal between Tm5/52 mice and wild-type littermates and 9d/89 and C57Bl/6J controls in both skeletal muscle and adipose tissue. The fact that GLUT4 levels did not change levels suggests that the changes in glucose uptake and tolerance are not due to an increase in the amount of GLUT4 available to transport glucose but in the rate of
GLUT4 translocation to the membrane.
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5.3.3 Tm5/52 transgenic mice have increased body fat
Alterations in metabolic state often coincide with changes in amounts of body fat
(Fujioka et al., 1987). The up-regulation of PPAR- (Section 4.3.3) and alterations of glucose tolerance in the Tm5NM1 over-expressing mice suggest that there may be changes in body fat levels. Mice between 13 and 14 week of age were weighed and body weights were compared between transgenic and control mice (2-11 mice per group). There was no difference detected in overall body weight between wild-type and transgenic mice in both females and males (Figure 5.7 A). Fat pads from the groin region, around the kidneys and from around the reproductive organs were removed and weighed. The combined weights of these fat pads from the abdominal region were expressed as a percentage of total body weight. Brown adipose tissue from the scapula region was also weighed and expressed as a percentage of total body weight. Female
(Figure 5.7 B) and male (Figure 5.7 C) Tm5/52 mice were found to have more abdominal fat than their age-matched controls. The amount of brown adipose tissue was similar amongst the different mouse strains showing that the effect of Tm5NM1 over- expression is specific to white adipose tissue.
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5.4 Discussion
5.4.1 Tm5NM1-defined actin filaments play a role in insulin-mediated glucose uptake
In this study, we show that altering the levels of Tm5NM1 impacts on glucose uptake in both adipose tissue and at a whole body level. The role of the cytoskeleton in
GLUT4 translocation and hence glucose uptake has been studied previously, showing that both microtubules and actin filaments play roles in this process (Wang et al.,
1998;Fletcher et al., 2000). Actin filaments have been shown to direct movement of
GLUT4 from intracellular storage sites to the plasma membrane (Kanzaki and Pessin,
2001) and that rearrangement of cortical actin promotes the insertion of GLUT4
vesicles into membrane ruffles at the cell surface (Tong et al., 2001). The co-
localisation of Tm5NM1 and both GLUT4 and syntaxin-4 in skeletal muscle, in the
absence of insulin stimulation, suggests an association between players in the glucose
uptake pathway and the actin filaments defined by Tm5NM1.
Intact actin filaments are required for the translocation of GLUT4-containing vesicles
to the plasma membrane and disruption of these filaments by depolymerising agents
inhibits the movement of these vesicles (Tsakiridis et al., 1994;Omata et al.,
2000;McCarthy et al., 2006). Tm5NM1 has been shown to provide stability to actin
filaments in cell culture, protecting them from the depolymerisation factor ADF/cofilin
(Bryce et al., 2003). The increased levels of Tm5NM1 in the transgenic mouse allow
the more efficient clearance of glucose from the blood stream while the absence of
Tm5NM1 in adipose tissue decreases the rate of glucose uptake. These results suggest
that the inclusion of Tm5NM1 on actin filaments involved in GLUT4 trafficking may
stabilise the actin filament, providing a scaffold for vesicle transport.
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Members of the myosin family drive movement of vesicles along actin filaments and
are also involved in the actin-mediated re-organisation of membrane structure
[reviewed in (Tuxworth and Titus, 2000)]. The unconventional myosin, myo1C, is
present in GLUT4-containing vesicles and has been shown to co-localise with GLUT4
and actin structures in insulin-stimulated adipocytes (Bose et al., 2002). Specifically, myo1C has been shown to facilitate the fusion of GLUT4-containing vesicles to the plasma membrane by co-ordinating membrane remodelling (Bose et al., 2004). A second myosin motor protein, myosin II, is involved in GLUT4 vesicle fusion (Steimle et al., 2005). Inhibition of myosin II by blebbistatin reduces insulin-stimulated glucose uptake, but does not prevent the translocation of GLUT4 to the membrane, suggesting this myosin is also involved in membrane fusion of GLUT4 vesicles (Steimle et al.,
2005).
Tm5NM1 has also been shown to attract myosin II to actin filaments in cultured neuronal cells (Bryce et al., 2003). Myosin IIA has been shown to play a role at the membrane of migrating cells facilitating interactions between actin and microtubules at the membrane, modulating membrane dynamics in this region (Even-Ram et al., 2007).
Tm5NM1-defined actin filaments may play a role in the organisation of membrane domains for the docking and fusion events of the GLUT4-contianing vesicles. At the membrane, vesicle docking and fusion is mediated by syntaxin-4-containing SNARE complexes (Ishiki and Klip, 2005). VAMP-2 on the vesicle membrane interacts with syntaxin-4 and munc18 on the plasma membrane to attach vesicles and elicit fusion to insert GLUT4 into the plasma membrane (James, 2005;D'Andrea-Merrins et al., 2007).
In this study, we have shown that Tm5NM1 co-localises with the plasma membrane- associated SNARE protein, syntaxin-4. Taken together, these results suggest that
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Tm5NM1-defined actin filaments may play a role in the recruitment of myosin II and
subsequent alteration in membrane structure to promote GLUT4 vesicle binding and
fusion to the plasma membrane.
5.4.2 Tm5NM1 impacts on adipose tissue
Obesity has been linked to insulin resistance leading to type II diabetes (Fujioka et al.,
1987). Despite the epidemiological links between obesity and this disease, a direct
molecular and physiological link is yet to be established (King et al., 1998;Mokdad et
al., 2003;Corcoran et al., 2007). In particular, excess abdominal fat has been linked to
insulin resistance and a general impairment of glucose metabolism (Fujioka et al.,
1987;Abate et al., 1995). In this study, we have shown that the Tm5/52 mice exhibit
increased levels of abdominal white adipose tissue and an increase in glucose clearance,
presenting an interesting phenotype.
Treatment of patients with type II diabetes with PPAR- agonists such as
thiazolidinediones (TZDs) has been shown to reduce the diabetic phenotype. PPAR-
agonists enhance insulin action and improve glycaemic control in patients with type II
diabetes (Zhang et al., 2007). These drugs also cause significant weight gain as a side-
effect in patients, most likely through promotion of adipocyte differentiation and
increased lipid storage since PPAR- plays a role in these processes (Nesto et al.,
2003;Sharma and Staels, 2007;Zhang et al., 2007). However, the effect of the PPAR-
agonists on adipose tissue is positive, preventing progression of insulin resistance in
this tissue. Similar to patients treated with TZDs, the Tm5NM1 over-expressing mice
have more efficient glucose clearance and increased adipose tissue. Tm5/52 mice were
found by microarray to have increased levels of PPAR- which was confirmed by
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protein analysis in skeletal muscle (Section 4.3.3). The increase in this protein suggests a role for Tm5NM1 in adipodicity or differentiation of adipocytes to form adipose tissue. The increase in PPAR- in these transgenic mice may be the underlying cause of the obese, non-diabetic phenotype observed; however, the cause of the up-regulation in
PPAR- requires further investigation.
.
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Section Three: General Discussion
Chapter Six: General discussion and future directions
Chapter Six
Actin filaments perform a diverse range of functions in cells. Their involvement in processes such as cytokinesis, cell motility, contractile force, intracellular transport and modulation of cell morphology and cell size poses questions about how a single filament system can accomplish such a range of functions. The functional diversity of
the actin filament system is achieved in part by the expression of different actin
isoforms in different cell types (Rubenstein, 1990), however actin binding proteins are
shown to play a pivotal role in specification of function (Gunning et al., 2005). It has been established that Tm isoforms specify actin filament function and have been shown to regulate the presence of other binding proteins on microfilaments (Gunning et al.,
2005).
The inclusion of the muscle Tm isoforms, αTmfast, Tm and αTmslow, onto the thin filament is highly regulated in muscle. Alterations of Tm composition on sarcomeric thin filaments have been shown to cause major disruption to contraction. In cardiac muscle, αTmfast is the predominant Tm isoform in the thin filament. Transgenic mice in which Tm was substituted for this isoform were generated to study the functional equivalence of the two isoforms (Muthuchamy et al., 1995;Palmiter et al., 1996). The isoform substitution impacted on heart function by increasing calcium sensitivity, showing the two isoforms differ in their function (Palmiter et al., 1996). The distinct roles of specific isoforms has also been shown through mutations in Tm genes which lead to human disease phenotypes, such as familial hypertrophic cardiomyopathy
(FHC) and nemaline myopathy (NM), showing that specific isoforms are required for normal striated muscle function (Laing et al., 1995;Donner et al., 2002;Tardiff, 2005).
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This study shows that in addition to muscle-specific Tm isoforms, cytoskeletal Tms are
important for the normal function of skeletal muscle.
6.1 Cytoskeletal Tms segregate to form functionally distinct compartments in
skeletal muscle
6.1.1 Tm-defined filament populations segregate with organelles and membrane
structures
Studies have shown that Tms form functionally distinct compartments in a range of cell
types including neurons (Gunning et al., 1998b;Bryce et al., 2003), fibroblasts (Percival
et al., 2000), osteoclasts (McMichael et al., 2006), epithelial cells (Dalby-Payne et al.,
2003) and skeletal muscle (Kee et al., 2004). The current study shows that skeletal muscle contains a number of Tm isoforms that compartmentalise to define actin filaments in association with various structures including the T-tubules, SR, MTJ and
NMJ. At the Z-LAC, Tm5NM1 defines filaments that are closely associated with the T- tubules. Tm5NM1-defined filaments are also localised to the sarcolemmal membrane.
Filaments defined by Tm4 are found at the Z-LAC, closely associated with the SR and also at the MTJ and NMJ membranes. Induction of regeneration alters the orientation of
Tm4-defined filaments, which localise to compartment perpendicular to the Z-lines.
This data is summarised in Figure 6.1.
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NMJ
MTJ Gamma actin
Membrane complexes Regeneration Tm4 Tm5NM1
Figure 6.1. Cytoskeletal Tms form discrete compartments in skeletal muscle Here is shown a schematic of skeletal muscle. The sarcomeric apparatus contains
the sarcomeric Tm isoforms. Tm4 is shown in blue. This isoform is present in a compartment adjacent to the Z-lines, in association with SR and at the NMJ and
MTJ. Tm5NM1, shown in black, also compartmentalises adjacent to the Z-lines, and is associated with the T-tubule system. It is also found at the sarcolemma.
During regeneration, Tm4 is also present in longitudinal filaments that run perpendicular to the developing contractile apparatus.
6.1.2 Tm isoforms are involved in the specification of γ-actin filaments in skeletal muscle
Cytoskeletal -actin has been shown to form attachments between myofibrils and dystrophin as a component of the costamere (Rybakova et al., 2000). -actin has also been proposed to function in a compensatory mechanism in dystrophic phenotypes.
This isoform is up-regulated in dystrophic muscle and is suggested to play a role in strengthening the disrupted costameric structures (Hanft et al., 2006). The structural
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Chapter Six: General discussion importance of -actin has been shown in a muscle specific -actin knockout mouse
(Sonnemann et al., 2006). This mouse exhibits a progressive myopathy with overt muscle weakness, but no signs of membrane damage (Sonnemann et al., 2006). Despite the disease phenotype, muscles are shown to progress normally through development, leading to the conclusion that -actin is not involved in muscle development
(Sonnemann et al., 2006). This contradicts previous publications. Lloyd et al. (2004) proposed that -actin plays a role in sarcomeric assembly in cultured myoblasts.
Derived from mouse muscle, the C2C12 cell line provides a model for muscle differentiation where mononucleated myoblasts fuse to form myotubes which establish the distinct sarcomeric structures (Yaffe and Saxel, 1977). -actin is found to form stress fibres in myoblasts and then re-arranges to form striations during sarcomere development (Lloyd et al., 2004). It has also been shown that these stress fibres contain cytoskeletal Tms (Schevzov, unpublished data). Tm4 is present in stress fibre-like structures in myoblasts. These stress fibres persist during establishment of the sarcomeres, suggesting a role for Tm4 in myoblast development. The current study shows that these Tm4-defined filaments are also present during muscle regeneration, a paradigm for muscle development. We hypothesise that longitudinally orientated, Tm4- defined -actin filaments play a role in the organisation of the contractile apparatus.
The phenotype of the -actin knockout mouse somewhat disagrees with this hypothesis as skeletal muscle develops normally in these mice. However, two explanations can be offered for this discrepancy in hypotheses. Firstly, during muscle development other actin isoforms may compensate for the absence of -actin. The levels of cytoskeletal - actin were analysed in mature muscle, but not in developing muscle of the -actin knockout (Sonnemann et al., 2006). In order to discount the role of -actin in development of sarcomeres it is necessary to analyse the levels of all actin isoforms
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during development. Secondly, it would be worthwhile analysing the regenerative
capacity of muscles from the -actin knockout mouse. It is possible that the role of
Tm4-defined -actin filaments is specific to the processes of regeneration and repair and these may be affected in the knockout mouse. Induction of regeneration by a myotoxin may identify flaws in the process of muscle repair and regeneration, revealing a specific role for -actin in this process. Alternatively, subsequent analysis of protein levels of - and smooth muscle actin isoforms may show that compensation for the absence of -actin exists during muscle regeneration.
Cytoskeletal -actin has been shown to localise to the novel cytoskeletal compartment; the Z-LAC (Kee et al., 2004). The current study shows that the Z-LAC is comprised of two separate filamentous populations in association with either the T-tubules or the SR.
The structure of these membranes was not specifically examined in the -actin knockout mouse. The removal of Tm5NM1 from -actin filaments in skeletal muscle lead to abnormal morphology of the T-tubules. Analysis of the morphology of the T- tubules and SR may be valuable in understanding the role of -actin in association with these structures. Tm4-defined filaments at the MTJ appear to be affected in the -actin knockout mouse. The -actin knockout mice were found to have deficits in muscle contraction consistent with defects in connectivity between fibres and at the MTJ
(Sonnemann et al., 2006). Taken together, these results suggest that Tm4-defined - actin filaments play a role in maintaining the stability of the MTJ.
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6.1.3 Tm5NM1 plays a unique, non-essential role defining γ-actin filaments in
association with the T-tubules and sarcolemma
The lack of prominent phenotype in both Tm5NM1 knockout and over-expressing mice
suggests that this isoform may not necessarily be essential to muscle function. This has
also been seen in yeast, where the knockout of TPM2 produced no phenotype, yet
deletion of both Tm genes is lethal (Drees et al., 1995). Similarly, removal of the 9c
exon-containing products in brain produces no brain defects, yet removal of all low
molecular weight products from the -gene prevents stem cell survival and hence
development (Hook et al., 2004;Vrhovski et al., 2004). While the removal of Tm5NM1
does not cause a muscle disease phenotype it is clear that the Z-LAC structure plays an
important role in skeletal muscle. This is shown by the subtle phenotypes including the
disruption of T-tubule and caveolae morphology and the changes in glucose uptake.
The more subtle phenotypes of the knockout and transgenic mice reveal that Tm5NM1
plays unique, but non-essential roles in muscle.
It is also possible that in the absence of Tm5NM1 another cytoskeletal isoform is
present in the T-tubule and sarcolemmal associated -actin filaments in skeletal muscle.
Tm4 and sarcomeric Tms were shown to be unaffected by the removal of Tm5NM1,
however other cytoskeletal isoforms have not yet been investigated. It is hypothesised
that actin filament structures associate with Tms with a gradation in the degree of
affinity for these Tms (Gunning et al., 2005). The Tm with the highest affinity for the actin structure will be bound to the filament, but in the absence of this particular isoform the actin filament binds the next most appropriate isoform for the particular site
(Gunning et al., 2005). In neuronal cells isolated from the brains of Tm5NM1 knockout mice, 9c exon-containing products from the -gene are up-regulated in response to the
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absence of the 9d-containing Tm5NM1 (Vrhovski et al., in preparation). The inclusion
of 9c-containing gene products on the filaments in skeletal muscle is unlikely as 9c
containing isoforms are expressed exclusively in the brain (Vrhovski et al., 2004).
However, it is possible that products containing the 9a exon may be included on -actin filaments in the absence of Tm5NM1. In skeletal muscle, the detection of changes in cytoskeletal products containing exon 9a is masked by the abundance of the sarcomeric
Tms which also contain the 9a exon.
Bryce et al. (2003) studied the regulation of Tm containing structures in cultured neuronal cells. In these studies Tm5NM1 was over-expressed and the exogenous protein was shown to incorporate into stress fibres, mirroring the localisation of endogenous Tm5NM1 (Bryce et al., 2003). Similarly, exogenous TmBr3 incorporates into structures at the growth cone where endogenous protein localises in untransfected cells (Bryce et al., 2003). This is also true for the ectopic Tm5NM1 in skeletal muscle which compartmentalises to the same region as the wild-type protein, at the Z-LAC.
This shows that Tm5NM1 has a strong affinity for the -actin filaments in this distinct cellular region. The expression of an exogenous Tm from the α-gene in neuronal cells, the high molecular weight Tm3, was used to study the specificity of Tm localisation.
Tm3 was distributed throughout the cell in a pattern of localisation similar to α-Tm gene products, Tm5a/5b. These genes differ only in their amino terminus. This phenomena suggests that the specificity of sorting may be due to the differences in protein sequences of the different Tm isoforms from different genes (Gunning et al.,
2005;Schevzov et al., 2005a).
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The altered expression of Tm isoforms has shown to impact on the composition of actin
filaments. Tm5a/5b, normally localised to the neurite is altered in Tm5NM1 over- expressing neurons and is present in the growth cones and absent from the neurite
(Schevzov et al., 2005a). This was also shown in cultured cells, where increased levels of Tm5NM1 leads to the inclusion of Tm5a in stress fibres (Bryce et al., 2003).
Tm5NM1 has been shown to form heterodimers with Tm5a/5b (Temm-Grove et al.,
1996). It is proposed that, when present at high levels, Tm5NM1 recruits Tm5a/5b to the growth cone and forms heterodimers with these isoforms (Schevzov et al., 2005a).
Tm5b is present in a number of muscles (Figure 2.1). Any changes in the localisation of this Tm in skeletal muscle would be difficult to detect as the antibody used to recognise
Tm5b, αfast9d, recognises other Tm isoforms in muscle. However, the relationship between Tm5NM1 and Tm5b in the skeletal muscles of Tm5/52 mice should be examined to detect possible differences in filament composition in these muscles.
Altered filament composition could play a role in the changes in glucose uptake seen in the Tm5/52 mice.
Studies have also shown that the increased levels of particular Tm isoforms lead to alterations in cell morphology and motility (Bryce et al., 2003;Schevzov et al., 2005a).
Embryonic neuronal cells isolated from Tm5/52 mice, over-expressing Tm5NM1, displayed significantly enlarged growth cones and an increase in axonal and dendritic branching (Schevzov et al., 2005a). In contrast the same cell type isolated from mice over-expressing the high molecular weight Tm3 displayed decreased neurite outgrowth and branching (Schevzov et al., 2005a). These alterations in cell structure implicate specific Tm isoforms in the maintenance of cell morphology. From this, it may be important to look more closely for morphological changes in the Tm5NM1 over-
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expressing mice. Further examination of Tm5NM1-containing filaments in association with T-tubules may also reveal alterations in filament composition contributing to the changes in the functioning of these filaments in the glucose uptake pathway.
6.1.4 Cytoskeletal Tm filaments are proposed to associate with other actin binding proteins in skeletal muscle
Actin binding proteins play an integral role in the function of the actin filament, co-
ordinating the dynamics of the filament. Tm isoforms have been shown to alter actin
filament organisation by recruiting or inhibiting other actin binding proteins. Studies of
the association between Tms and various proteins are often performed using only one
Tm isoform. Drawing conclusions from these results, can lead to inaccurate statements
about the association of or competition between Tms with proteins, as different Tm
isoforms have been shown to associate with molecularly distinct actin filaments (Bryce
et al., 2003;Gunning et al., 2005). Understanding the interactions between various
proteins and the different Tms isoforms is essential to determining the functions of
these isoforms.
ADF/cofilin proteins modulate actin dynamics by enhancing the rate of filament
turnover by severing and depolymerising actin filaments (Bamburg et al., 1999;Ono,
2003). It was originally proposed that cofilin inhibits the interactions of actin with Tm
(Nishida et al., 1984). Subsequent studies identified that the reciprocal of this was also
true with inclusion of Tm isoforms on actin filaments shown to inhibit ADF/cofilin
binding (Ono and Ono, 2002). Bryce et al. (2003) showed that the association of
ADF/cofilin with the actin/Tm filament differed depending on the Tm isoform present.
TmBr3 was shown to immunoprecipitate with ADF-containing actin filaments. In
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Tm5NM1 (Bryce et al., 2003). In cultured skeletal muscle cells ADF/cofilin is localised to the I-band (Nakashima et al., 2005). The staining pattern of ADF/cofilin shown by
Nakashima et al. is similar to the pattern of localisation of Tm isoforms in the Z-LAC.
As Tm5NM1 does not associate with ADF/cofilin, the presence of this Tm isoform in close proximity to regions containing ADF/cofilin suggests that Tm5NM1 is protecting
-actin filaments from depolymerisation. In this way Tm5NM1 may be creating a more stable filament in association with the T-tubules.
Non-muscle myosins play a role in the process of myofibrillogenesis (Sanger et al.,
2002;Sanger et al., 2006). Early on in sarcomere development non-muscle myosins are present in immature myofibrils. These are replaced with muscle specific isoforms as the myofibrils mature (Sanger et al., 2006). The presence of non-muscle myosins in mature muscle has not been studied in detail. One region that has shown to contain non-muscle myosins is the NMJ (Vega-Riveroll et al., 2005). Both myosin IIA and IIB have been shown to localise to the NMJ and are presemt beneath the membrane and adjacent to the ACh receptors (Vega-Riveroll et al., 2005). We have shown in this study that Tm4 also localises to this region. Unlike Tm5NM1, there have not yet been studies to show that Tm4 recruits non-muscle myosins. However, it is possible that at the NMJ Tm4 and myosin IIA/IIB decorate the same actin filaments.
The process of glucose transport in adipocytes has been shown to involve non-muscle myosins (Bose et al., 2002;Bose et al., 2004;Steimle et al., 2005). GLUT4 vesicles translocate from intracellular storage sites to the membrane of the cell in an actin dependent manner. At the membrane myosin IIA and IC have been shown to be
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involved with GLUT4 vesicle fusion (Bose et al., 2004;Steimle et al., 2005). These myosins modulate membrane dynamics, enabling the vesicle to attach and fuse to incorporate the GLUT4 protein in the plasma membrane. Bryce et al. (2003) showed that Tm5NM1 recruits myosin II to actin filaments. We have shown that Tm5NM1 plays a role in glucose uptake, this process being affected by increased or lack of
Tm5NM1. This isoform is present at two membrane regions in skeletal muscle; at the sarcolemma and the T-tubules. Taken together, these results implicate Tm5NM1- containing filaments, in association with myosin motors, in the fusion of GLUT4 vesicles at the membrane. It may also be suggested that GLUT4 vesicles use a non- muscle myosin motor to facilitate translocation along Tm5NM1-containing actin filaments.
A novel actin binding protein, Xin, has been shown to localise to stress fibres in C2C12 cells in culture (Sinn et al., 2002). In these cells Xin was found in stress fibre-like structures and co-localised with sarcomeric Tms recognised by the CH-1 antibody
(Sinn et al., 2002). This protein was shown to bind actin through a binding motif termed the Xin-repeat region (Pacholsky et al., 2004). When bound to actin, the Xin- repeat region was found to stabilise stress fibre-like structures in cultured cells shown by resistance to depolymerisation by Latrunculin A (Pacholsky et al., 2004). In mature muscle Xin localises to the MTJ, co-localising with vinculin in this region (Sinn et al.,
2002). Subsequent studies have shown that Xin is associated with components of the troponin complex, gelsolin and actin itself (Jung-Ching et al., 2005). The current study has shown that Tm4 accumulates at the MTJ and unpublished work by Schevzov et al.
(2006) has demonstrated that Tm4 is present in stress fibres in cultured C2C12 cells.
Taken together, this data suggests that Tm4 may associate with Xin on -actin
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filaments. Pacholsky et al. (2004) investigated the relationship between Xin and Tm in an in vitro assay using the repeat domains of the Xin protein and concluded that Xin repeats inhibit Tm binding to the actin filament. This theory was supported by identification that Xin binds the actin filament at the same position as Tm
(Cherepanova et al., 2006). This result does not explain the co-localisation seen between Xin and Tms in stress fibres (Sinn et al., 2002). The difference may be due to the Tm isoforms analysed in these studies. Pacholsky et al. (2004) used smooth muscle
Tm, a high molecular weight isoform, in contrast to the low molecular weight Tm4.
Different Tm isoforms have been shown to have different affinities for actin binding proteins, such as ADF/cofilin, this may be the case with Xin (Bryce et al., 2003).
Different Tm isoforms have also been shown to occupy different positions on the actin filament, which may be the case with Tm4 and the smooth muscle Tm analysed in conjunction with Xin binding (Lehman et al., 2000). Conversely, individual filaments at the MTJ and in stress fibres may contain either Xin or Tm4. It is hypothesised that
Xin and Tm4 work to stabilise the actin filament, therefore it is possible that actin filaments require only one of these two proteins.
Caveolae are invaginations of the plasma membrane that contain cholesterol, sphingolipids, and the structural protein caveolin (van Deurs et al., 2003). The altered morphology of the caveolae in skeletal muscle from the Tm5NM1-null mice suggests that this Tm isoform may interact with proteins present in the caveolae for accurate localisation of these structures. The principal component of caveolar membranes is a protein known as caveolin, with caveolin-3 the predominant isoform expressed in muscle (Tang et al., 1996). Similar to Tm5NM1, caveolin-3 is shown to localise to the sarcolemmal and T-tubule membranes in skeletal muscle (Ralston and Ploug, 1999).
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Mutations in caveolin-3 lead to a deficiency in the protein and cause a limb girdle
muscular dystrophy (Galbiati et al., 2001). In patients carrying mutations in caveolin-3, skeletal muscle contains reduced amounts of caveolae structures and those present had abnormal morphology (Minetti et al., 2002). The T-tubules were also found to be disrupted (Minetti et al., 2002). In addition to this caveolin-3 knockout mice displayed
increased adipodicity and changes in metabolic state (Capozza et al., 2005). The similarities in localisation patterns of caveolin-3 and Tm5NM1 and the phenotypes of mice lacking either of these proteins suggest that the two proteins may interact in some way. A role for actin has been established in the maintenance of the position of caveolae in the membrane (van Deurs et al., 2003). The disruption of actin filaments by
cytochalasin D in cultured cells results in the reduction of caveolae structures at the
membrane (Head et al., 2006). We suggest that Tm5NM1 may decorate actin filaments
involved in caveolae structure and function.
6.2 A role for Tm4 in the regeneration and repair of muscle tissue
The current study shows that Tm4 is up-regulated in muscle diseases in both human
patients and mouse models of human muscle disease. This up-regulation is also seen
upon induction of regeneration and repair by administration of a myotoxin or by stretch immobilisation. The increased Tm4 coincided with the presence of longitudinally oriented Tm4-defined filaments which ran perpendicular to the sarcomeric apparatus.
Here, the role of Tm4 in the processes of regeneration and repair is suggested and the implications of this role in disease phenotypes are investigated.
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6.2.1 A role for Tm4 in organisation of muscle structure during regeneration/repair
The presence of Tm4-defined longitudinal filaments in regenerating and repairing
muscle and its association with the SR in mature muscle suggests a role for Tm4 during
regeneration in the organisation of SR membranes. For the correct function of skeletal
muscle the SR must be positioned correctly (Takekura et al., 2001). Following notexin-
induced degeneration of soleus and EDL muscle, markers of the SR are depleted
(Mendler et al., 1998;Zador et al., 1998). At the electron microscope level, changes in the SR are detected two days after induction of regeneration by a myotoxin, with muscle displaying dilated and fragmented terminal SR (Politi et al., 2006). Markers of
the SR begin to recover at three days post injection, increasing further by day ten of
regeneration (Mendler et al., 1998;Zador et al., 1998). This increase in the presence of
SR proteins coincides with the appearance of longitudinal Tm4 filaments in
regenerating muscle. During development of the muscle fibre the SR are found to
associate with the Z-lines early on (Takekura et al., 2001). The positioning of these
membrane structures at the Z-line may be reliant on the Tm4-defined filaments.
The activation and subsequent fusion of satellite cells is essential for the process of
regeneration during disease or following injury (Charge and Rudnicki, 2004). Once
activated the satellite cells proliferate and then align and fuse to form myotubes which
mature to form a new muscle fibre (Charge and Rudnicki, 2004). Tm4 is found not only
in longitudinal filaments but is also expressed in satellite cells and during regeneration.
The migration of the satellite cell to the centre of the muscle bed requires a number of
cellular processes (Dedieu et al., 2004). Actin filament rearrangements are required for
the formation of projections of the cell membrane which are necessary for the migration
of the cell (Lambrechts et al., 2004). The actin-rich lamellipodia and filopodia project
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Chapter Six: General discussion from the leading edge of the cell to coordinate the forward movement of the cell into its new location (Lambrechts et al., 2004). In migrating myoblasts disruptions to the actin cytoskeleton by mRNA knockdown of calpains, result in decreased migration of these cells (Mazeres et al., 2006). In neuronal cells Tm4 is proposed to be involved in cell motility and growth of neuronal processes (Had et al., 1994). In the satellite cell Tm4 may play a role in the organisation of actin filaments for migration of these cells.
Actin filaments are also involved in the positioning of the nucleus in a range of cell types (Starr and Han, 2003). Stable actin filaments anchor the nucleus into a set position, while dynamic filaments are involved in force production for movement of nuclei (Starr and Han, 2003). During regeneration the myonuclei are aligned in the centre of the fibre and the presence of centralised nuclei is a hallmark of a regenerated fibre (Ehrhardt and Morgan, 2005). In skeletal muscle that has been induced to undergo degeneration and subsequent regeneration Tm4 staining surrounds centrally located nuclei (Figure 3.5). This staining pattern suggests that Tm4 may define actin filaments that are involved in the positioning of the central nuclei in the regenerating fibre.
During the regeneration of muscle fibres the contractile apparatus is assembled in a fashion that mimics muscle development (Bischoff, 1994). The sarcomeres develop and the Z-lines then align to form the ordered array of myofibrils in the repair muscle fibre.
In C2C12 cells stress fibres, composed of -actin, are found in the early stages of sarcomeric assembly (Lloyd et al., 2004). Tm4 has also been shown to localise to these structures in C2C12 cells (Schevzov, unpublished data). These -actin stress fibres relocate during assembly of the contractile apparatus to flank the Z-lines (Lloyd et al.,
2004) in a pattern similar to what we have found in mature muscle. These structures are
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proposed to form a ‘scaffold’ for the construction of the sarcomeres and function in the alignment of the Z-bodies and Z-lines (Lloyd et al., 2004). The longitudinal filaments found in regenerating muscle resemble stress fibre-like structures. Tm4 also may define
-actin filaments which function in the coordination of the correct assembly of sarcomeres.
6.2.2 A role for Tm4 in muscle disease
The plasma membrane of the muscle fibre is very susceptible damage due to the high mechanical activity the muscle experiences (McNeil and Steinhardt, 2003). Therefore it is important that the cells are able to maintain sarcolemmal integrity under mechanical stress and effectively repair membrane damage (Bansal et al., 2003). The inability to effectively repair damaged muscle fibres has been shown to lead to a form of muscular dystrophy (Bansal et al., 2003). Mutations in the dysferlin gene have been shown to cause limb girdle muscular dystrophy type 2B and also Miyoshi myopathy (Bansal and
Campbell, 2004). Muscles with defects in dysferlin have a stable plasma membrane; however, they slowly develop progressive muscular dystrophy. This dystrophy is caused by defects in the mechanisms of muscle fibre repair (Bansal et al., 2003;Bansal and Campbell, 2004). The up-regulation of Tm4 during regeneration, repair and disease pathologies and the presence of longitudinally orientated filaments implicates this Tm in the process of muscle regeneration. Defects in Tm4 may lead to a muscular dystrophy phenotype similar to that seen with defects in dysferlin. If Tm4 is unable to function in the regeneration of muscle tissue, damage caused by mechanical stress or muscle injury may not be repaired as effectively leading to a dystrophic pathology.
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6.3 A role for Tm5NM1 and other cytoskeletal Tms in intracellular transport
Actin has been established as a component in the pathway of vesicle transport (DePina and Langford, 1999;Stamnes, 2002;Eitzen, 2003). Actin filaments have been shown to play roles in the transport of vesicles to the membrane from intracellular storage sites
(Kanzaki, 2006b), membrane dynamics during the fusion of vesicles to the membrane
(Eitzen, 2003) and in the endocytosis pathway (Smythe and Ayscough, 2006).
6.3.1 Cytoskeletal Tms are involved in the transport of vesicles to the membrane.
The current study proposes a role for Tm5NM1 in the process of GLUT4 translocation
from intracellular storage sites to the plasma membrane. The specific function of
Tm5NM1 in the translocation of GLUT4-contianing vesicles has not yet been
deciphered but increased or decreased expression of this Tm isoform impacts on
glucose uptake. We have proposed that Tm5NM1-defined actin filaments facilitate the
movement of GLUT4 vesicles to the plasma membrane, where these filaments are
involved in myosin II directed fusion of vesicles to the membrane (Steimle et al.,
2005).
A role has been established for Tms in the transport of secretory vesicles in budding
yeast (Pruyne et al., 1998). The Tm encoded by TPM1, one of the two Tm genes in
yeast cells, binds to actin cables in yeast cells. In the absence of TPM1, yeast cells lack
actin cables and accumulate vesicles (Liu and Bretscher, 1992). The formation and
accumulation of vesicles is unaffected; however, the cells lose the ability to directly
secrete these vesicles, suggesting a role for Tm-containing actin cables in this process
(Liu and Bretscher, 1992). The disruption of this protein also impacts on the
localisation of the yeast myosin isoform Myo2p, suggesting that the localisation of
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myosin to the actin cables is reliant on Tm (Pruyne et al., 1998). In addition to this, the disruption of both Tm and myosin genes in yeast leads to lethality, suggesting that Tm and myosin interactions are required for the important secretory pathway (Liu and
Bretscher, 1992). In tissue from Tm5NM1 knockout mice, the rate of glucose uptake is reduced suggesting a problem in the transport of GLUT-containing vesicles to the plasma membrane. The role of Tm5NM1 in this pathway may be similar to the role of
Tpm1 in vesicle secretion in yeast. Tm5NM1 may play a role in the maintenance of the integrity of actin filaments, which are required for efficient vesicle transport.
Interactions exist between Tm and myosin in the secretory pathway in yeast. We have also suggested this may be the case in the translocation of GLUT4. These interactions between Tm5NM1 and myosin have been discussed previously (Section 6.1.4).
A role for cytoskeletal Tms has been shown in the transport of the cystic fibrosis transmembrane conductance regulator (CFTR) in epithelial cells (Dalby-Payne et al.,
2003). This protein is present in the plasma membranes of epithelial cells and plays a role in the regulation of ions and water (Ganeshan et al., 2007). Impaired expression of
CFTR at the plasma membrane hinders the cell’s ability to clear chloride ions from the cell, creating an osmotic imbalance as water cannot leave the cell. This causes thickening of the mucus in the exterior of the cell leading to the disease phenotype cystic fibrosis which is characterised by the accumulation of mucus in organs such as the lungs and pancreas (Okiyoneda and Lukacs, 2007). Disruption to actin filaments in epithelial cells impacts on the regulation of CFTR at the membrane. Incubation of cells with the actin filament disrupting agent Cytochalsin D causes an increase in chloride ion transport at the membrane, indicative of an increase in CFTR at the membrane (Prat et al., 1995). The prevention of actin filament formation by DNAse-1 inhibits cAMP-
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stimulated chloride secretion (Prat et al., 1995). Prat et al. (1995) concluded that these
findings were consistent with the hypothesis that activation of the CFTR is dependent
on short actin filaments, rather than actin monomers or long filaments.
The association of Tm isoforms Tm5a/5b with CFTR has been shown in epithelial cells
(Dalby-Payne et al., 2003). Knocking down the amount of Tm5a/5b in epithelial cells leads to an increase in CFTR at the cell surface and increased chloride efflux (Dalby-
Payne et al., 2003). Myosin VI has been proposed to facilitate the endocytosis of CFTR from the plasma membrane (Swiatecka-Urban et al., 2004). Inhibition of the function of the actin-dependant myosin VI resulted in a decrease in the endocytosis of CFTR,
consistent with an increase in surface expression of this protein (Swiatecka-Urban et
al., 2004). Taken together these results suggest a role for Tm5a/5b in the endocytosis of
CFTR. The results seen following reduction of Tm5a/5b were in contrast to the
reduction of Tm5NM1, which lead to decreased transport of GLUT4 to the membrane.
These differences may imply that specific Tm isoforms are involved in the various
aspects of vesicle transport. Tm5NM1 may be involved in the transport of vesicles to
the membrane; whereas, Tm5a/5b is involved in the internalisation of vesicles from the
plasma membrane. The different roles of these isoforms may also be indicative of
differences in vesicle trafficking in various cell types and tissues.
6.3.2 Implications of aberrant Tm expression in diseases related to vesicle trafficking
The incidence of type II diabetes has increased over the last 10 years as a consequence
of elevated incidence of obesity (Mokdad et al., 2003). Patients develop insulin
resistance, a condition whereby the insulin-stimulated glucose uptake is low in tissues,
leading to high levels of glucose remaining in the blood. Insulin release is then
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increased leading to a state of hyperinsulinemia. Eventually this increased insulin
production leads to failure of the -cells of the pancreas, dropping insulin levels leading to glucose tolerance (Nehlin et al., 2006). A role for Tm5NM1 in the uptake of glucose into cells, implicates this Tm in the early stages of a diabetic disease state. A lack of
Tm5NM1 leads to a reduction of glucose uptake into adipose tissue. Mutations in the
9d exon of the -Tm gene may lead to a reduction in protein expression in humans
which may, in turn, predispose the carrier of the mutation to type II diabetes.
A therapeutic option for the treatment of type II diabetes may involve delivery of the
Tm5NM1 protein to patients. The increased levels of Tm5NM1 in the transgenic mice increase the clearance of glucose from the blood stream, possibly by a more rapid delivery of GLUT4-containing vesicles to the plasma membrane. Studies using TZDs,
PPAR- activators, have shown that these glucose lowering agents have follow-on effects leading to improvements in peripheral and hepatic insulin sensitivity (Yamauchi et al., 2001). These drugs also cause a weight gain, most likely due to an increase in
adipocyte formation. Detailed studies on the adipose tissue shows that TZDs have
effects on the morphology of adipose cells and also reduce hepatic fat, said to
contribute to the improvements in the disease state (Sharma and Staels, 2007). The
phenotype of the Tm5NM1 mice indicates that the increased levels of this Tm mimic
the action of TZDs in some respects. Further investigation is required including
examining hepatic fat and adipocyte morphology to understand the mechanisms of
Tm5NM1 in the metabolism of glucose.
The identification of a cytoskeletal Tm isoform in one vesicle transport pathway opens
the door for analysis of cytoskeletal Tms in other pathways of vesicle transport.
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Diseases such as Dent’s disease, which results in progressive renal failure, and Usher’s syndrome, resulting in blindness and deafness, arise from defects in vesicle transport
(Howell et al., 2006). Investigations into pathways of vesicle secretion should take into account the possibility that cytoskeletal Tms and their actin backbone play a role in the pathogenesis of disease.
6.4 A role for cytoskeletal Tms in membrane stability and dynamics
It has been shown that actin dynamics play an important role at the membrane in processes of migration and vesicle fusion (Pollard and Borisy, 2003;Eitzen, 2003). Tm isoforms have also been shown to associate with membranes in neurons, erythrocytes, epithelial cells and skeletal muscle (Bryce et al., 2003;Dalby-Payne et al., 2003;Kee et al., 2004;An et al., 2007) In this study we show that in skeletal muscle cytoskeletal
Tms associate with various membranes including the sarcolemma, MTJ, NMJ, T-
tubules and the SR.
6.4.1 Cytoskeletal Tms play a role in membrane stabilisation in skeletal muscle.
In the current study we have identified a cytoskeletal Tm, Tm4, in association with the
NMJ and MTJ. These two regions of the sarcolemma have been shown to contain a
complex underlying cytoskeleton including proteins such as intermediate filament proteins: desmin, vinculin and talin. An actin cytoskeleton has been found in both of these regions together with actin associating proteins such as α-actinin and Tm2 (Bloch and Hall, 1983;Marazzi et al., 1989). The presence of Tm4 in these regions implicates
this Tm in the stabilisation of actin and also the membrane structure. Tm isoforms have
previously been shown to stabilise membrane structures. The plasma membrane of the
red blood cell undergoes dramatic distortions due to the demands of circulation. Due to
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this a complex cytoskeleton comprised of spectrin tetramers and actin filaments is
found beneath the membrane, to confer elastic properties, while still providing the stability necessary to protect the membrane from damage (An et al., 2007). Tm has been shown to be a component of this cytoskeletal structure, with Tm depletion shown to cause reduced resistance to membrane shearing (An et al., 2007). Introduction of a skeletal muscle isoform in place of the erythroid Tm (Tm5a/5b), does not compensate efficiently showing that isoform specificity is important for maintenance of membrane stability in erythrocytes (An et al., 2007). The specific inclusion of Tm4 onto actin filaments at the sites of high force transduction suggests that the properties of this isoform are required for membrane stabilisation. Tm4 is also found in the fibroblast- like cells present in the tendon itself (Figure 2.3). Actin has been shown to play an important role in the organisation of these cells in the tendon, forming stress fibres which maintain links between cells and provide a basis for sensing mechanical strain
(Ralphs et al., 2002). Tm4 in these cells may stabilise the actin filaments that maintain the integrity of cell-cell contacts and hence provide stability to the cell as a whole.
Tm5NM1 is shown in the current study to localise to the T-tubule membranes of skeletal muscle. Removal of Tm5NM1 from skeletal muscle results in alterations of the
T-tubule morphology and also an altered ability for uptake of Ruthenium Red dye into this membrane system. This suggests that Tm5NM1 is responsible for the correct localisation of T-tubules and possibly for the connectivity between the T-tubules and the SR.
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6.4.2 A role for cytoskeletal Tms in membrane dynamics associated with vesicle fusion
Reorganisation of membrane structures by actin filaments has been shown in migratory
cells, where actin is responsible for the membrane formations that allow cell motility
(Pollard and Borisy, 2003;Small and Resch, 2005). Tm isoforms have also been found
in association with membranes undergoing structural organisation such as the
localisation of Tm4 at the plasma membrane of podosomes and attachment structures of
osteoclast cells (McMichael et al., 2006). Tm5NM1 has also been shown to localise to
membranes undergoing structural changes, for example at the tip of the growing neural
processes (Schevzov et al., 2005a). It has been shown that the binding of insulin to its receptor in response to increased glucose levels not only triggers signalling cascades but also acts to remodel actin filaments (Tong et al., 2001). Insulin-dependent remodelling of the actin cytoskeleton leads to ruffle formation where the t-SNAREs, syntaxin4 and SNAP-23, concentrate in preparation for the docking of GLUT4- containing vesicles (Tong et al., 2001). It is possible that reorganization of the actin network situated beneath the plasma membrane reorganises membrane structures in order to allow GLUT4 vesicles to fuse with the plasma membrane after insulin- stimulation (Khayat et al., 2000). As we have shown that the absence or increased
expression of Tm5NM1 influences the glucose uptake pathway it is reasonable to say
that Tm5NM1 may influence actin dynamics involved with membrane remodelling in
preparation for vesicle fusion.
Membranes do not undergo spontaneous fusion. The reorganisation of both the vesicle
and the plasma membrane are required for fusion events to occur (Eitzen, 2003). Actin and its binding proteins have been shown to be heavily involved in these events and to facilitate fusion (Eitzen, 2003). It has been shown that myosins II and IC are required
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Chapter Six: General discussion for the reorganisation of membrane structures for GLUT4 vesicle fusion (Bose et al.,
2004;Steimle et al., 2005). These proteins mediate contraction of the actin cytoskeleton can also lead to localised remodeling of the cell cortex, including induction of membrane ruffles that may be required for vesicle fusion with the plasma membrane.
We propose that Tm5NM1 may recruit myosin II to actin filaments associated with the
T-tubule and plasma membranes in skeletal muscle, allowing this motor protein to facilitate membrane reorganisation events necessary for vesicle fusion. We hypothesise that the removal of Tm5NM1 from these actin filaments, as seen in the Tm5NM1 knockout mouse, decreases the efficiency of this recruitment, slowing down the rate of
GLUT4 vesicle delivery to the membrane. In the presence of excess Tm5NM1, as in the Tm5NM1 transgenic mouse, actin filaments are able to recruit more myosin II allowing more rapid fusion of GLUT4 vesicles.
6.5 A role for Tm5NM1 in the regulation of adipose tissue
Although the current study was focused primarily on deciphering the roles of cytoskeletal Tms in skeletal muscle we discovered a phenotype involving adipose tissue. Tm5NM1 over-expressing mice were found to have elevated levels of abdominal adipose tissue in comparison to wild-type controls. Tm5NM1 was also found to impact on the glucose uptake pathway which takes place primarily in both skeletal muscle and adipose tissue.
6.5.1 Tm5NM1 impacts on adipogenicity and PPAR-γ levels
Increased levels of adipose tissue have been found to be a major risk factor for the development of type II diabetes and a combination of these two metabolic dysfunctions is considered a major risk factor for cardiovascular disease (Spinler, 2006). It is due to
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this reason that research into obesity and type II diabetes has intensified over recent
years, with a focus on development of treatments for these conditions (Nehlin et al.,
2006). PPAR- agonists are one such treatment that has shown promise in the treatment of type II diabetes. These drugs have been shown to activate PPAR- , a nuclear receptor that has been shown to be a major regulator in the function of adipose cells
(Sharma and Staels, 2007). The mechanism of action of the PPAR- agonists has been difficult to understand as these drugs improve the disease status of patients with type II diabetes by an, as yet, unknown mechanism (Balakumar et al., 2007). In the Tm5NM1 over-expressing mice the levels of PPAR- were found to be elevated in skeletal muscle in comparison to wild-type controls. The mechanisms by which the increased levels of
Tm5NM1 are able to increase the levels of PPAR- are not known. It is possible that the impact of Tm5NM1 on glucose uptake may have downstream effects leading to increased PPAR- in response to increases in glucose uptake of the muscle. It may also be that the underlying cause of increased adipose tissue is the elevated levels of PPAR-
, a known factor in increased adipocyte proliferation and differentiation (Balakumar et al., 2007).
6.5.2 A possible role for Tm5NM1 for the treatment of obesity
Obesity is a risk factor for not only type II diabetes, but cardiovascular disease, chronic kidney disease and non-alcoholic fatty liver disease to name a few (Spinler,
2006;Kramer and Luke, 2007;Neuschwander-Tetri, 2007). The search for drugs that can be used to reduce obesity has been the focus of research as a decrease in the incidence of obesity should lead to a reduction in the prevalence of the diseases listed above. The current study shows that an increase in Tm5NM1 leads to an increase in abdominal adipose tissue by an unknown mechanism. It is then possible that a decrease
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Chapter Six: General discussion
in the levels of Tm5NM1 may lead to a reduction in levels of adipose tissue, making
Tm5NM1 a possible target for drug development. The levels of fat tissue in the
Tm5NM1 knockout mouse are yet to be studied; however, it would be predicted that
adipose tissue would be reduced in these mice. However, we have also proposed a role
for Tm5NM1 in glucose uptake and reduction of this Tm isoform has been associated
with impairment in glucose uptake in adipose tissue.
The use of PPAR- agonists in the treatment of diabetes has shown to also cause weight
gain in treated patients. The use of these drugs to treat diabetes has been debated in the
literature as the mechanism of treatment is unknown (Sharma and Staels, 2007;Barak
and Kim, 2007;Balakumar et al., 2007). Studies have shown that this weight gain may
not be detrimental to the health of the patient as adipocytes, while in abundance, seem
to operate more efficiently. PPAR- is involved in adipocyte proliferation and
differentiation and its agonists improve insulin sensitivity by promoting fatty acid
storage and inhibiting adipokine synthesis (Balakumar et al., 2007). Due to the increased levels of PPAR- found in Tm5NM1 transgenic mice further investigation into the relationship between PPAR- and Tm5NM1 is warranted before any conclusions can be drawn about the role of Tm5NM1 in obesity.
6.6 Future directions and developments
The research reported in this study presents data that implicates cytoskeletal Tms in the maintenance of muscle structure along with roles in physiological processes in muscle.
These results warrant further investigation into the various pathways that may be affected by the cytoskeletal Tms.
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6.6.1 Generation of Tm4-null mice
To study the specific roles of Tm4 in organisation of SR membranes and contractile
apparatus and regeneration and repair of skeletal muscle fibres a Tm4 knockout mouse
is required. Tm4 is also found to be highly expressed in a number of organs including
the lungs, spleen and kidneys with expression also found in cardiac muscle and neural
tissue (Schevzov et al., 2005b). It is advisable that a conditional knockout be produced
to specifically remove Tm4 from skeletal muscle as it is predicted that a global ablation
of Tm4 may be embryonically lethal.
It is hypothesised that Tm4 is involved in the organisation of the SR. To investigate
this, the levels of calsequestrin and other proteins located in the terminal SR (e.g.,
triadin, junctin and RyR) should be examined in Tm4 knockout mice by western blot
analysis. The ultrastructure of the skeletal muscle of these mice will be examined to
identify specific changes in this membrane system. The NMJ and MTJ regions of Tm4
knockout skeletal muscle will also be examined to analyse the membranes at these
regions. It could be predicted that these membrane structures may be more fragile and susceptible to damage in Tm4 knockout skeletal muscle due to the specific localisation of Tm4 to these regions.
It is expected that a Tm4 knockout mouse will have problems with muscle regeneration and repair. The induction of regeneration in these mice should provide information about the mechanism of Tm4 action in regeneration. The induction of degeneration/regeneration by Notexin injection should be used to asses the regenerative capacity of skeletal muscle from Tm4 knockout mice. We predict that skeletal muscle
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Chapter Six: General discussion from the Tm4 knockout mice will not be able to regenerate as quickly as wild-type muscle. During the process of regeneration the structure of the developing SR should be examined to elucidate the function of Tm4 on the positioning of these membranes.
As Tm4 is associated with SR in mature muscle, we hypothesise that these membranes may not be positioned correctly following muscle regeneration. Due to the high expression of Tm4 in satellite cells, these cells should also be examined specifically.
Isolation of satellite cells from regenerating Tm4 knockout muscle and analysis in culture may provide information about the capability of these cells to migrate efficiently.
Compensation from other cytoskeletal Tms will also need to be examined. It could be predicted that other cytoskeletal Tms may not compensate fully for the loss of Tm4.
6.6.2 Further investigation into the role of Tm5NM1 at the T-tubules
The specific localisation of Tm5NM1 at the T-tubules implies a role for this Tm in the structure of these membranes. Further investigation into the role of Tm5NM1 at the T- tubules may include analysis of the ion channels at the triad. Tm5NM1 may have a role in positioning ion channels at the triad. Analysis of a variety of ion channels in the
Tm5NM1 transgenic and knockout mice may reveal subtle differences in ion channels such as alterations in the localisation of the DHPR, RyRs and terminal SR localised
SERCA. Further analysis is also required into the morphological differences seen between Tm5NM1 knockout and wild-type mice, analysing multiple muscles from these mice using EM analysis.
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6.6.3 Further analysis into the role of Tm5NM1 in the translocation of GLUT4
The current study shows that altered regulation of Tm5NM1 impacts on the glucose uptake pathway. Further analysis is required to elucidate the specific role of this Tm in this pathway. Firstly the glucose tolerance testing performed on Tm5NM1 over- expressing mice should be performed in Tm5NM1-null mice. We would expect that the results obtained in the knockout mice would contrast with those found in the transgenic mice and that these mice would show a decrease in the rate of glucose uptake from the blood stream. Similarly, the glucose uptake analysis performed on adipose explants from knockout mice should be repeated using adipose tissue from transgenic mice. We predict that the adipose tissue from transgenic mice will take up glucose more rapidly than wild-type controls. These studies would demonstrate that loss and gain of function
of Tm5NM1 produce results in direct contrast to one another. Insulin tolerance should
also be analysed to determine if the effects seen are due to an increase in the insulin
sensitivity of the mice.
Further investigation into the role of Tm5NM1 in the translocation of GLUT4 should
be performed in adipocytes. Analysis of isolated adipocytes from Tm5NM1 null and
over-expressing mice will provide insight into the pathways of GLUT4 translocation.
GLUT4 vesicle trafficking can be analysed by introducing a GFP-tagged version of
GLUT4 into these cells. Visualisation of this protein in live cell imaging would enable
us to track vesicle trafficking following stimulation by insulin. Comparing the cells
derived from the different mouse strains in this analysis would provide a detailed
description of the role of the Tm5NM1 protein in each step of the GLUT4 translocation
pathway.
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Chapter Six: General discussion
A number of molecules have been identified in the various steps of the glucose uptake
pathway. We hypothesise that the Tm-defined actin cytoskeletal regulates vesicle translocation to the membrane and then facilitates vesicle fusion. The SNARE proteins,
found on both the vesicle and plasma membranes, may facilitate the interactions
between vesicles and the membrane (Bryant et al., 2002;Kanzaki, 2006a). SNARE proteins include VAMP-2 on the vesicle membrane and syntaxin-4 and SNAP-23
(Kanzaki, 2006a). It is possible that Tm5NM1-defined actin filaments interact with these proteins to facilitate vesicle movement and fusion. In order to determine if these
Tm5NM1 interacts with these proteins further analysis must be performed. Studies on these interactions should include looking for co-localisation of these molecules in wild- type skeletal muscle cells and adipocytes and analysis of the levels and localisation of these proteins in Tm5NM1 null and over-expressing tissue. If Tm5NM1 interacts or forms a complex with these proteins their abundance and/or localisation may be altered in response to increased/decreased levels of Tm5NM1 as seen in the transgenic and knockout mice
6.6.4 Analysis of cytoskeletal Tms in the process of adipodicity
The Tm5NM1 transgenic mouse was found to have elevated levels of abdominal adipose tissue (Section 5.3.4). In order to understand the impact of Tm5NM1 on adipose tissue further investigation is required into the role of Tm5NM1 in adipocytes.
Experimentation could include isolation of adipocytes from adipose tissue from wild- type mice as well as those with increased and decreased levels of Tm5NM1 protein.
Differentiation of these cells should be studied to examine the role of Tm5NM1 in
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Chapter Six: General discussion
adipogenesis. It would also be interesting to analyse the proliferative capacity of these cells, to determine if Tm5NM1 increases the ability of adipocytes to propagate.
Investigation into the mechanisms by which Tm5NM1 leads to an increase in PPAR-
levels in skeletal muscle is necessary. As PPAR- plays a number of roles in adipose
tissue this analysis could prove difficult. The up-regulation of this molecule in skeletal
muscle of Tm5NM1 transgenic mice may be due to increased levels of intermuscular
fat deposits. Analysis of skeletal muscle from Tm5NM1 transgenic mice at the EM
level should be completed, to examine for the presence of lipid accumulation. If lipid
deposits are found in skeletal muscle, this would suggest that the elevated levels of
PPAR- in skeletal muscle could be due to the presence of PPAR- in lipid droplets
found within skeletal muscle, rather than in the muscle cells themselves.
6.7 Concluding remarks
We have identified two cytoskeletal Tms that are present in skeletal muscle and
compartmentalise to form functionally distinct filaments. One filament system, defined
by Tm4, is present in muscle regeneration and in response to disease or injury. We
suggest that Tm4-defined filaments play a role in the process of regeneration and
repair. We also hypothesise that mutations in the Tm4 gene, δ-Tm, may cause a disease pathology, resulting from problems in the regenerative process in response to injury and muscle membrane damage. This finding is significant as a number of myopathies have unknown genetic causes, identification of a new mutation increases the likelihood of successful treatment by methods such as gene therapy.
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Chapter Six: General discussion
The Tm5NM1-defined cytoskeleton has been shown to be involved in the morphology
of the T-tubule and sarcolemmal membranes and as well as plays a role in glucose
uptake. This finding is important as it implicates this cytoskeleton in the regulation of
GLUT vesicle trafficking which may lead to a role in type II diabetes. As the incidence of this lifestyle disease is on the increase research has intensified to identify drug targets for treatment of type II diabetes and obesity. The identification of Tm5NM1 in these processes and the link to PPAR- may provide a new target for anti-diabetes
drugs.
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Reference List
Aarimaa,V., Rantanen,J., Best,T., Schultz,E., Corr,D., Kalimo,H. (2004). Mild eccentric stretch injury in skeletal muscle causes transient effects on tensile load and cell proliferation. Scand J Med Sci Sports. 14, 367-372.
Abate,N., Garg,A., Peshock,R.M., Stray-Gundersen,J., Grundy,S.M. (1995). Relationships of generalized and regional adiposity to insulin sensitivity in men. J Clin Invest. 96, 88-98.
Agarkova,I., Ehler,E., Lange,S., Schoenauer,R., Perriard,J.C. (2003). M-band: a safeguard for sarcomere stability? J Muscle Res Cell Motil 24, 191-203.
Ai,H., Ralston,E., Lauritzen,H.P., Galbo,H., Ploug,T. (2003). Disruption of microtubules in rat skeletal muscle does not inhibit insulin- or contraction-stimulated glucose transport. Am J Physiol Endocrinol Metab. 285, E836-E844.
Alberts,B., Bray,D., Johnson,A., Lewis,J., Raff,M., Roberts,K., Walter,P. (1998). Essential Cell Biology. New York: Garland Publishing.
An,X., Salomao,M., Guo,X., Gratzer,W., Mohandas,N. (2007). Tropomyosin modulates erythrocyte membrane stability. Blood. 109, 1284-1288.
Askanas,V., Bornemann,A., Engel,W.K. (1990). Immunocytochemical localization of desmin at human neuromuscular junctions. Neurology. 40, 949-953.
Atilgan,E., Wirtz,D., Sun,S.X. (2005). Morphology of the lamellipodium and organization of actin filaments at the leading edge of crawling cells. Biophys J. 89, 3589-3602.
Bailly,M., Macaluso,F., Cammer,M., Chan,A., Segall,J.E., Condeelis,J.S. (1999). Relationship between Arp2/3 complex and the barbed ends of actin filaments at the leading edge of carcinoma cells after epidermal growth factor stimulation. J Cell Biol. %19;145, 331-345.
Balakumar,P., Rose,M., Ganti,S.S., Krishan,P., Singh,M. (2007). PPAR dual agonists: Are they opening Pandora's Box? Pharmacol Res. ..
Bamburg,J.R., McGough,A., Ono,S. (1999). Putting a new twist on actin: ADF/cofilins modulate actin dynamics. Trends Cell Biol. 9, 364-370.
Bansal,D., Campbell,K.P. (2004). Dysferlin and the plasma membrane repair in muscular dystrophy. Trends Cell Biol. 14, 206-213.
Bansal,D., Miyake,K., Vogel,S.S., Groh,S., Chen,C.C., Williamson,R., McNeil,P.L., Campbell,K.P. (2003). Defective membrane repair in dysferlin-deficient muscular dystrophy. Nature 423, 168-172.
Barak,Y., Kim,S. (2007). Genetic Manipulations of PPARs: Effects on Obesity and Metabolic Disease. PPAR.Res. 2007:12781., 12781.
Page 198
Becker,B.E., Cassimeris,L. (2005). Cytoskeleton: microtubules born on the run. Curr Biol. 15, R551-R554.
Bellin,R.M., Huiatt,T.W., Critchley,D.R., Robson,R.M. (2001). Synemin may function to directly link muscle cell intermediate filaments to both myofibrillar Z-lines and costameres. J Biol Chem 276, 32330-32337.
Berg,J.S., Powell,B.C., Cheney,R.E. (2001). A millennial myosin census. Mol Biol Cell. 12, 780-794.
Bernstein,B.W., Bamburg,J.R. (1982). Tropomyosin binding to F-actin protects the F- actin from disassembly by brain actin-depolymerizing factor (ADF). Cell Motil. 2, 1-8.
Berthier,C., Blaineau,S. (1997). Supramolecular organization of the subsarcolemmal cytoskeleton of adult skeletal muscle fibers. A review. Biol Cell 89, 413-434.
Bischoff,R. (1994). The Satellite Cell and Muscle Regeneration. In: Myology, ed. Engel Andrew G, C.Franzini-ArmstrongNew York: MrGraw Hill, 97-118.
Blake,D.J., Weir,A., Newey,S.E., Davies,K.E. (2002). Function and genetics of dystrophin and dystrophin-related proteins in muscle. Physiol Rev. 82, 291-329.
Blanchard,E.M., Iizuka,K., Christe,M., Conner,D.A., Geisterfer-Lowrance,A., Schoen,F.J., Maughan,D.W., Seidman,C.E., Seidman,J.G. (1997). Targeted ablation of the murine alpha-tropomyosin gene. Circ Res. 81, 1005-1010.
Blau,H.M., Webster,C., Pavlath,G.K. (1983). Defective myoblasts identified in Duchenne muscular dystrophy. Proc Natl Acad Sci U S A. 80, 4856-4860.
Bloch,R.J., Capetanaki,Y., O'Neill,A., Reed,P., Williams,M.W., Resneck,W.G., Porter,N.C., Ursitti,J.A. (2002). Costameres: repeating structures at the sarcolemma of skeletal muscle. Clin Orthop S203-S210.
Bloch,R.J., Hall,Z.W. (1983). Cytoskeletal components of the vertebrate neuromuscular junction: vinculin, alpha-actinin, and filamin. J Cell Biol. 97, 217-223.
Booth,F.W. (1977). Time course of muscular atrophy during immobilization of hindlimbs in rats. J Appl Physiol. 43, 656-661.
Bose,A., Guilherme,A., Robida,S.I., Nicoloro,S.M., Zhou,Q.L., Jiang,Z.Y., Pomerleau,D.P., Czech,M.P. (2002). Glucose transporter recycling in response to insulin is facilitated by myosin Myo1c. Nature. %19-26;420, 821-824.
Bose,A., Robida,S., Furcinitti,P.S., Chawla,A., Fogarty,K., Corvera,S., Czech,M.P. (2004). Unconventional myosin Myo1c promotes membrane fusion in a regulated exocytic pathway. Mol Cell Biol. 24, 5447-5458.
Bottinelli,R., Pellegrino,M.A., Canepari,M., Rossi,R., Reggiani,C. (1999). Specific contributions of various muscle fibre types to human muscle performance: an in vitro study. J Electromyogr.Kinesiol. 9, 87-95.
Page 199
Brown,P.O., Botstein,D. (1999). Exploring the new world of the genome with DNA microarrays. Nat Genet. 21, 33-37.
Bryant,N.J., Govers,R., James,D.E. (2002). Regulated transport of the glucose transporter GLUT4. Nat Rev Mol Cell Biol. 3, 267-277.
Bryce,N.S., Schevzov,G., Ferguson,V., Percival,J.M., Lin,J.J., Matsumura,F., Bamburg,J.R., Jeffrey,P.L., Hardeman,E.C., Gunning,P., Weinberger,R.P. (2003). Specification of actin filament function and molecular composition by tropomyosin isoforms. Mol Biol Cell 14, 1002-1016.
Buckingham,M., Bajard,L., Chang,T., Daubas,P., Hadchouel,J., Meilhac,S., Montarras,D., Rocancourt,D., Relaix,F. (2003). The formation of skeletal muscle: from somite to limb. J Anat 202, 59-68.
Burgunder,J.M., Hunziker,W. (2003). Hereditary spastic paraplegia: clues from a rare disorder for a common problem? IUBMB Life. 55, 347-352.
Capell,B.C., Collins,F.S. (2006). Human laminopathies: nuclei gone genetically awry. Nat Rev Genet. 7, 940-952.
Capetanaki,Y. (2002). Desmin cytoskeleton: a potential regulator of muscle mitochondrial behavior and function. Trends Cardiovasc Med. 12, 339-348.
Capetanaki,Y., Milner,D.J. (1998). Desmin cytoskeleton in muscle integrity and function. Subcell.Biochem. 31, 463-495.
Capozza,F., Combs,T.P., Cohen,A.W., Cho,Y.R., Park,S.Y., Schubert,W., Williams,T.M., Brasaemle,D.L., Jelicks,L.A., Scherer,P.E., Kim,J.K., Lisanti,M.P. (2005). Caveolin-3 knockout mice show increased adiposity and whole body insulin resistance, with ligand-induced insulin receptor instability in skeletal muscle. Am J Physiol Cell Physiol. 288, C1317-C1331.
Carlier,M.F., Pantaloni,D. (1997). Control of actin dynamics in cell motility. J Mol Biol. %20;269, 459-467.
Carlson,B.M., Faulkner,J.A. (1983). The regeneration of skeletal muscle fibers following injury: a review. Med Sci Sports Exerc. 15, 187-198.
Carlsson,L., Li,Z.L., Paulin,D., Price,M.G., Breckler,J., Robson,R.M., Wiche,G., Thornell,L.E. (2000). Differences in the distribution of synemin, paranemin, and plectin in skeletal muscles of wild-type and desmin knock-out mice. Histochem Cell Biol. 114, 39-47.
Chang,L., Chiang,S.H., Saltiel,A.R. (2004). Insulin signaling and the regulation of glucose transport. Mol Med. 10, 65-71.
Charge,S.B., Rudnicki,M.A. (2004). Cellular and molecular regulation of muscle regeneration. Physiol Rev. 84, 209-238.
Page 200
Chen,Y., Wang,S.M., Wu,J.C., Huang,S.H. (2006). Effects of PPARgamma agonists on cell survival and focal adhesions in a Chinese thyroid carcinoma cell line. J Cell Biochem. 98, 1021-1035.
Cherepanova,O., Orlova,A., Galkin,V.E., van der Ven,P.F., Furst,D.O., Jin,J.P., Egelman,E.H. (2006). Xin-repeats and nebulin-like repeats bind to F-actin in a similar manner. J Mol Biol. 356, 714-723.
Chevalier-Larsen,E., Holzbaur,E.L. (2006). Axonal transport and neurodegenerative disease. Biochim Biophys Acta. 1762, 1094-1108.
Clark,K.A., McElhinny,A.S., Beckerle,M.C., Gregorio,C.C. (2002). Striated muscle cytoarchitecture: an intricate web of form and function. Annu Rev Cell Dev Biol 18, 637-706.
Clarkson,E., Costa,C.F., Machesky,L.M. (2004). Congenital myopathies: diseases of the actin cytoskeleton. J Pathol. 204, 407-417.
Corbett,M.A., Robinson,C.S., Dunglison,G.F., Yang,N., Joya,J.E., Stewart,A.W., Schnell,C., Gunning,P.W., North,K.N., Hardeman,E.C. (2001). A mutation in alpha- tropomyosin(slow) affects muscle strength, maturation and hypertrophy in a mouse model for nemaline myopathy. Hum Mol Genet 10, 317-328.
Corcoran,M.P., Lamon-Fava,S., Fielding,R.A. (2007). Skeletal muscle lipid deposition and insulin resistance: effect of dietary fatty acids and exercise. Am J Clin Nutr. 85, 662-677.
Coulombe,P.A., Wong,P. (2004). Cytoplasmic intermediate filaments revealed as dynamic and multipurpose scaffolds. Nat Cell Biol. 6, 699-706.
Coviello,D.A., Maron,B.J., Spirito,P., Watkins,H., Vosberg,H.P., Thierfelder,L., Schoen,F.J., Seidman,J.G., Seidman,C.E. (1997). Clinical features of hypertrophic cardiomyopathy caused by mutation of a "hot spot" in the alpha-tropomyosin gene. J Am Coll.Cardiol. 29, 635-640.
Craig,R. (1994). The Structure of the Contractile Filaments. In: Myology, ed. Engel Andrew G, C.Franzini-ArmstrongNew York: McGraw Hill, 134-175.
Crawford,K., Flick,R., Close,L., Shelly,D., Paul,R., Bove,K., Kumar,A., Lessard,J. (2002). Mice lacking skeletal muscle actin show reduced muscle strength and growth deficits and die during the neonatal period. Mol Cell Biol. 22, 5887-5896.
D'Andrea-Merrins,M., Chang,L., Lam,A.D., Ernst,S.A., Stuenkel,E.L. (2007). Munc18c interaction with syntaxin4 monomers and SNARE complex intermediates in GLUT4 vesicle trafficking. J Biol Chem. ..
Dai,Z., Luo,X., Xie,H., Peng,H.B. (2000). The actin-driven movement and formation of acetylcholine receptor clusters. J Cell Biol 150, 1321-1334.
Dalby-Payne,J.R., O'Loughlin,E.V., Gunning,P. (2003). Polarization of specific tropomyosin isoforms in gastrointestinal epithelial cells and their impact on CFTR at the apical surface. Mol Biol Cell 14, 4365-4375.
Page 201
Dalkilic,I., Kunkel,L.M. (2003). Muscular dystrophies: genes to pathogenesis. Curr Opin Genet Dev 13, 231-238.
Dantzig,J.A., Liu,T.Y., Goldman,Y.E. (2006). Functional studies of individual Myosin molecules. Ann N Y Acad Sci. 1080:1-18., 1-18.
Das,K.M., Dasgupta,A., Mandal,A., Geng,X. (1993). Autoimmunity to cytoskeletal protein tropomyosin. A clue to the pathogenetic mechanism for ulcerative colitis. J Immunol. 150, 2487-2493.
Davies,K.E., Nowak,K.J. (2006). Molecular mechanisms of muscular dystrophies: old and new players. Nat Rev Mol Cell Biol. 7, 762-773.
Decostre,V., Ben Yaou,R., Bonne,G. (2005). Laminopathies affecting skeletal and cardiac muscles: clinical and pathophysiological aspects. Acta Myol. 24, 104-109.
Dedieu,S., Poussard,S., Mazeres,G., Grise,F., Dargelos,E., Cottin,P., Brustis,J.J. (2004). Myoblast migration is regulated by calpain through its involvement in cell attachment and cytoskeletal organization. Exp Cell Res. 292, 187-200.
Dellavalle,A., Sampaolesi,M., Tonlorenzi,R., Tagliafico,E., Sacchetti,B., Perani,L., Innocenzi,A., Galvez,B.G., Messina,G., Morosetti,R., Li,S., Belicchi,M., Peretti,G., Chamberlain,J.S., Wright,W.E., Torrente,Y., Ferrari,S., Bianco,P., Cossu,G. (2007). Pericytes of human skeletal muscle are myogenic precursors distinct from satellite cells. Nat Cell Biol. 9, 255-267.
Dent,E.W., Gertler,F.B. (2003). Cytoskeletal dynamics and transport in growth cone motility and axon guidance. Neuron. 40, 209-227.
Denti,M.A., Rosa,A., D'Antona,G., Sthandier,O., De Angelis,F.G., Nicoletti,C., Allocca,M., Pansarasa,O., Parente,V., Musaro,A., Auricchio,A., Bottinelli,R., Bozzoni,I. (2006). Body-wide gene therapy of Duchenne muscular dystrophy in the mdx mouse model. Proc Natl Acad Sci U S A. 103, 3758-3763.
DePina,A.S., Langford,G.M. (1999). Vesicle transport: the role of actin filaments and myosin motors. Microsc Res Tech. 47, 93-106.
DesMarais,V., Ichetovkin,I., Condeelis,J., Hitchcock-DeGregori,S.E. (2002). Spatial regulation of actin dynamics: a tropomyosin-free, actin-rich compartment at the leading edge. J Cell Sci 115, 4649-4660.
Dezawa,M., Ishikawa,H., Itokazu,Y., Yoshihara,T., Hoshino,M., Takeda,S., Ide,C., Nabeshima,Y. (2005). Bone marrow stromal cells generate muscle cells and repair muscle degeneration. Science. 309, 314-317.
Dickson,G., Azad,A., Morris,G.E., Simon,H., Noursadeghi,M., Walsh,F.S. (1992). Co- localization and molecular association of dystrophin with laminin at the surface of mouse and human myotubes. J Cell Sci. 103, 1223-1233.
DiMario,J.X., Uzman,A., Strohman,R.C. (1991). Fiber regeneration is not persistent in dystrophic (MDX) mouse skeletal muscle. Dev Biol. 148, 314-321.
Page 202
Dlugosz,A.A., Antin,P.B., Nachmias,V.T., Holtzer,H. (1984). The relationship between stress fiber-like structures and nascent myofibrils in cultured cardiac myocytes. J Cell Biol. 99, 2268-2278.
Donner,K., Ollikainen,M., Ridanpaa,M., Christen,H.J., Goebel,H.H., de Visser,M., Pelin,K., Wallgren-Pettersson,C. (2002). Mutations in the beta-tropomyosin (TPM2) gene--a rare cause of nemaline myopathy. Neuromuscul.Disord 12, 151-158.
Drees,B., Brown,C., Barrell,B.G., Bretscher,A. (1995). Tropomyosin is essential in yeast, yet the TPM1 and TPM2 products perform distinct functions. J Cell Biol 128, 383-392.
Dugani,C.B., Klip,A. (2005). Glucose transporter 4: cycling, compartments and controversies. EMBO Rep. 6, 1137-1142.
Durbeej,M., Campbell,K.P. (2002). Muscular dystrophies involving the dystrophin- glycoprotein complex: an overview of current mouse models. Curr Opin Genet Dev 12, 349-361.
Eckes,B., Dogic,D., Colucci-Guyon,E., Wang,N., Maniotis,A., Ingber,D., Merckling,A., Langa,F., Aumailley,M., Delouvee,A., Koteliansky,V., Babinet,C., Krieg,T. (1998). Impaired mechanical stability, migration and contractile capacity in vimentin-deficient fibroblasts. J Cell Sci. 111, 1897-1907.
Ehrhardt,J., Morgan,J. (2005). Regenerative capacity of skeletal muscle. Curr Opin Neurol. 18, 548-553.
Eitzen,G. (2003). Actin remodeling to facilitate membrane fusion. Biochim Biophys Acta. 1641, 175-181.
El Amraoui,A., Petit,C. (2005). Usher I syndrome: unravelling the mechanisms that underlie the cohesion of the growing hair bundle in inner ear sensory cells. J Cell Sci. 118, 4593-4603.
Engel,A.G., Yamamoto,M., Fischbeck,K.H. (1994). Dystrophinopathies. In: Myology, ed. Engel Andrew G, C.Franzini-ArmstrongNew York: McGraw-Hill INC, 1133-1187.
Ervasti,J.M. (2003). Costameres: the Achilles' heel of Herculean muscle. J Biol Chem 278, 13591-13594.
Ervasti,J.M., Campbell,K.P. (1993). A role for the dystrophin-glycoprotein complex as a transmembrane linker between laminin and actin. J Cell Biol. 122, 809-823.
Even-Ram,S., Doyle,A.D., Conti,M.A., Matsumoto,K., Adelstein,R.S., Yamada,K.M. (2007). Myosin IIA regulates cell motility and actomyosin-microtubule crosstalk. Nat Cell Biol. 9, 299-309.
Fanning,A.S., Wolenski,J.S., Mooseker,M.S., Izant,J.G. (1994). Differential regulation of skeletal muscle myosin-II and brush border myosin-I enzymology and mechanochemistry by bacterially produced tropomyosin isoforms. Cell Motil Cytoskeleton. 29, 29-45.
Page 203
Fatima,S., Yaghini,F.A., Pavicevic,Z., Kalyankrishna,S., Jafari,N., Luong,E., Estes,A., Malik,K.U. (2005). Intact actin filaments are required for cytosolic phospholipase A2 translocation but not for its activation by norepinephrine in vascular smooth muscle cells. J Pharmacol Exp Ther. 313, 1017-1026.
Faulkner,G., Lanfranchi,G., Valle,G. (2001). Telethonin and other new proteins of the Z-disc of skeletal muscle. IUBMB Life. 51, 275-282.
Fischer,R.S., Fowler,V.M. (2003). Tropomodulins: life at the slow end. Trends Cell Biol. 13, 593-601.
Fletcher,L.M., Welsh,G.I., Oatey,P.B., Tavare,J.M. (2000). Role for the microtubule cytoskeleton in GLUT4 vesicle trafficking and in the regulation of insulin-stimulated glucose uptake. Biochem J. 352 Pt 2:267-76., 267-276.
Flucher,B.E., Andrews,S.B., Daniels,M.P. (1994). Molecular organization of transverse tubule/sarcoplasmic reticulum junctions during development of excitation-contraction coupling in skeletal muscle. Mol Biol Cell. 5, 1105-1118.
Flucher,B.E., Phillips,J.L., Powell,J.A., Andrews,S.B., Daniels,M.P. (1992). Coordinated development of myofibrils, sarcoplasmic reticulum and transverse tubules in normal and dysgenic mouse skeletal muscle, in vivo and in vitro. Dev Biol. 150, 266- 280.
Flucher,B.E., Takekura,H., Franzini-Armstrong,C. (1993). Development of the excitation-contraction coupling apparatus in skeletal muscle: association of sarcoplasmic reticulum and transverse tubules with myofibrils. Dev Biol. 160, 135-147.
Foster,K., Foster,H., Dickson,J.G. (2006). Gene therapy progress and prospects: Duchenne muscular dystrophy. Gene Ther. 13, 1677-1685.
Foster,L.J., Li,D., Randhawa,V.K., Klip,A. (2001). Insulin accelerates inter-endosomal GLUT4 traffic via phosphatidylinositol 3-kinase and protein kinase B. J Biol Chem. 276, 44212-44221.
Fowler,V.M., Sussmann,M.A., Miller,P.G., Flucher,B.E., Daniels,M.P. (1993). Tropomodulin is associated with the free (pointed) ends of the thin filaments in rat skeletal muscle. J Cell Biol. 120, 411-420.
Frank,D., Kuhn,C., Katus,H.A., Frey,N. (2006). The sarcomeric Z-disc: a nodal point in signalling and disease. J Mol Med. 84, 446-468.
Franzen,B., Linder,S., Uryu,K., Alaiya,A.A., Hirano,T., Kato,H., Auer,G. (1996). Expression of tropomyosin isoforms in benign and malignant human breast lesions. Br J Cancer. 73, 909-913.
Franzini-Armstrong,C. (1991). Simultaneous maturation of transverse tubules and sarcoplasmic reticulum during muscle differentiation in the mouse. Dev Biol. 146, 353- 363.
Page 204
Franzini-Armstrong,C. (1994). The Sarcoplasmic Reticulum and the Transverse Tubules. In: Myology, ed. Engel Andrew G, C.Franzini-ArmstrongNew York: McGraw Hill, 176-199.
Franzini-Armstrong,C., Fischman,D.A. (1994). Morphogenesis of Skeletal Muscle Fibers. In: Myology, ed. Engel Andrew G, C.Franzini-ArmstrongNew York: McGraw Hill, 74-96.
Fuchs,E., Cleveland,D.W. (1998). A structural scaffolding of intermediate filaments in health and disease. Science. 279, 514-519.
Fuchs,E., Weber,K. (1994). Intermediate filaments: structure, dynamics, function, and disease. Annu Rev Biochem. 63:345-82., 345-382.
Fujioka,S., Matsuzawa,Y., Tokunaga,K., Tarui,S. (1987). Contribution of intra- abdominal fat accumulation to the impairment of glucose and lipid metabolism in human obesity. Metabolism. 36, 54-59.
Fujisawa,T., Kostyukova,A., Maeda,Y. (2001). The shapes and sizes of two domains of tropomodulin, the P-end-capping protein of actin-tropomyosin. FEBS Lett. 498, 67-71.
Gadde,S., Heald,R. (2004). Mechanisms and molecules of the mitotic spindle. Curr Biol. 14, R797-R805.
Galbiati,F., Razani,B., Lisanti,M.P. (2001). Caveolae and caveolin-3 in muscular dystrophy. Trends Mol Med. 7, 435-441.
Ganeshan,R., Nowotarski,K., Di,A., Nelson,D.J., Kirk,K.L. (2007). CFTR surface expression and chloride currents are decreased by inhibitors of N-WASP and actin polymerization. Biochim Biophys Acta. 1773, 192-200.
Goebel,H.H., Muller,H.D. (2006). Protein aggregate myopathies. Semin Pediatr Neurol 13, 96-103.
Goldspink,G. (1999). Changes in muscle mass and phenotype and the expression of autocrine and systemic growth factors by muscle in response to stretch and overload. J Anat 194 ( Pt 3), 323-334.
Goldstein,L.S. (2003). Do disorders of movement cause movement disorders and dementia? Neuron. 40, 415-425.
Goley,E.D., Welch,M.D. (2006). The ARP2/3 complex: an actin nucleator comes of age. Nat Rev Mol Cell Biol. 7, 713-726.
Gordon,A.M., Homsher,E., Regnier,M. (2000). Regulation of contraction in striated muscle. Physiol Rev. 80, 853-924.
Grady,R.M., Merlie,J.P., Sanes,J.R. (1997a). Subtle neuromuscular defects in utrophin- deficient mice. J Cell Biol 136, 871-882.
Page 205
Grady,R.M., Teng,H., Nichol,M.C., Cunningham,J.C., Wilkinson,R.S., Sanes,J.R. (1997b). Skeletal and cardiac myopathies in mice lacking utrophin and dystrophin: a model for Duchenne muscular dystrophy. Cell. 90, 729-738.
Granger,B.L., Lazarides,E. (1978). The existence of an insoluble Z disc scaffold in chicken skeletal muscle. Cell. 15, 1253-1268.
Gregorevic,P., Allen,J.M., Minami,E., Blankinship,M.J., Haraguchi,M., Meuse,L., Finn,E., Adams,M.E., Froehner,S.C., Murry,C.E., Chamberlain,J.S. (2006). rAAV6- microdystrophin preserves muscle function and extends lifespan in severely dystrophic mice. Nat Med. 12, 787-789.
Gregorio,C.C., Trombitas,K., Centner,T., Kolmerer,B., Stier,G., Kunke,K., Suzuki,K., Obermayr,F., Herrmann,B., Granzier,H., Sorimachi,H., Labeit,S. (1998). The NH2 terminus of titin spans the Z-disc: its interaction with a novel 19-kD ligand (T-cap) is required for sarcomeric integrity. J Cell Biol. 143, 1013-1027.
Griffiths,G., McDowall,A., Back,R., Dubochet,J. (1984). On the preparation of cryosections for immunocytochemistry. J Ultrastruct.Res. 89, 65-78.
Gunning,P., Gordon,M., Wade,R., Gahlmann,R., Lin,C.S., Hardeman,E. (1990). Differential control of tropomyosin mRNA levels during myogenesis suggests the existence of an isoform competition-autoregulatory compensation control mechanism. Dev Biol. 138, 443-453.
Gunning,P., Hardeman,E., Jeffrey,P., Weinberger,R. (1998a). Creating intracellular structural domains: spatial segregation of actin and tropomyosin isoforms in neurons. Bioessays 20, 892-900.
Gunning, P., O'neill, G. M., and Hardeman, E. Tropomyosin-based regulation of the actin cytoskeleton in time and space. J Physiol . 2007. Ref Type: In Press
Gunning,P., Weinberger,R., Jeffrey,P., Hardeman,E. (1998b). Isoform sorting and the creation of intracellular compartments. Annu Rev Cell Dev Biol 14:339-72., 339-372.
Gunning,P.W., Schevzov,G., Kee,A.J., Hardeman,E.C. (2005). Tropomyosin isoforms: divining rods for actin cytoskeleton function. Trends Cell Biol. 15, 333-341.
Gupton,S.L., Anderson,K.L., Kole,T.P., Fischer,R.S., Ponti,A., Hitchcock- DeGregori,S.E., Danuser,G., Fowler,V.M., Wirtz,D., Hanein,D., Waterman- Storer,C.M. (2005). Cell migration without a lamellipodium: translation of actin dynamics into cell movement mediated by tropomyosin. J Cell Biol 168, 619-631.
Guzik,B.W., Goldstein,L.S. (2004). Microtubule-dependent transport in neurons: steps towards an understanding of regulation, function and dysfunction. Curr Opin Cell Biol. 16, 443-450.
Had,L., Faivre-Sarrailh,C., Legrand,C., Mery,J., Brugidou,J., Rabie,A. (1994). Tropomyosin isoforms in rat neurons: the different developmental profiles and distributions of TM-4 and TMBr-3 are consistent with different functions. J Cell Sci. 107, 2961-2973.
Page 206
Had,L., Faivre-Sarrailh,C., Legrand,C., Rabie,A. (1993). The expression of tropomyosin genes in pure cultures of rat neurons, astrocytes and oligodendrocytes is highly cell-type specific and strongly regulated during development. Brain Res Mol Brain Res. 18, 77-86.
Hall,Z.W., Sanes,J.R. (1993). Synaptic structure and development: the neuromuscular junction. Cell 72 Suppl:99-121., 99-121.
Hanft,L.M., Rybakova,I.N., Patel,J.R., Rafael-Fortney,J.A., Ervasti,J.M. (2006). Cytoplasmic gamma-actin contributes to a compensatory remodeling response in dystrophin-deficient muscle. Proc Natl Acad Sci U S A. 103, 5385-5390.
Hannan,A.J., Gunning,P., Jeffrey,P.L., Weinberger,R.P. (1998). Structural compartments within neurons: developmentally regulated organization of microfilament isoform mRNA and protein. Mol Cell Neurosci 11, 289-304.
Hannan,A.J., Schevzov,G., Gunning,P., Jeffrey,P.L., Weinberger,R.P. (1995). Intracellular localization of tropomyosin mRNA and protein is associated with development of neuronal polarity. Mol Cell Neurosci. 6, 397-412.
He,A., Liu,X., Liu,L., Chang,Y., Fang,F. (2007). How many signals impinge on GLUT4 activation by insulin? Cell Signal. 19, 1-7.
Head,B.P., Patel,H.H., Roth,D.M., Murray,F., Swaney,J.S., Niesman,I.R., Farquhar,M.G., Insel,P.A. (2006). Microtubules and actin microfilaments regulate lipid raft/caveolae localization of adenylyl cyclase signaling components. J Biol Chem. 281, 26391-26399.
Helfand,B.T., Chang,L., Goldman,R.D. (2004). Intermediate filaments are dynamic and motile elements of cellular architecture. J Cell Sci. 117, 133-141.
Hernandez-Deviez,D.J., Martin,S., Laval,S.H., Lo,H.P., Cooper,S.T., North,K.N., Bushby,K., Parton,R.G. (2006). Aberrant dysferlin trafficking in cells lacking caveolin or expressing dystrophy mutants of caveolin-3. Hum Mol Genet. 15, 129-142.
Hijikata,T., Murakami,T., Ishikawa,H., Yorifuji,H. (2003). Plectin tethers desmin intermediate filaments onto subsarcolemmal dense plaques containing dystrophin and vinculin. Histochem Cell Biol. 119, 109-123.
Hill,M., Wernig,A., Goldspink,G. (2003). Muscle satellite (stem) cell activation during local tissue injury and repair. J Anat. 203, 89-99.
Hirokawa,N., Takemura,R. (2003). Biochemical and molecular characterization of diseases linked to motor proteins. Trends Biochem Sci. 28, 558-565.
Hook,J., Lemckert,F., Qin,H., Schevzov,G., Gunning,P. (2004). Gamma tropomyosin gene products are required for embryonic development. Mol Cell Biol 24, 2318-2323.
Hornef,N., Olbrich,H., Horvath,J., Zariwala,M.A., Fliegauf,M., Loges,N.T., Wildhaber,J., Noone,P.G., Kennedy,M., Antonarakis,S.E., Blouin,J.L., Bartoloni,L., Nusslein,T., Ahrens,P., Griese,M., Kuhl,H., Sudbrak,R., Knowles,M.R., Reinhardt,R.,
Page 207
Omran,H. (2006). DNAH5 mutations are a common cause of primary ciliary dyskinesia with outer dynein arm defects. Am J Respir Crit Care Med. 174, 120-126.
Horowits,R., Kempner,E.S., Bisher,M.E., Podolsky,R.J. (1986). A physiological role for titin and nebulin in skeletal muscle. Nature. 323, 160-164.
Hotulainen,P., Lappalainen,P. (2006). Stress fibers are generated by two distinct actin assembly mechanisms in motile cells. J Cell Biol. 173, 383-394.
Howard,J., Hyman,A.A. (2003). Dynamics and mechanics of the microtubule plus end. Nature. 422, 753-758.
Howell,G.J., Holloway,Z.G., Cobbold,C., Monaco,A.P., Ponnambalam,S. (2006). Cell biology of membrane trafficking in human disease. Int Rev Cytol. 252:1-69., 1-69.
Huang,J., Imamura,T., Babendure,J.L., Lu,J.C., Olefsky,J.M. (2005). Disruption of microtubules ablates the specificity of insulin signaling to GLUT4 translocation in 3T3- L1 adipocytes. J Biol Chem. 280, 42300-42306.
Hughes,B.W., Kusner,L.L., Kaminski,H.J. (2006). Molecular architecture of the neuromuscular junction. Muscle Nerve. 33, 445-461.
Hutchinson,D.O., Charlton,A., Laing,N.G., Ilkovski,B., North,K.N. (2006). Autosomal dominant nemaline myopathy with intranuclear rods due to mutation of the skeletal muscle ACTA1 gene: clinical and pathological variability within a kindred. Neuromuscul.Disord 16, 113-121.
Ilkovski,B., Cooper,S.T., Nowak,K., Ryan,M.M., Yang,N., Schnell,C., Durling,H.J., Roddick,L.G., Wilkinson,I., Kornberg,A.J., Collins,K.J., Wallace,G., Gunning,P., Hardeman,E.C., Laing,N.G., North,K.N. (2001). Nemaline myopathy caused by mutations in the muscle alpha-skeletal-actin gene. Am J Hum Genet. 68, 1333-1343.
Ishikawa,R., Yamashiro,S., Matsumura,F. (1989a). Annealing of gelsolin-severed actin fragments by tropomyosin in the presence of Ca2+. Potentiation of the annealing process by caldesmon. J Biol Chem. 264, 16764-16770.
Ishikawa,R., Yamashiro,S., Matsumura,F. (1989b). Differential modulation of actin- severing activity of gelsolin by multiple isoforms of cultured rat cell tropomyosin. Potentiation of protective ability of tropomyosins by 83-kDa nonmuscle caldesmon. J Biol Chem. 264, 7490-7497.
Ishiki,M., Klip,A. (2005). Minireview: recent developments in the regulation of glucose transporter-4 traffic: new signals, locations, and partners. Endocrinology. 146, 5071- 5078.
James,D.E. (2005). MUNC-ing around with insulin action. J Clin Invest. 115, 219-221.
James,D.E., Strube,M., Mueckler,M. (1989). Molecular cloning and characterization of an insulin-regulatable glucose transporter. Nature. 338, 83-87.
Janson,M.E., de Dood,M.E., Dogterom,M. (2003). Dynamic instability of microtubules is regulated by force. J Cell Biol. 161, 1029-1034.
Page 208
Jarvinen,T.A., Jarvinen,T.L., Kaariainen,M., Kalimo,H., Jarvinen,M. (2005). Muscle injuries: biology and treatment. Am J Sports Med. 33, 745-764.
Jebailey,L., Wanono,O., Niu,W., Roessler,J., Rudich,A., Klip,A. (2007). Ceramide- and oxidant-induced insulin resistance involve loss of insulin-dependent rac-activation and actin remodeling in muscle cells. Diabetes. 56, 394-403.
Jongbloed,R.J., Marcelis,C.L., Doevendans,P.A., Schmeitz-Mulkens,J.M., Van Dockum,W.G., Geraedts,J.P., Smeets,H.J. (2003). Variable clinical manifestation of a novel missense mutation in the alpha-tropomyosin (TPM1) gene in familial hypertrophic cardiomyopathy. J Am Coll.Cardiol. %19;41, 981-986.
Joya,J.E., Kee,A.J., Nair-Shalliker,V., Ghoddusi,M., Nguyen,M.A., Luther,P., Hardeman,E.C. (2004). Muscle weakness in a mouse model of nemaline myopathy can be reversed with exercise and reveals a novel myofiber repair mechanism. Hum Mol Genet. 13, 2633-2645.
Judge,L.M., Chamberlain,J.S. (2005). Gene therapy for Duchenne muscular dystrophy: AAV leads the way. Acta Myol. 24, 184-193.
Jung-Ching,L.J., Gustafson-Wagner,E.A., Sinn,H.W., Choi,S., Jaacks,S.M., Wang,D.Z., Evans,S., Li-Chun,L.J. (2005). Structure, Expression, and Function of a Novel Intercalated Disc Protein, Xin. J Med Sci. 25, 215-222.
Kanzaki,M. (2006a). Insulin receptor signals regulating GLUT4 translocation and actin dynamics. Endocr J. 53, 267-293.
Kanzaki,M. (2006b). Insulin receptor signals regulating GLUT4 translocation and actin dynamics. Endocr J. 53, 267-293.
Kanzaki,M., Pessin,J.E. (2001). Insulin-stimulated GLUT4 translocation in adipocytes is dependent upon cortical actin remodeling. J Biol Chem. 276, 42436-42444.
Kee,A.J., Schevzov,G., Nair-Shalliker,V., Robinson,C.S., Vrhovski,B., Ghoddusi,M., Qiu,M.R., Lin,J.J., Weinberger,R., Gunning,P.W., Hardeman,E.C. (2004). Sorting of a nonmuscle tropomyosin to a novel cytoskeletal compartment in skeletal muscle results in muscular dystrophy. J Cell Biol 166, 685-696.
Khayat,Z.A., Tong,P., Yaworsky,K., Bloch,R.J., Klip,A. (2000). Insulin-induced actin filament remodeling colocalizes actin with phosphatidylinositol 3-kinase and GLUT4 in L6 myotubes. J Cell Sci. 113 Pt 2:279-90., 279-290.
King,H., Aubert,R.E., Herman,W.H. (1998). Global burden of diabetes, 1995-2025: prevalence, numerical estimates, and projections. Diabetes Care. 21, 1414-1431.
Kirfel,J., Magin,T.M., Reichelt,J. (2003). Keratins: a structural scaffold with emerging functions. Cell Mol Life Sci. 60, 56-71.
Kostin,S., Scholz,D., Shimada,T., Maeno,Y., Mollnau,H., Hein,S., Schaper,J. (1998). The internal and external protein scaffold of the T-tubular system in cardiomyocytes. Cell Tissue Res. 294, 449-460.
Page 209
Kramer,H., Luke,A. (2007). Obesity and kidney disease: a big dilemma. Curr Opin Nephrol Hypertens. 16, 237-241.
Krendel,M., Mooseker,M.S. (2005). Myosins: tails (and heads) of functional diversity. Physiology (Bethesda.). 20:239-51., 239-251.
Kumar,A., Crawford,K., Close,L., Madison,M., Lorenz,J., Doetschman,T., Pawlowski,S., Duffy,J., Neumann,J., Robbins,J., Boivin,G.P., O'Toole,B.A., Lessard,J.L. (1997). Rescue of cardiac alpha-actin-deficient mice by enteric smooth muscle gamma-actin. Proc Natl Acad Sci U S A. 94, 4406-4411.
Kwiatkowska,M., Poplonska,K., Stepinski,D. (2005). Actin filaments connected with the microtubules of lipotubuloids, cytoplasmic domains rich in lipid bodies and microtubules. Protoplasma. 226, 163-167.
Laing,N.G., Nowak,K.J. (2005). When contractile proteins go bad: the sarcomere and skeletal muscle disease. Bioessays. 27, 809-822.
Laing,N.G., Wilton,S.D., Akkari,P.A., Dorosz,S., Boundy,K., Kneebone,C., Blumbergs,P., White,S., Watkins,H., Love,D.R., . (1995). A mutation in the alpha tropomyosin gene TPM3 associated with autosomal dominant nemaline myopathy. Nat Genet 9, 75-79.
Lambrechts,A., Van Troys,M., Ampe,C. (2004). The actin cytoskeleton in normal and pathological cell motility. Int J Biochem Cell Biol. 36, 1890-1909.
Lauritzen,H.P., Ploug,T., Prats,C., Tavare,J.M., Galbo,H. (2006). Imaging of insulin signaling in skeletal muscle of living mice shows major role of T-tubules. Diabetes. 55, 1300-1306.
Lehman,W., Hatch,V., Korman,V., Rosol,M., Thomas,L., Maytum,R., Geeves,M.A., Van Eyk,J.E., Tobacman,L.S., Craig,R. (2000). Tropomyosin and actin isoforms modulate the localization of tropomyosin strands on actin filaments. J Mol Biol. 302, 593-606.
Lehmann,J.M., Moore,L.B., Smith-Oliver,T.A., Wilkison,W.O., Willson,T.M., Kliewer,S.A. (1995). An antidiabetic thiazolidinedione is a high affinity ligand for peroxisome proliferator-activated receptor gamma (PPAR gamma). J Biol Chem. 270, 12953-12956.
Li,S., Kimura,E., Fall,B.M., Reyes,M., Angello,J.C., Welikson,R., Hauschka,S.D., Chamberlain,J.S. (2005). Stable transduction of myogenic cells with lentiviral vectors expressing a minidystrophin. Gene Ther. 12, 1099-1108.
Lin,Z.X., Holtzer,S., Schultheiss,T., Murray,J., Masaki,T., Fischman,D.A., Holtzer,H. (1989). Polygons and adhesion plaques and the disassembly and assembly of myofibrils in cardiac myocytes. J Cell Biol. 108, 2355-2367.
Littlefield,R., Fowler,V.M. (1998). Defining actin filament length in striated muscle: rulers and caps or dynamic stability? Annu Rev Cell Dev Biol. 14:487-525., 487-525.
Page 210
Liu,H., Bretscher,A. (1992). Characterization of TPM1 disrupted yeast cells indicates an involvement of tropomyosin in directed vesicular transport. J Cell Biol. 118, 285- 299.
Liu,L.B., Omata,W., Kojima,I., Shibata,H. (2003). Insulin recruits GLUT4 from distinct compartments via distinct traffic pathways with differential microtubule dependence in rat adipocytes. J Biol Chem. 278, 30157-30169.
Liu,Q., Muruve,D.A. (2003). Molecular basis of the inflammatory response to adenovirus vectors. Gene Ther. 10, 935-940.
Lloyd,C.M., Berendse,M., Lloyd,D.G., Schevzov,G., Grounds,M.D. (2004). A novel role for non-muscle gamma-actin in skeletal muscle sarcomere assembly. Exp Cell Res. 297, 82-96.
Lo,C.M., Buxton,D.B., Chua,G.C., Dembo,M., Adelstein,R.S., Wang,Y.L. (2004). Nonmuscle myosin IIb is involved in the guidance of fibroblast migration. Mol Biol Cell. 15, 982-989.
Lopez-Soriano,J., Chiellini,C., Maffei,M., Grimaldi,P.A., Argiles,J.M. (2006). Roles of skeletal muscle and peroxisome proliferator-activated receptors in the development and treatment of obesity. Endocr Rev. 27, 318-329.
Lucas,C.A., Hoh,J.F. (2003). Distribution of developmental myosin heavy chains in adult rabbit extraocular muscle: identification of a novel embryonic isoform absent in fetal limb. Invest Ophthalmol Vis Sci. 44, 2450-2456.
Marazzi,G., Bard,F., Klymkowsky,M.W., Rubin,L.L. (1989). Microinjection of a monoclonal antibody against a 37-kD protein (tropomyosin 2) prevents the formation of new acetylcholine receptor clusters. J Cell Biol 109, 2337-2344.
Martini,F.H., Ober,W.C., Garrison,C.W., Welch,K., Hutching,R.T. (2001). Anatomy and Physiology. London: Prentice Hall.
Maupin,P., Phillips,C.L., Adelstein,R.S., Pollard,T.D. (1994). Differential localization of myosin-II isozymes in human cultured cells and blood cells. J Cell Sci. 107, 3077- 3090.
Mayer,U., Jurgens,G. (2002). Microtubule cytoskeleton: a track record. Curr Opin Plant Biol. 5, 494-501.
Mazeres,G., Leloup,L., Daury,L., Cottin,P., Brustis,J.J. (2006). Myoblast attachment and spreading are regulated by different patterns by ubiquitous calpains. Cell Motil Cytoskeleton. 63, 193-207.
McCarthy,A.M., Spisak,K.O., Brozinick,J.T., Elmendorf,J.S. (2006). Loss of cortical actin filaments in insulin-resistant skeletal muscle cells impairs GLUT4 vesicle trafficking and glucose transport. Am J Physiol Cell Physiol. 291, C860-C868.
McKillop,D.F., Geeves,M.A. (1993). Regulation of the interaction between actin and myosin subfragment 1: evidence for three states of the thin filament. Biophys J. 65, 693-701.
Page 211
McLoon,L.K., Rowe,J., Wirtschafter,J., McCormick,K.M. (2004). Continuous myofiber remodeling in uninjured extraocular myofibers: myonuclear turnover and evidence for apoptosis. Muscle Nerve 29, 707-715.
McMichael,B.K., Kotadiya,P., Singh,T., Holliday,L.S., Lee,B.S. (2006). Tropomyosin isoforms localize to distinct microfilament populations in osteoclasts. Bone. 39, 694- 705.
McNeil,P.L., Steinhardt,R.A. (2003). Plasma membrane disruption: repair, prevention, adaptation. Annu Rev Cell Dev Biol. 19:697-731., 697-731.
Mendler,L., Zador,E., Dux,L., Wuytack,F. (1998). mRNA levels of myogenic regulatory factors in rat slow and fast muscles regenerating from notexin-induced necrosis. Neuromuscul.Disord 8, 533-541.
Miki,H., Okada,Y., Hirokawa,N. (2005). Analysis of the kinesin superfamily: insights into structure and function. Trends Cell Biol. 15, 467-476.
Milner,D.J., Mavroidis,M., Weisleder,N., Capetanaki,Y. (2000). Desmin cytoskeleton linked to muscle mitochondrial distribution and respiratory function. J Cell Biol. 150, 1283-1298.
Minetti,C., Bado,M., Broda,P., Sotgia,F., Bruno,C., Galbiati,F., Volonte,D., Lucania,G., Pavan,A., Bonilla,E., Lisanti,M.P., Cordone,G. (2002). Impairment of caveolae formation and T-system disorganization in human muscular dystrophy with caveolin-3 deficiency. Am J Pathol. 160, 265-270.
Mitchell,R.D., Saito,A., Palade,P., Fleischer,S. (1983). Morphology of isolated triads. J Cell Biol. 96, 1017-1029.
Mokdad,A.H., Ford,E.S., Bowman,B.A., Dietz,W.H., Vinicor,F., Bales,V.S., Marks,J.S. (2003). Prevalence of obesity, diabetes, and obesity-related health risk factors, 2001. JAMA. 289, 76-79.
Molero,J.C., Jensen,T.E., Withers,P.C., Couzens,M., Herzog,H., Thien,C.B., Langdon,W.Y., Walder,K., Murphy,M.A., Bowtell,D.D., James,D.E., Cooney,G.J. (2004). c-Cbl-deficient mice have reduced adiposity, higher energy expenditure, and improved peripheral insulin action. J Clin Invest. 114, 1326-1333.
Montarras,D., Morgan,J., Collins,C., Relaix,F., Zaffran,S., Cumano,A., Partridge,T., Buckingham,M. (2005). Direct isolation of satellite cells for skeletal muscle regeneration. Science. 309, 2064-2067.
Muthuchamy,M., Grupp,I.L., Grupp,G., O'Toole,B.A., Kier,A.B., Boivin,G.P., Neumann,J., Wieczorek,D.F. (1995). Molecular and physiological effects of overexpressing striated muscle beta-tropomyosin in the adult murine heart. J Biol Chem. 270, 30593-30603.
Myers,K.A., He,Y., Hasaka,T.P., Baas,P.W. (2006). Microtubule transport in the axon: Re-thinking a potential role for the actin cytoskeleton. Neuroscientist. 12, 107-118.
Page 212
Nair-Shalliker,V., Kee,A.J., Joya,J.E., Lucas,C.A., Hoh,J.F., Hardeman,E.C. (2004). Myofiber adaptational response to exercise in a mouse model of nemaline myopathy. Muscle Nerve. 30, 470-480.
Nakashima,K., Sato,N., Nakagaki,T., Abe,H., Ono,S., Obinata,T. (2005). Two mouse cofilin isoforms, muscle-type (MCF) and non-muscle type (NMCF), interact with F- actin with different efficiencies. J Biochem (Tokyo). 138, 519-526.
Nehlin,J.O., Mogensen,J.P., Petterson,I., Jeppesen,L., Fleckner,J., Wulff,E.M., Sauerberg,P. (2006). Selective PPAR agonists for the treatment of type 2 diabetes. Ann N Y Acad Sci. 1067:448-53., 448-453.
Nesto,R.W., Bell,D., Bonow,R.O., Fonseca,V., Grundy,S.M., Horton,E.S., Le Winter,M., Porte,D., Semenkovich,C.F., Smith,S., Young,L.H., Kahn,R. (2003). Thiazolidinedione use, fluid retention, and congestive heart failure: a consensus statement from the American Heart Association and American Diabetes Association. October 7, 2003. Circulation. 108, 2941-2948.
Neuschwander-Tetri,B.A. (2007). Fatty liver and the metabolic syndrome. Curr Opin Gastroenterol. 23, 193-198.
Nguyen,T.M., Ellis,J.M., Love,D.R., Davies,K.E., Gatter,K.C., Dickson,G., Morris,G.E. (1991). Localization of the DMDL gene-encoded dystrophin-related protein using a panel of nineteen monoclonal antibodies: presence at neuromuscular junctions, in the sarcolemma of dystrophic skeletal muscle, in vascular and other smooth muscles, and in proliferating brain cell lines. J Cell Biol. 115, 1695-1700.
Nieminen,M., Henttinen,T., Merinen,M., Marttila-Ichihara,F., Eriksson,J.E., Jalkanen,S. (2006). Vimentin function in lymphocyte adhesion and transcellular migration. Nat Cell Biol. 8, 156-162.
Nishida,E., Maekawa,S., Sakai,H. (1984). Cofilin, a protein in porcine brain that binds to actin filaments and inhibits their interactions with myosin and tropomyosin. Biochemistry. 23, 5307-5313.
North,K.N., Beggs,A.H. (1996). Deficiency of a skeletal muscle isoform of alpha- actinin (alpha-actinin-3) in merosin-positive congenital muscular dystrophy. Neuromuscul.Disord. 6, 229-235.
O'Neill,A., Williams,M.W., Resneck,W.G., Milner,D.J., Capetanaki,Y., Bloch,R.J. (2002). Sarcolemmal organization in skeletal muscle lacking desmin: evidence for cytokeratins associated with the membrane skeleton at costameres. Mol Biol Cell 13, 2347-2359.
Obermann,W.M., Gautel,M., Weber,K., Furst,D.O. (1997). Molecular structure of the sarcomeric M band: mapping of titin and myosin binding domains in myomesin and the identification of a potential regulatory phosphorylation site in myomesin. EMBO J. 16, 211-220.
Okiyoneda,T., Lukacs,G.L. (2007). Cell surface dynamics of CFTR: The ins and outs. Biochim Biophys Acta. 1773, 476-479.
Page 213
Olefsky,J.M., Saltiel,A.R. (2000). PPAR gamma and the treatment of insulin resistance. Trends Endocrinol Metab. 11, 362-368.
Olson,A.L., Eyster,C.A., Duggins,Q.S., Knight,J.B. (2003). Insulin promotes formation of polymerized microtubules by a phosphatidylinositol 3-kinase-independent, actin- dependent pathway in 3T3-L1 adipocytes. Endocrinology 144, 5030-5039.
Olson,A.L., Knight,J.B., Pessin,J.E. (1997). Syntaxin 4, VAMP2, and/or VAMP3/cellubrevin are functional target membrane and vesicle SNAP receptors for insulin-stimulated GLUT4 translocation in adipocytes. Mol Cell Biol 17, 2425-2435.
Olson,T.M., Kishimoto,N.Y., Whitby,F.G., Michels,V.V. (2001). Mutations that alter the surface charge of alpha-tropomyosin are associated with dilated cardiomyopathy. J Mol Cell Cardiol. 33, 723-732.
Omary,M.B., Coulombe,P.A., McLean,W.H. (2004). Intermediate filament proteins and their associated diseases. N Engl J Med. 351, 2087-2100.
Omata,W., Shibata,H., Li,L., Takata,K., Kojima,I. (2000). Actin filaments play a critical role in insulin-induced exocytotic recruitment but not in endocytosis of GLUT4 in isolated rat adipocytes. Biochem J. 346 Pt 2:321-8., 321-328.
Ono,S. (2003). Regulation of actin filament dynamics by actin depolymerizing factor/cofilin and actin-interacting protein 1: new blades for twisted filaments. Biochemistry. 42, 13363-13370.
Ono,S., Ono,K. (2002). Tropomyosin inhibits ADF/cofilin-dependent actin filament dynamics. J Cell Biol. 156, 1065-1076.
Pacholsky,D., Vakeel,P., Himmel,M., Lowe,T., Stradal,T., Rottner,K., Furst,D.O., van der Ven,P.F. (2004). Xin repeats define a novel actin-binding motif. J Cell Sci. 117, 5257-5268.
Palmiter,K.A., Kitada,Y., Muthuchamy,M., Wieczorek,D.F., Solaro,R.J. (1996). Exchange of beta- for alpha-tropomyosin in hearts of transgenic mice induces changes in thin filament response to Ca2+, strong cross-bridge binding, and protein phosphorylation. J Biol Chem. 271, 11611-11614.
Paolini,C., Fessenden,J.D., Pessah,I.N., Franzini-Armstrong,C. (2004). Evidence for conformational coupling between two calcium channels. Proc Natl Acad Sci U S A. 101, 12748-12752.
Papa,I., Astier,C., Kwiatek,O., Raynaud,F., Bonnal,C., Lebart,M.C., Roustan,C., Benyamin,Y. (1999). Alpha actinin-CapZ, an anchoring complex for thin filaments in Z-line. J Muscle Res Cell Motil. 20, 187-197.
Pardo,J.V., Siliciano,J.D., Craig,S.W. (1983). A vinculin-containing cortical lattice in skeletal muscle: transverse lattice elements ("costameres") mark sites of attachment between myofibrils and sarcolemma. Proc Natl Acad Sci U S A. 80, 1008-1012.
Paulin,D., Huet,A., Khanamyrian,L., Xue,Z. (2004). Desminopathies in muscle disease. J Pathol. 204, 418-427.
Page 214
Percival,J.M., Hughes,J.A., Brown,D.L., Schevzov,G., Heimann,K., Vrhovski,B., Bryce,N., Stow,J.L., Gunning,P.W. (2004). Targeting of a tropomyosin isoform to short microfilaments associated with the Golgi complex. Mol Biol Cell 15, 268-280.
Percival,J.M., Thomas,G., Cock,T.A., Gardiner,E.M., Jeffrey,P.L., Lin,J.J., Weinberger,R.P., Gunning,P. (2000). Sorting of tropomyosin isoforms in synchronised NIH 3T3 fibroblasts: evidence for distinct microfilament populations. Cell Motil Cytoskeleton 47, 189-208.
Perry,S.V. (2001). Vertebrate tropomyosin: distribution, properties and function. J Muscle Res Cell Motil. 22, 5-49.
Petit,C. (2001). Usher syndrome: from genetics to pathogenesis. Annu Rev Genomics Hum Genet. 2:271-97., 271-297.
Pittenger,M.F., Kistler,A., Helfman,D.M. (1995). Alternatively spliced exons of the beta tropomyosin gene exhibit different affinities for F-actin and effects with nonmuscle caldesmon. J Cell Sci. 108, 3253-3265.
Plant, D. R., Colarossi, F. E., and Lynch, G. S. Notexin causes greater myotoxic damage and slower functional repair in mouse skeletal muscles than bupivacaine. Muscle Nerve . 2006. Ref Type: Generic
Ploug,T., van Deurs,B., Ai,H., Cushman,S.W., Ralston,E. (1998). Analysis of GLUT4 distribution in whole skeletal muscle fibers: identification of distinct storage compartments that are recruited by insulin and muscle contractions. J Cell Biol. 142, 1429-1446.
Politi,P.K., Havaki,S., Manta,P., Lyritis,G. (2006). Bupivacaine-induced regeneration of rat soleus muscle: ultrastructural and immunohistochemical aspects. Ultrastruct.Pathol. 30, 461-469.
Pollard,T.D., Blanchoin,L., Mullins,R.D. (2000). Molecular mechanisms controlling actin filament dynamics in nonmuscle cells. Annu Rev Biophys Biomol Struct. 29:545- 76., 545-576.
Pollard,T.D., Borisy,G.G. (2003). Cellular motility driven by assembly and disassembly of actin filaments. Cell. 112, 453-465.
Pollard,T.D., Cooper,J.A. (1986). Actin and actin-binding proteins. A critical evaluation of mechanisms and functions. Annu Rev Biochem. 55:987-1035., 987-1035.
Porter,N.C., Resneck,W.G., O'Neill,A., Van Rossum,D.B., Stone,M.R., Bloch,R.J. (2005). Association of small ankyrin 1 with the sarcoplasmic reticulum. Mol Membr Biol. 22, 421-432.
Porteus,M.H., Connelly,J.P., Pruett,S.M. (2006). A look to future directions in gene therapy research for monogenic diseases. PLoS Genet. 2, e133.
Prat,A.G., Xiao,Y.F., Ausiello,D.A., Cantiello,H.F. (1995). cAMP-independent regulation of CFTR by the actin cytoskeleton. Am J Physiol. 268, C1552-C1561.
Page 215
Price,F.D., Kuroda,K., Rudnicki,M.A. (2007). Stem cell based therapies to treat muscular dystrophy. Biochim Biophys Acta. 1772, 272-283.
Procaccio,V., Salazar,G., Ono,S., Styers,M.L., Gearing,M., Davila,A., Jimenez,R., Juncos,J., Gutekunst,C.A., Meroni,G., Fontanella,B., Sontag,E., Sontag,J.M., Faundez,V., Wainer,B.H. (2006). A mutation of beta -actin that alters depolymerization dynamics is associated with autosomal dominant developmental malformations, deafness, and dystonia. Am J Hum Genet. 78, 947-960.
Pruyne,D.W., Schott,D.H., Bretscher,A. (1998). Tropomyosin-containing actin cables direct the Myo2p-dependent polarized delivery of secretory vesicles in budding yeast. J Cell Biol. 143, 1931-1945.
Quenneville,S.P., Chapdelaine,P., Skuk,D., Paradis,M., Goulet,M., Rousseau,J., Xiao,X., Garcia,L., Tremblay,J.P. (2007). Autologous transplantation of muscle precursor cells modified with a lentivirus for muscular dystrophy: human cells and primate models. Mol Ther. 15, 431-438.
Quenneville,S.P., Tremblay,J.P. (2006). Ex vivo modification of cells to induce a muscle-based expression. Curr Gene Ther. 6, 625-632.
Ralphs,J.R., Waggett,A.D., Benjamin,M. (2002). Actin stress fibres and cell-cell adhesion molecules in tendons: organisation in vivo and response to mechanical loading of tendon cells in vitro. Matrix Biol. 21, 67-74.
Ralston,E., Ploug,T. (1999). Caveolin-3 is associated with the T-tubules of mature skeletal muscle fibers. Exp Cell Res. 246, 510-515.
Rando,T.A. (2001). The dystrophin-glycoprotein complex, cellular signaling, and the regulation of cell survival in the muscular dystrophies. Muscle Nerve. 24, 1575-1594.
Rando,T.A. (2007). Non-viral gene therapy for Duchenne muscular dystrophy: progress and challenges. Biochim Biophys Acta. 1772, 263-271.
Rappaport,L., Oliviero,P., Samuel,J.L. (1998). Cytoskeleton and mitochondrial morphology and function. Mol Cell Biochem. 184, 101-105.
Reggiani,C., Bottinelli,R., Stienen,G.J. (2000). Sarcomeric Myosin Isoforms: Fine Tuning of a Molecular Motor. News Physiol Sci 15:26-33., 26-33.
Reid,E., Kloos,M., Ashley-Koch,A., Hughes,L., Bevan,S., Svenson,I.K., Graham,F.L., Gaskell,P.C., Dearlove,A., Pericak-Vance,M.A., Rubinsztein,D.C., Marchuk,D.A. (2002). A kinesin heavy chain (KIF5A) mutation in hereditary spastic paraplegia (SPG10). Am J Hum Genet. 71, 1189-1194.
Rethinasamy,P., Muthuchamy,M., Hewett,T., Boivin,G., Wolska,B.M., Evans,C., Solaro,R.J., Wieczorek,D.F. (1998). Molecular and physiological effects of alpha- tropomyosin ablation in the mouse. Circ Res 82, 116-123.
Rhee,D., Sanger,J.M., Sanger,J.W. (1994). The premyofibril: evidence for its role in myofibrillogenesis. Cell Motil Cytoskeleton. 28, 1-24.
Page 216
Robbins,J. (1998). Alpha-tropomyosin knockouts: a blow against transcriptional chauvinism. Circ Res. 82, 134-136.
Rodnick,K.J., Slot,J.W., Studelska,D.R., Hanpeter,D.E., Robinson,L.J., Geuze,H.J., James,D.E. (1992). Immunocytochemical and biochemical studies of GLUT4 in rat skeletal muscle. J Biol Chem. 267, 6278-6285.
Rubenstein,P.A. (1990). The functional importance of multiple actin isoforms. Bioessays. 12, 309-315.
Ruiz-Canada,C., Budnik,V. (2006). Synaptic cytoskeleton at the neuromuscular junction. Int Rev Neurobiol. 75:217-36., 217-236.
Rybakova,I.N., Patel,J.R., Ervasti,J.M. (2000). The dystrophin complex forms a mechanically strong link between the sarcolemma and costameric actin. J Cell Biol 150, 1209-1214.
Saito,A., Seiler,S., Chu,A., Fleischer,S. (1984). Preparation and morphology of sarcoplasmic reticulum terminal cisternae from rabbit skeletal muscle. J Cell Biol. 99, 875-885.
Sampaolesi,M., Blot,S., D'Antona,G., Granger,N., Tonlorenzi,R., Innocenzi,A., Mognol,P., Thibaud,J.L., Galvez,B.G., Barthelemy,I., Perani,L., Mantero,S., Guttinger,M., Pansarasa,O., Rinaldi,C., Cusella De Angelis,M.G., Torrente,Y., Bordignon,C., Bottinelli,R., Cossu,G. (2006). Mesoangioblast stem cells ameliorate muscle function in dystrophic dogs. Nature. 444, 574-579.
Samstag,Y., Eibert,S.M., Klemke,M., Wabnitz,G.H. (2003). Actin cytoskeletal dynamics in T lymphocyte activation and migration. J Leukoc Biol. 73, 30-48.
Sanes,J.R., Lichtman,J.W. (1999). Development of the vertebrate neuromuscular junction. Annu Rev Neurosci 22:389-442., 389-442.
Sanger,J.W., Chowrashi,P., Shaner,N.C., Spalthoff,S., Wang,J., Freeman,N.L., Sanger,J.M. (2002). Myofibrillogenesis in skeletal muscle cells. Clin Orthop Relat Res. S153-S162.
Sanger,J.W., Kang,S., Siebrands,C.C., Freeman,N., Du,A., Wang,J., Stout,A.L., Sanger,J.M. (2006). How to build a myofibril. J Muscle Res Cell Motil. 1-12.
Sanoudou,D., Corbett,M.A., Han,M., Ghoddusi,M., Nguyen,M.A., Vlahovich,N., Hardeman,E.C., Beggs,A.H. (2006). Skeletal muscle repair in a mouse model of nemaline myopathy. Hum Mol Genet. 15, 2603-2612.
Sanoudou,D., Haslett,J.N., Kho,A.T., Guo,S., Gazda,H.T., Greenberg,S.A., Lidov,H.G., Kohane,I.S., Kunkel,L.M., Beggs,A.H. (2003). Expression profiling reveals altered satellite cell numbers and glycolytic enzyme transcription in nemaline myopathy muscle. Proc Natl Acad Sci U S A. 100, 4666-4671.
Schevzov, G. 2006. Ref Type: Unpublished Work
Page 217
Schevzov,G., Bryce,N.S., Almonte-Baldonado,R., Joya,J., Lin,J.J., Hardeman,E., Weinberger,R., Gunning,P. (2005a). Specific features of neuronal size and shape are regulated by tropomyosin isoforms. Mol Biol Cell. 16, 3425-3437.
Schevzov,G., Gunning,P., Jeffrey,P.L., Temm-Grove,C., Helfman,D.M., Lin,J.J., Weinberger,R.P. (1997). Tropomyosin localization reveals distinct populations of microfilaments in neurites and growth cones. Mol Cell Neurosci. 8, 439-454.
Schevzov,G., Lloyd,C., Gunning,P. (1992). High level expression of transfected beta- and gamma-actin genes differentially impacts on myoblast cytoarchitecture. J Cell Biol. 117, 775-785.
Schevzov,G., Vrhovski,B., Bryce,N.S., Elmir,S., Qiu,M.R., O'neill,G.M., Yang,N., Verrills,N.M., Kavallaris,M., Gunning,P.W. (2005b). Tissue-specific tropomyosin isoform composition. J Histochem Cytochem. 53, 557-570.
Schildmeyer,L.A., Braun,R., Taffet,G., Debiasi,M., Burns,A.E., Bradley,A., Schwartz,R.J. (2000). Impaired vascular contractility and blood pressure homeostasis in the smooth muscle alpha-actin null mouse. FASEB J. 14, 2213-2220.
Schnorrer,F., Dickson,B.J. (2004). Muscle building; mechanisms of myotube guidance and attachment site selection. Dev Cell 7, 9-20.
Schoenauer,R., Bertoncini,P., Machaidze,G., Aebi,U., Perriard,J.C., Hegner,M., Agarkova,I. (2005). Myomesin is a molecular spring with adaptable elasticity. J Mol Biol. 349, 367-379.
Schroder,R., Reimann,J., Salmikangas,P., Clemen,C.S., Hayashi,Y.K., Nonaka,I., Arahata,K., Carpen,O. (2003). Beyond LGMD1A: myotilin is a component of central core lesions and nemaline rods. Neuromuscul.Disord. 13, 451-455.
Schwenk,F., Baron,U., Rajewsky,K. (1995). A cre-transgenic mouse strain for the ubiquitous deletion of loxP-flanked gene segments including deletion in germ cells. Nucleic Acids Res. 23, 5080-5081.
Sealock,R., Paschal,B., Beckerle,M., Burridge,K. (1986). Talin is a post-synaptic component of the rat neuromuscular junction. Exp Cell Res. 163, 143-150.
Semple,R.K., Chatterjee,V.K., O'Rahilly,S. (2006). PPAR gamma and human metabolic disease. J Clin Invest. 116, 581-589.
Sharma,A.M., Staels,B. (2007). Review: Peroxisome proliferator-activated receptor gamma and adipose tissue--understanding obesity-related changes in regulation of lipid and glucose metabolism. J Clin Endocrinol Metab. 92, 386-395.
Shawlot,W., Deng,J.M., Fohn,L.E., Behringer,R.R. (1998). Restricted beta- galactosidase expression of a hygromycin-lacZ gene targeted to the beta-actin locus and embryonic lethality of beta-actin mutant mice. Transgenic Res. 7, 95-103.
Sicinski,P., Geng,Y., Ryder-Cook,A.S., Barnard,E.A., Darlison,M.G., Barnard,P.J. (1989). The molecular basis of muscular dystrophy in the mdx mouse: a point mutation. Science. 244, 1578-1580.
Page 218
Sinn,H.W., Balsamo,J., Lilien,J., Lin,J.J. (2002). Localization of the novel Xin protein to the adherens junction complex in cardiac and skeletal muscle during development. Dev Dyn. 225, 1-13.
Skuk,D., Roy,B., Goulet,M., Chapdelaine,P., Bouchard,J.P., Roy,R., Dugre,F.J., Lachance,J.G., Deschenes,L., Helene,S., Sylvain,M., Tremblay,J.P. (2004). Dystrophin expression in myofibers of Duchenne muscular dystrophy patients following intramuscular injections of normal myogenic cells. Mol Ther. 9, 475-482.
Small,J.V., Resch,G.P. (2005). The comings and goings of actin: coupling protrusion and retraction in cell motility. Curr Opin Cell Biol. 17, 517-523.
Smith,D.A., Geeves,M.A. (2003). Cooperative regulation of myosin-actin interactions by a continuous flexible chain II: actin-tropomyosin-troponin and regulation by calcium. Biophys J. 84, 3168-3180.
Smythe,E., Ayscough,K.R. (2006). Actin regulation in endocytosis. J Cell Sci. 119, 4589-4598.
Sonnemann,K.J., Fitzsimons,D.P., Patel,J.R., Liu,Y., Schneider,M.F., Moss,R.L., Ervasti,J.M. (2006). Cytoplasmic gamma-actin is not required for skeletal muscle development but its absence leads to a progressive myopathy. Dev Cell. 11, 387-397.
Sparrow,J.C., Nowak,K.J., Durling,H.J., Beggs,A.H., Wallgren-Pettersson,C., Romero,N., Nonaka,I., Laing,N.G. (2003). Muscle disease caused by mutations in the skeletal muscle alpha-actin gene (ACTA1). Neuromuscul.Disord 13, 519-531.
Spence,H.J., Chen,Y.J., Winder,S.J. (2002). Muscular dystrophies, the cytoskeleton and cell adhesion. Bioessays 24, 542-552.
Spinler,S.A. (2006). Challenges associated with metabolic syndrome. Pharmacotherapy. 26, 209S-217S.
Staiger,C.J., Blanchoin,L. (2006). Actin dynamics: old friends with new stories. Curr Opin Plant Biol. 9, 554-562.
Stamnes,M. (2002). Regulating the actin cytoskeleton during vesicular transport. Curr Opin Cell Biol. 14, 428-433.
Starr,D.A., Han,M. (2003). ANChors away: an actin based mechanism of nuclear positioning. J Cell Sci. 116, 211-216.
Steimle,P.A., Fulcher,F.K., Patel,Y.M. (2005). A novel role for myosin II in insulin- stimulated glucose uptake in 3T3-L1 adipocytes. Biochem Biophys Res Commun. 331, 1560-1565.
Strober,J.B. (2006). Therapeutics in duchenne muscular dystrophy. NeuroRx. 3, 225- 234.
Suter,D.M., Espindola,F.S., Lin,C.H., Forscher,P., Mooseker,M.S. (2000). Localization of unconventional myosins V and VI in neuronal growth cones. J Neurobiol. 42, 370- 382.
Page 219
Svitkina,T.M., Borisy,G.G. (1999). Arp2/3 complex and actin depolymerizing factor/cofilin in dendritic organization and treadmilling of actin filament array in lamellipodia. J Cell Biol. 145, 1009-1026.
Swiatecka-Urban,A., Boyd,C., Coutermarsh,B., Karlson,K.H., Barnaby,R., Aschenbrenner,L., Langford,G.M., Hasson,T., Stanton,B.A. (2004). Myosin VI regulates endocytosis of the cystic fibrosis transmembrane conductance regulator. J Biol Chem. 279, 38025-38031.
Takeda,K., Yu,Z.X., Qian,S., Chin,T.K., Adelstein,R.S., Ferrans,V.J. (2000). Nonmuscle myosin II localizes to the Z-lines and intercalated discs of cardiac muscle and to the Z-lines of skeletal muscle. Cell Motil Cytoskeleton. 46, 59-68.
Takekura,H., Flucher,B.E., Franzini-Armstrong,C. (2001). Sequential docking, molecular differentiation, and positioning of T-Tubule/SR junctions in developing mouse skeletal muscle. Dev Biol 239, 204-214.
Tanabe,Y., Esaki,K., Nomura,T. (1986). Skeletal muscle pathology in X chromosome- linked muscular dystrophy (mdx) mouse. Acta Neuropathol (Berl) 69, 91-95.
Tang,Z., Scherer,P.E., Okamoto,T., Song,K., Chu,C., Kohtz,D.S., Nishimoto,I., Lodish,H.F., Lisanti,M.P. (1996). Molecular cloning of caveolin-3, a novel member of the caveolin gene family expressed predominantly in muscle. J Biol Chem. 271, 2255- 2261.
Taniguchi,M., Geng,X., Glazier,K.D., Dasgupta,A., Lin,J.J., Das,K.M. (2001). Cellular immune response against tropomyosin isoform 5 in ulcerative colitis. Clin Immunol. 101, 289-295.
Tardiff,J.C. (2005). Sarcomeric proteins and familial hypertrophic cardiomyopathy: linking mutations in structural proteins to complex cardiovascular phenotypes. Heart Fail.Rev. 10, 237-248.
Tellam,J.T., Macaulay,S.L., McIntosh,S., Hewish,D.R., Ward,C.W., James,D.E. (1997). Characterization of Munc-18c and syntaxin-4 in 3T3-L1 adipocytes. Putative role in insulin-dependent movement of GLUT-4. J Biol Chem. 272, 6179-6186.
Temm-Grove,C.J., Guo,W., Helfman,D.M. (1996). Low molecular weight rat fibroblast tropomyosin 5 (TM-5): cDNA cloning, actin-binding, localization, and coiled-coil interactions. Cell Motil Cytoskeleton. 33, 223-240.
Thong,F.S., Bilan,P.J., Klip,A. (2007). The Rab GTPase-Activating Protein AS160 Integrates Akt, Protein Kinase C, and AMP-Activated Protein Kinase Signals Regulating GLUT4 Traffic. Diabetes. 56, 414-423.
Tidball,J.G., Daniel,T.L. (1986). Myotendinous junctions of tonic muscle cells: structure and loading. Cell Tissue Res. 245, 315-322.
Tilney,L.G., Bonder,E.M., DeRosier,D.J. (1981). Actin filaments elongate from their membrane-associated ends. J Cell Biol. 90, 485-494.
Page 220
Tong,P., Khayat,Z.A., Huang,C., Patel,N., Ueyama,A., Klip,A. (2001). Insulin-induced cortical actin remodeling promotes GLUT4 insertion at muscle cell membrane ruffles. J Clin Invest. 108, 371-381.
Torok,D., Patel,N., Jebailey,L., Thong,F.S., Randhawa,V.K., Klip,A., Rudich,A. (2004). Insulin but not PDGF relies on actin remodeling and on VAMP2 for GLUT4 translocation in myoblasts. J Cell Sci. 117, 5447-5455.
Tsakiridis,T., Tong,P., Matthews,B., Tsiani,E., Bilan,P.J., Klip,A., Downey,G.P. (1999). Role of the actin cytoskeleton in insulin action. Microsc Res Tech. 47, 79-92.
Tsakiridis,T., Vranic,M., Klip,A. (1994). Disassembly of the actin network inhibits insulin-dependent stimulation of glucose transport and prevents recruitment of glucose transporters to the plasma membrane. J Biol Chem. 269, 29934-29942.
Tskhovrebova,L., Trinick,J. (2003). Titin: properties and family relationships. Nat Rev Mol Cell Biol. 4, 679-689.
Tuxworth,R.I., Titus,M.A. (2000). Unconventional myosins: anchors in the membrane traffic relay. Traffic. 1, 11-18.
Urish,K., Kanda,Y., Huard,J. (2005). Initial failure in myoblast transplantation therapy has led the way toward the isolation of muscle stem cells: potential for tissue regeneration. Curr Top Dev Biol. 68:263-80., 263-280.
Vafiadaki,E., Reis,A., Keers,S., Harrison,R., Anderson,L.V., Raffelsberger,T., Ivanova,S., Hoger,H., Bittner,R.E., Bushby,K., Bashir,R. (2001). Cloning of the mouse dysferlin gene and genomic characterization of the SJL-Dysf mutation. Neuroreport 12, 625-629.
Vallotton,P., Gupton,S.L., Waterman-Storer,C.M., Danuser,G. (2004). Simultaneous mapping of filamentous actin flow and turnover in migrating cells by quantitative fluorescent speckle microscopy. Proc Natl Acad Sci U S A. 101, 9660-9665. van der Ven,P.F., Obermann,W.M., Lemke,B., Gautel,M., Weber,K., Furst,D.O. (2000). Characterization of muscle filamin isoforms suggests a possible role of gamma- filamin/ABP-L in sarcomeric Z-disc formation. Cell Motil Cytoskeleton. 45, 149-162.
van Deurs,B., Roepstorff,K., Hommelgaard,A.M., Sandvig,K. (2003). Caveolae: anchored, multifunctional platforms in the lipid ocean. Trends Cell Biol. 13, 92-100.
Van Dijk,J., Knight,A.E., Molloy,J.E., Chaussepied,P. (2002). Characterization of three regulatory states of the striated muscle thin filament. J Mol Biol 323, 475-489.
Vega-Riveroll,L.J., Wylie,S.R., Loughna,P.T., Parson,S.H., Chantler,P.D. (2005). Nonmuscle myosins IIA and IIB are present in adult motor nerve terminals. Neuroreport. 16, 1143-1146.
Vera,C., Sood,A., Gao,K.M., Yee,L.J., Lin,J.J., Sung,L.A. (2000). Tropomodulin- binding site mapped to residues 7-14 at the N-terminal heptad repeats of tropomyosin isoform 5. Arch Biochem Biophys. 378, 16-24.
Page 221
Vrhovski,B., Lemckert,F., Gunning,P. (2004). Modification of the tropomyosin isoform composition of actin filaments in the brain by deletion of an alternatively spliced exon. Neuropharmacology. 47, 684-693.
Vrhovski,B., McKay,K., Schevzov,G., Gunning,P.W., Weinberger,R.P. (2005). Smooth muscle-specific alpha tropomyosin is a marker of fully differentiated smooth muscle in lung. J Histochem Cytochem. 53, 875-883.
Wahli,W., Braissant,O., Desvergne,B. (1995). Peroxisome proliferator activated receptors: transcriptional regulators of adipogenesis, lipid metabolism and more... Chem Biol. 2, 261-266.
Wallgren-Pettersson,C., Jasani,B., Newman,G.R., Morris,G.E., Jones,S., Singhrao,S., Clarke,A., Virtanen,I., Holmberg,C., Rapola,J. (1995). Alpha-actinin in nemaline bodies in congenital nemaline myopathy: immunological confirmation by light and electron microscopy. Neuromuscul.Disord 5, 93-104.
Wang,J., Shaner,N., Mittal,B., Zhou,Q., Chen,J., Sanger,J.M., Sanger,J.W. (2005). Dynamics of Z-band based proteins in developing skeletal muscle cells. Cell Motil Cytoskeleton. 61, 34-48.
Wang,K., Wright,J. (1988). Architecture of the sarcomere matrix of skeletal muscle: immunoelectron microscopic evidence that suggests a set of parallel inextensible nebulin filaments anchored at the Z line. J Cell Biol. 107, 2199-2212.
Wang,L., Hayashi,H., Ebina,Y. (1999). Transient effect of platelet-derived growth factor on GLUT4 translocation in 3T3-L1 adipocytes. J Biol Chem. 274, 19246-19253.
Wang,Q., Bilan,P.J., Tsakiridis,T., Hinek,A., Klip,A. (1998). Actin filaments participate in the relocalization of phosphatidylinositol3-kinase to glucose transporter- containing compartments and in the stimulation of glucose uptake in 3T3-L1 adipocytes. Biochem J. 331, 917-928.
Wang,Z., Kuhr,C.S., Allen,J.M., Blankinship,M., Gregorevic,P., Chamberlain,J.S., Tapscott,S.J., Storb,R. (2007). Sustained AAV-mediated Dystrophin Expression in a Canine Model of Duchenne Muscular Dystrophy with a Brief Course of Immunosuppression. Mol Ther. ..
Warrick,H.M., Spudich,J.A. (1987). Myosin structure and function in cell motility. Annu Rev Cell Biol. 3:379-421., 379-421.
Watchko,J.F., O'Day,T.L., Hoffman,E.P. (2002). Functional characteristics of dystrophic skeletal muscle: insights from animal models. J Appl Physiol 93, 407-417.
Wattanasirichaigoon,D., Swoboda,K.J., Takada,F., Tong,H.Q., Lip,V., Iannaccone,S.T., Wallgren-Pettersson,C., Laing,N.G., Beggs,A.H. (2002). Mutations of the slow muscle alpha-tropomyosin gene, TPM3, are a rare cause of nemaline myopathy. Neurology 59, 613-617.
Weber,A., Pennise,C.R., Babcock,G.G., Fowler,V.M. (1994). Tropomodulin caps the pointed ends of actin filaments. J Cell Biol. 127, 1627-1635.
Page 222
Wegner,A. (1982). Kinetic analysis of actin assembly suggests that tropomyosin inhibits spontaneous fragmentation of actin filaments. J Mol Biol. 161, 217-227.
Wehrle-Haller,B., Imhof,B.A. (2003). Actin, microtubules and focal adhesion dynamics during cell migration. Int J Biochem Cell Biol. 35, 39-50.
Weinberger,R.P., Henke,R.C., Tolhurst,O., Jeffrey,P.L., Gunning,P. (1993). Induction of neuron-specific tropomyosin mRNAs by nerve growth factor is dependent on morphological differentiation. J Cell Biol. 120, 205-215.
Weir,A. (2000). Muscular dystrophy. Curr Biol. 10, R92.
Weller,A.H., Magliato,S.A., Bell,K.P., Rosenberg,N.L. (1997). Spontaneous myopathy in the SJL/J mouse: pathology and strength loss. Muscle Nerve 20, 72-82.
Whiteman,E.L., Chen,J.J., Birnbaum,M.J. (2003). Platelet-derived growth factor (PDGF) stimulates glucose transport in 3T3-L1 adipocytes overexpressing PDGF receptor by a pathway independent of insulin receptor substrates. Endocrinology. 144, 3811-3820.
Wieczorek,D.F., Periasamy,M., Butler-Browne,G.S., Whalen,R.G., Nadal-Ginard,B. (1985). Co-expression of multiple myosin heavy chain genes, in addition to a tissue- specific one, in extraocular musculature. J Cell Biol 101, 618-629.
Wieczorek,D.F., Smith,C.W., Nadal-Ginard,B. (1988). The rat alpha-tropomyosin gene generates a minimum of six different mRNAs coding for striated, smooth, and nonmuscle isoforms by alternative splicing. Mol Cell Biol. 8, 679-694.
Witt,C.C., Burkart,C., Labeit,D., McNabb,M., Wu,Y., Granzier,H., Labeit,S. (2006). Nebulin regulates thin filament length, contractility, and Z-disk structure in vivo. EMBO J. 25, 3843-3855.
Wojtaszewski,J.F., Hansen,B.F., Urso,B., Richter,E.A. (1996). Wortmannin inhibits both insulin- and contraction-stimulated glucose uptake and transport in rat skeletal muscle. J Appl Physiol. 81, 1501-1509.
Yaffe,D., Saxel,O. (1977). Serial passaging and differentiation of myogenic cells isolated from dystrophic mouse muscle. Nature. 270, 725-727.
Yager,M.L., Hughes,J.A., Lovicu,F.J., Gunning,P.W., Weinberger,R.P., O'neill,G.M. (2003). Functional analysis of the actin-binding protein, tropomyosin 1, in neuroblastoma. Br J Cancer. 89, 860-863.
Yamauchi,T., Kamon,J., Waki,H., Murakami,K., Motojima,K., Komeda,K., Ide,T., Kubota,N., Terauchi,Y., Tobe,K., Miki,H., Tsuchida,A., Akanuma,Y., Nagai,R., Kimura,S., Kadowaki,T. (2001). The mechanisms by which both heterozygous peroxisome proliferator-activated receptor gamma (PPARgamma) deficiency and PPARgamma agonist improve insulin resistance. J Biol Chem. 276, 41245-41254.
Yang,Y., Makita,T. (1996). Immunocytochemical localization of desmin in human fetal skeletal muscle. J Electron Microsc (Tokyo). 45, 401-406.
Page 223
Young,P., Ferguson,C., Banuelos,S., Gautel,M. (1998). Molecular structure of the sarcomeric Z-disk: two types of titin interactions lead to an asymmetrical sorting of alpha-actinin. EMBO J. 17, 1614-1624.
Zador,E., Szakonyi,G., Racz,G., Mendler,L., Ver,H.M., Lebacq,J., Dux,L., Wuytack,F. (1998). Expression of the sarco/endoplasmic reticulum Ca(2+)-transport ATPase protein isoforms during regeneration from notexin-induced necrosis of rat soleus muscle. Acta Histochem. 100, 355-369.
Zhang,F., Lavan,B.E., Gregoire,F.M. (2007). Selective Modulators of PPAR-gamma Activity: Molecular Aspects Related to Obesity and Side-Effects. PPAR.Res. 2007:32696., 32696.
Zhang,H., Berg,J.S., Li,Z., Wang,Y., Lang,P., Sousa,A.D., Bhaskar,A., Cheney,R.E., Stromblad,S. (2004). Myosin-X provides a motor-based link between integrins and the cytoskeleton. Nat Cell Biol. 6, 523-531.
Zhao,C., Takita,J., Tanaka,Y., Setou,M., Nakagawa,T., Takeda,S., Yang,H.W., Terada,S., Nakata,T., Takei,Y., Saito,M., Tsuji,S., Hayashi,Y., Hirokawa,N. (2001). Charcot-Marie-Tooth disease type 2A caused by mutation in a microtubule motor KIF1Bbeta. Cell. 105, 587-597.
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Appendix A
The following table shows gene changes detected in the microarray of soleus muscle from 9d/89 knockout mice. Genes were grouped into ontology pathways by the GeneSifter program (VisX Labs). The column entitled ‘List’ indicates the number of genes deregulated in comparison to control samples, in the particular pathway, which is further specified into up- or down-regulated categories. The column entitled ‘Array’ shows the number of genes with the specific ontology assayed in the microarray. The Z-scores indicate whether the deregulation of each ontology occurs more or less frequently than expected. Positive numbers greater than 2.0 indicate that the genes in the pathway are deregulated more frequently than expected whereas a number less than -2.0 indicates the change is less frequently than expected. Z-score Pathway List Up Down Array Up Down Neuroactive ligand-receptor interaction 123 50 73 268 3.28 2.16 MAPK signaling pathway 90 30 60 264 -0.49 0.30 Regulation of actin cytoskeleton 76 19 57 192 -1.05 2.65 Calcium signaling pathway 74 25 49 166 1.10 2.40 Cytokine-cytokine receptor interaction 71 30 41 219 0.64 -1.20 Wnt signaling pathway 70 23 47 143 1.40 3.21 Cell adhesion molecules (CAMs) 63 25 38 121 2.84 2.55 Axon guidance 60 26 34 127 2.85 1.33 Jak-STAT signaling pathway 54 25 29 134 2.27 -0.10 Focal adhesion 52 15 37 181 -1.69 -0.51 Natural killer cell mediated cytotoxicity 51 17 34 109 1.06 2.36 Leukocyte transendothelial migration 46 18 28 111 1.27 0.84 T cell receptor signaling pathway 44 16 28 93 1.45 1.92 Insulin signaling pathway 43 12 31 133 -1.18 0.38 Tight junction 42 13 29 112 -0.23 1.02 Adherens junction 39 16 23 73 2.52 1.98 Cell cycle 38 9 29 107 -1.25 1.30 Purine metabolism 38 13 25 133 -0.91 -0.90 Hematopoietic cell lineage 35 13 22 77 1.23 1.41 Cell Communication 33 11 22 99 -0.37 0.06 Colorectal cancer 31 12 19 88 0.38 -0.09 Toll-like receptor signaling pathway 31 16 15 84 1.90 -0.92 Fc epsilon RI signaling pathway 30 14 16 75 1.69 -0.14 GnRH signaling pathway 30 9 21 91 -0.71 0.26 VEGF signaling pathway 30 12 18 71 1.19 0.69 Glycan structures - biosynthesis 1 29 10 19 98 -0.65 -0.63 Long-term potentiation 29 12 17 67 1.41 0.68 Pancreatic cancer 29 11 18 73 0.72 0.56 B cell receptor signaling pathway 28 8 20 64 0.04 1.81 ECM-receptor interaction 27 10 17 80 0.05 -0.16 Antigen processing and presentation 26 13 13 58 2.36 0.08 Apoptosis 26 8 18 81 -0.68 0.05 Complement and coagulation cascades 25 10 15 63 0.87 0.35 Pyrimidine metabolism 25 7 18 81 -1.02 0.05 Chronic myeloid leukemia 24 4 20 74 -1.83 1.06 Glioma 24 7 17 62 -0.25 1.04 Page 225
Phosphatidylinositol signaling system 24 11 13 72 0.77 -0.81 Hedgehog signaling pathway 23 7 16 53 0.20 1.45 Glycerophospholipid metabolism 22 8 14 73 -0.36 -0.58 Long-term depression 22 8 14 76 -0.48 -0.76 Metabolism of xenobiotics by cytochrome P450 22 9 13 52 1.10 0.53 TGF-beta signaling pathway 22 11 11 80 0.39 -1.80 Tryptophan metabolism 22 8 14 70 -0.23 -0.40 Arachidonic acid metabolism 21 10 11 65 0.76 -0.99 PPAR signaling pathway 20 10 10 67 0.65 -1.41 Glycan structures - biosynthesis 2 19 5 14 60 -0.95 0.25 Ribosome 19 6 13 82 -1.39 -1.36 Type I diabetes mellitus 19 7 12 40 1.00 1.23 Adipocytokine signaling pathway 18 5 13 70 -1.33 -0.70 Gap junction 17 4 13 87 -2.22 -1.60 Glycerolipid metabolism 17 5 12 46 -0.30 0.68 Glycolysis / Gluconeogenesis 17 8 9 52 0.68 -0.82 Inositol phosphate metabolism 17 8 9 51 0.74 -0.75 Type II diabetes mellitus 17 4 13 44 -0.66 1.22 Tyrosine metabolism 17 4 13 49 -0.89 0.77 Linoleic acid metabolism 16 10 6 41 2.36 -1.14 mTOR signaling pathway 16 6 10 50 -0.07 -0.34 Androgen and estrogen metabolism 15 3 12 35 -0.68 1.77 Notch signaling pathway 15 5 10 44 -0.19 0.12 Starch and sucrose metabolism 15 6 9 49 -0.02 -0.62 Butanoate metabolism 14 8 6 44 1.19 -1.34 Oxidative phosphorylation 14 6 8 110 -2.22 -3.78 Maturity onset diabetes of the young 13 4 9 25 0.56 1.70 Nicotinate and nicotinamide metabolism 13 4 9 39 -0.39 0.17 Olfactory transduction 13 3 10 28 -0.26 1.76 Sphingolipid metabolism 13 5 8 33 0.50 0.31 Neurodegenerative Disorders 12 7 5 32 1.65 -0.87 ABC transporters - General 11 3 8 39 -0.88 -0.22 Ether lipid metabolism 11 4 7 29 0.24 0.28 Fructose and mannose metabolism 11 3 8 49 -1.33 -0.96 Glycine, serine and threonine metabolism 11 4 7 46 -0.75 -1.11 Lysine degradation 11 7 4 47 0.54 -2.24 Pentose phosphate pathway 11 2 9 25 -0.66 1.70 Aminosugars metabolism 10 0 10 31 -2.10 1.39 Arginine and proline metabolism 10 4 6 51 -0.98 -1.77 Bile acid biosynthesis 10 2 8 34 -1.15 0.22 DNA polymerase 10 2 8 26 -0.72 1.09 gamma-Hexachlorocyclohexane degradation 10 5 5 22 1.49 0.08 Glutamate metabolism 10 2 8 31 -1.00 0.52 Glutathione metabolism 10 3 7 35 -0.68 -0.28 Histidine metabolism 10 4 6 33 -0.03 -0.53 N-Glycan biosynthesis 10 4 6 41 -0.50 -1.14 Taste transduction 10 1 9 29 -1.46 1.18 Alkaloid biosynthesis II 9 4 5 21 0.94 0.20 Dorso-ventral axis formation 9 2 7 27 -0.78 0.50 Fatty acid metabolism 9 2 7 40 -1.42 -0.69 Galactose metabolism 9 3 6 33 -0.57 -0.53 Glycan structures - degradation 9 4 5 25 0.56 -0.24 Phenylalanine metabolism 9 3 6 26 -0.12 0.14 Porphyrin and chlorophyll metabolism 9 0 9 23 -1.80 1.99 SNARE interactions in vesicular transport 9 4 5 31 0.10 -0.79 Valine, leucine and isoleucine degradation 9 2 7 46 -1.66 -1.11 Amyotrophic lateral sclerosis (ALS) 8 5 3 17 2.15 -0.43 Citrate cycle (TCA cycle) 8 3 5 26 -0.12 -0.34 Huntington's disease 8 4 4 27 0.40 -0.90 Limonene and pinene degradation 8 2 6 25 -0.66 0.24
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Propanoate metabolism 8 4 4 32 0.03 -1.30 Regulation of autophagy 8 4 4 25 0.56 -0.72 1- and 2-Methylnaphthalene degradation 7 0 7 21 -1.72 1.26 Benzoate degradation via CoA ligation 7 1 6 30 -1.50 -0.26 Chondroitin sulfate biosynthesis 7 3 4 17 0.67 0.15 Cysteine metabolism 7 3 4 19 0.46 -0.10 Folate biosynthesis 7 3 4 32 -0.51 -1.30 Glycosaminoglycan degradation 7 3 4 15 0.91 0.44 Glycosphingolipid biosynthesis - ganglioseries 7 2 5 16 0.02 0.90 Glycosphingolipid biosynthesis - neo-lactoseries 7 2 5 18 -0.16 0.60 Nitrogen metabolism 7 2 5 20 -0.32 0.33 O-Glycan biosynthesis 7 0 7 19 -1.64 1.57 Pantothenate and CoA biosynthesis 7 2 5 19 -0.24 0.46 Pyruvate metabolism 7 4 3 40 -0.45 -2.22 Selenoamino acid metabolism 7 3 4 28 -0.26 -0.99 Alanine and aspartate metabolism 6 2 4 31 -1.00 -1.23 Alzheimer's disease 6 2 4 21 -0.39 -0.33 beta-Alanine metabolism 6 2 4 23 -0.53 -0.53 C21-Steroid hormone metabolism 6 1 5 12 -0.42 1.65 Circadian rhythm 6 3 3 17 0.67 -0.43 Ethylbenzene degradation 6 1 5 15 -0.67 1.06 Glyoxylate and dicarboxylate metabolism 6 1 5 14 -0.59 1.24 Heparan sulfate biosynthesis 6 3 3 19 0.46 -0.65 One carbon pool by folate 6 3 3 16 0.78 -0.31 Riboflavin metabolism 6 2 4 17 -0.07 0.15 RNA polymerase 6 2 4 20 -0.32 -0.21 Aminophosphonate metabolism 5 2 3 14 0.22 -0.05 Basal transcription factors 5 1 4 30 -1.50 -1.15 Carbon fixation 5 1 4 23 -1.17 -0.53 Dentatorubropallidoluysian atrophy (DRPLA) 5 1 4 13 -0.51 0.77 Glycosphingolipid biosynthesis - lactoseries 5 1 4 10 -0.22 1.38 Methane metabolism 5 3 2 10 1.70 -0.15 Proteasome 5 1 4 29 -1.46 -1.07 Urea cycle and metabolism of amino groups 5 1 4 23 -1.17 -0.53 Ascorbate and aldarate metabolism 4 2 2 15 0.12 -0.81 Biosynthesis of steroids 4 1 3 16 -0.74 -0.31 Glycosphingolipid biosynthesis - globoseries 4 1 3 13 -0.51 0.10 Glycosylphosphatidylinositol(GPI)-anchor 4 2 2 20 -0.32 -1.30 biosynthesis Keratan sulfate biosynthesis 4 1 3 13 -0.51 0.10 Methionine metabolism 4 2 2 19 -0.24 -1.21 N-Glycan degradation 4 2 2 13 0.34 -0.58 Parkinson's disease 4 1 3 13 -0.51 0.10 Phenylalanine, tyrosine and tryptophan 4 2 2 10 0.74 -0.15 biosynthesis Prion disease 4 2 2 11 0.59 -0.30 Reductive carboxylate cycle (CO2 fixation) 4 2 2 11 0.59 -0.30 Sulfur metabolism 4 1 3 10 -0.22 0.61 Taurine and hypotaurine metabolism 4 1 3 8 0.02 1.06 Valine, leucine and isoleucine biosynthesis 4 1 3 9 -0.11 0.82 Vitamin B6 metabolism 4 1 3 12 -0.42 0.25 2,4-Dichlorobenzoate degradation 3 2 1 6 1.57 -0.31 Aminoacyl-tRNA biosynthesis 3 1 2 28 -1.41 -1.90 Bisphenol A degradation 3 1 2 17 -0.81 -1.02 Caprolactam degradation 3 1 2 13 -0.51 -0.58 Lysine biosynthesis 3 2 1 4 2.29 0.15 Nitrobenzene degradation 3 1 2 12 -0.42 -0.45 Pentose and glucuronate interconversions 3 0 3 12 -1.30 0.25 Retinol metabolism 3 0 3 5 -0.84 2.05 Thiamine metabolism 3 1 2 9 -0.11 0.02
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Atrazine degradation 2 0 2 4 -0.75 1.35 Benzoate degradation via hydroxylation 2 0 2 6 -0.92 0.67 Cyanoamino acid metabolism 2 1 1 5 0.52 -0.11 Fatty acid biosynthesis 2 0 2 5 -0.84 0.97 Nucleotide sugars metabolism 2 1 1 16 -0.74 -1.52 Streptomycin biosynthesis 2 2 0 9 0.90 -1.59 Terpenoid biosynthesis 2 0 2 5 -0.84 0.97 Tetrachloroethene degradation 2 1 1 10 -0.22 -0.92 1,1,1-Trichloro-2,2-bis(4-chlorophenyl)ethane 1 0 1 2 -0.53 0.96 (DDT) degradation Alkaloid biosynthesis I 1 0 1 5 -0.84 -0.11 C5-Branched dibasic acid metabolism 1 0 1 2 -0.53 0.96 D-Glutamine and D-glutamate metabolism 1 0 1 3 -0.65 0.47 Fatty acid elongation in mitochondria 1 0 1 9 -1.13 -0.79 Protein export 1 0 1 11 -1.24 -1.03 Stilbene, coumarine and lignin biosynthesis 1 1 0 6 0.32 -1.30 Styrene degradation 1 1 0 4 0.77 -1.06 Synthesis and degradation of ketone bodies 1 0 1 7 -0.99 -0.49
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Appendix B
The table below details the gene changes detected in the microarray of soleus muscle from Tm5/52 mice. An explanation of the table can be found in appendix A. Z-score Pathway List Up Down Array Up Down Neuroactive ligand-receptor interaction 126 75 51 268 4.35 2.60 MAPK signaling pathway 92 48 44 264 0.02 1.42 Cytokine-cytokine receptor interaction 78 45 33 219 0.96 0.58 Calcium signaling pathway 67 32 35 166 0.39 2.80 Regulation of actin cytoskeleton 65 29 36 192 -1.12 2.06 Cell adhesion molecules (CAMs) 62 43 19 121 5.05 0.63 Axon guidance 53 25 28 127 0.46 2.76 Wnt signaling pathway 50 25 25 143 -0.21 1.32 Focal adhesion 47 24 23 181 -1.75 -0.42 T cell receptor signaling pathway 45 26 19 93 2.49 1.89 Jak-STAT signaling pathway 42 25 17 134 0.16 -0.37 Natural killer cell mediated cytotoxicity 41 24 17 109 1.07 0.57 Leukocyte transendothelial migration 40 18 22 111 -0.53 1.88 Insulin signaling pathway 39 25 14 133 0.20 -1.10 Cell cycle 38 23 15 107 0.92 0.08 Tight junction 38 14 24 112 -1.57 2.39 Purine metabolism 37 24 13 133 -0.03 -1.36 Hematopoietic cell lineage 33 15 18 77 0.31 2.48 Adherens junction 29 13 16 73 -0.07 2.04 Colorectal cancer 28 15 13 88 -0.27 0.28 Pyrimidine metabolism 28 17 11 81 0.67 -0.05 Glycan structures - biosynthesis 1 27 15 12 98 -0.74 -0.44 GnRH signaling pathway 27 14 13 91 -0.69 0.15 Toll-like receptor signaling pathway 27 13 14 84 -0.64 0.78 Antigen processing and presentation 26 14 12 58 1.20 1.54 B cell receptor signaling pathway 26 12 14 64 0.13 1.90 Cell Communication 26 13 13 99 -1.31 -0.18 Fc epsilon RI signaling pathway 26 14 12 75 0.12 0.57 TGF-beta signaling pathway 25 16 9 80 0.44 -0.66 Apoptosis 24 14 10 81 -0.20 -0.37 Chronic myeloid leukemia 24 11 13 74 -0.74 0.96 Gap junction 24 12 12 87 -1.06 0.01 Hedgehog signaling pathway 24 18 6 53 3.01 -0.52 Adipocytokine signaling pathway 23 13 10 70 0.10 0.13 Glioma 23 12 11 62 0.25 0.92 Long-term depression 23 12 11 76 -0.54 0.18 Pancreatic cancer 23 11 12 73 -0.69 0.67 Tryptophan metabolism 23 13 10 70 0.10 0.13 VEGF signaling pathway 23 13 10 71 0.04 0.08 Glycerophospholipid metabolism 21 13 8 73 -0.07 -0.70 Long-term potentiation 21 10 11 67 -0.69 0.64 Phosphatidylinositol signaling system 21 14 7 72 0.29 -1.00 Arachidonic acid metabolism 20 11 9 65 -0.26 0.02 ECM-receptor interaction 20 13 7 80 -0.44 -1.32 Glycan structures - biosynthesis 2 20 9 11 60 -0.64 1.04 Type I diabetes mellitus 20 13 7 40 2.37 0.69 Complement and coagulation cascades 18 5 13 63 -2.12 1.60 Metabolism of xenobiotics by cytochrome 17 9 8 52 -0.16 0.34
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P450 Inositol phosphate metabolism 16 11 5 51 0.64 -0.83 mTOR signaling pathway 16 9 7 50 -0.02 0.05 Oxidative phosphorylation 15 7 8 110 -3.25 -2.01 PPAR signaling pathway 15 11 4 67 -0.37 -1.87 Olfactory transduction 14 6 8 28 0.45 2.28 Androgen and estrogen metabolism 13 7 6 35 0.29 0.58 Ribosome 13 5 8 82 -2.86 -1.06 Starch and sucrose metabolism 13 5 8 49 -1.45 0.53 Type II diabetes mellitus 13 9 4 44 0.40 -0.90 Fructose and mannose metabolism 12 7 5 49 -0.70 -0.73 Glutathione metabolism 12 9 3 35 1.17 -0.90 Glycerolipid metabolism 12 5 7 46 -1.29 0.29 Sphingolipid metabolism 12 7 5 33 0.46 0.23 Glycine, serine and threonine metabolism 11 7 4 46 -0.52 -1.00 Glycolysis / Gluconeogenesis 11 7 4 52 -0.88 -1.28 Pantothenate and CoA biosynthesis 11 8 3 19 2.72 0.26 Tyrosine metabolism 11 7 4 49 -0.70 -1.14 Lysine degradation 10 8 2 47 -0.20 -1.90 Maturity onset diabetes of the young 10 5 5 25 0.24 0.91 Basal transcription factors 9 5 4 30 -0.21 -0.07 Dorso-ventral axis formation 9 5 4 27 0.05 0.16 Linoleic acid metabolism 9 8 1 41 0.23 -2.12 Neurodegenerative Disorders 9 5 4 32 -0.37 -0.21 Nicotinate and nicotinamide metabolism 9 4 5 39 -1.28 -0.17 SNARE interactions in vesicular transport 9 9 0 31 1.58 -2.23 Taste transduction 9 3 6 29 -1.09 1.09 Valine, leucine and isoleucine degradation 9 4 5 46 -1.67 -0.57 ABC transporters - General 8 6 2 39 -0.45 -1.57 Bile acid biosynthesis 8 7 1 34 0.37 -1.84 Chondroitin sulfate biosynthesis 8 7 1 17 2.47 -0.94 Fatty acid metabolism 8 5 3 40 -0.93 -1.16 Galactose metabolism 8 8 0 33 0.91 -2.30 Glycosphingolipid biosynthesis - neo- 8 4 4 18 0.45 1.04 lactoseries Notch signaling pathway 8 4 4 44 -1.57 -0.90 O-Glycan biosynthesis 8 5 3 19 0.93 0.26 Amyotrophic lateral sclerosis (ALS) 7 5 2 17 1.21 -0.24 Butanoate metabolism 7 4 3 44 -1.57 -1.34 Citrate cycle (TCA cycle) 7 3 4 26 -0.88 0.24 Folate biosynthesis 7 3 4 32 -1.29 -0.21 Glycosphingolipid biosynthesis - lactoseries 7 3 4 10 0.98 2.41 Pentose phosphate pathway 7 4 3 25 -0.28 -0.26 Alkaloid biosynthesis II 6 4 2 21 0.11 -0.57 Alzheimer's disease 6 5 1 21 0.68 -1.20 Carbon fixation 6 4 2 23 -0.09 -0.71 Glycan structures - degradation 6 3 3 25 -0.80 -0.26 Glycosphingolipid biosynthesis - globoseries 6 4 2 13 1.18 0.17 N-Glycan biosynthesis 6 1 5 41 -2.62 -0.29 Nitrogen metabolism 6 3 3 20 -0.36 0.16 Pyruvate metabolism 6 3 3 40 -1.76 -1.16 RNA polymerase 6 2 4 20 -0.95 0.81 Selenoamino acid metabolism 6 6 0 28 0.45 -2.12 1- and 2-Methylnaphthalene degradation 5 2 3 21 -1.03 0.07 Arginine and proline metabolism 5 1 4 51 -3.02 -1.24 Benzoate degradation via CoA ligation 5 3 2 30 -1.16 -1.13 Cysteine metabolism 5 2 3 19 -0.86 0.26 DNA polymerase 5 3 2 26 -0.88 -0.90 Ether lipid metabolism 5 4 1 29 -0.61 -1.62 Glutamate metabolism 5 4 1 31 -0.76 -1.71
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Glycosaminoglycan degradation 5 2 3 15 -0.48 0.70 Glyoxylate and dicarboxylate metabolism 5 2 3 14 -0.37 0.83 Histidine metabolism 5 5 0 33 -0.45 -2.30 Huntington's disease 5 2 3 27 -1.45 -0.40 Prion disease 5 1 4 11 -0.78 2.18 Regulation of autophagy 5 3 2 25 -0.80 -0.84 Urea cycle and metabolism of amino groups 5 2 3 23 -1.18 -0.10 Alanine and aspartate metabolism 4 3 1 31 -1.23 -1.71 Aminoacyl-tRNA biosynthesis 4 3 1 28 -1.02 -1.57 Aminosugars metabolism 4 1 3 31 -2.16 -0.66 beta-Alanine metabolism 4 3 1 23 -0.64 -1.31 Biosynthesis of steroids 4 3 1 16 0.06 -0.87 Circadian rhythm 4 3 1 17 -0.05 -0.94 Dentatorubropallidoluysian atrophy (DRPLA) 4 3 1 13 0.46 -0.64 Ethylbenzene degradation 4 2 2 15 -0.48 -0.05 gamma-Hexachlorocyclohexane 4 3 1 22 -0.55 -1.26 degradation Heparan sulfate biosynthesis 4 2 2 19 -0.86 -0.41 Limonene and pinene degradation 4 2 2 25 -1.32 -0.84 Methionine metabolism 4 4 0 19 0.33 -1.75 Phenylalanine metabolism 4 2 2 26 -1.39 -0.90 Porphyrin and chlorophyll metabolism 4 1 3 23 -1.72 -0.10 Propanoate metabolism 4 1 3 32 -2.21 -0.72 Reductive carboxylate cycle (CO2 fixation) 4 3 1 11 0.79 -0.45 Valine, leucine and isoleucine biosynthesis 4 4 0 9 2.05 -1.20 Aminophosphonate metabolism 3 3 0 14 0.32 -1.50 Ascorbate and aldarate metabolism 3 3 0 15 0.19 -1.55 Bisphenol A degradation 3 3 0 17 -0.05 -1.65 C21-Steroid hormone metabolism 3 2 1 12 -0.13 -0.55 Glycosphingolipid biosynthesis - 3 0 3 16 -1.89 0.58 ganglioseries Keratan sulfate biosynthesis 3 1 2 13 -0.98 0.17 Nitrobenzene degradation 3 3 0 12 0.62 -1.39 One carbon pool by folate 3 3 0 16 0.06 -1.60 Parkinson's disease 3 1 2 13 -0.98 0.17 Pentose and glucuronate interconversions 3 2 1 12 -0.13 -0.55 Protein export 3 3 0 11 0.79 -1.33 Riboflavin metabolism 3 1 2 17 -1.31 -0.24 Sulfur metabolism 3 2 1 10 0.15 -0.35 Tetrachloroethene degradation 3 2 1 10 0.15 -0.35 Alkaloid biosynthesis I 2 2 0 5 1.27 -0.89 Benzoate degradation via hydroxylation 2 1 1 6 -0.09 0.21 Caprolactam degradation 2 1 1 13 -0.98 -0.64 Fatty acid biosynthesis 2 1 1 5 0.11 0.41 Fatty acid elongation in mitochondria 2 2 0 9 0.32 -1.20 Glycosylphosphatidylinositol(GPI)-anchor 2 2 0 20 -0.95 -1.79 biosynthesis Lysine biosynthesis 2 2 0 4 1.65 -0.80 Nucleotide sugars metabolism 2 2 0 16 -0.59 -1.60 Phenylalanine, tyrosine and tryptophan 2 1 1 10 -0.67 -0.35 biosynthesis Proteasome 2 2 0 29 -1.58 -2.16 Streptomycin biosynthesis 2 2 0 9 0.32 -1.20 Synthesis and degradation of ketone bodies 2 0 2 7 -1.25 1.14 Thiamine metabolism 2 1 1 9 -0.55 -0.23 1,1,1-Trichloro-2,2-bis(4- 1 0 1 2 -0.67 1.49 chlorophenyl)ethane (DDT) degradation 2,4-Dichlorobenzoate degradation 1 1 0 6 -0.09 -0.98 Atrazine degradation 1 1 0 4 0.36 -0.80 Biotin metabolism 1 1 0 9 -0.55 -1.20 C5-Branched dibasic acid metabolism 1 1 0 2 1.17 -0.56
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Methane metabolism 1 1 0 10 -0.67 -1.26 N-Glycan degradation 1 1 0 13 -0.98 -1.44 Peptidoglycan biosynthesis 1 0 1 2 -0.67 1.49 Stilbene, coumarine and lignin biosynthesis 1 1 0 6 -0.09 -0.98 Vitamin B6 metabolism 1 1 0 12 -0.88 -1.39
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Appendix C
The following results are from quantitative RT-PCR on cDNA samples from soleus muscles from Tm5NM1 transgenic and knockout mice and their respective controls. All genes were not significantly changed between transgenic/knockout mice and wild-type littermates. Statistic evaluations are shown here. All concentrations are expressed as
ng/µL.
Cofilin SUMMARY Groups Count Sum Average Variance Wild-type 2 0.1374 0.0687 2.05E-05 Tm5/52 3 0.2563 0.085433 0.000503
ANOVA Source of Variation SS df MS F P-value F crit Between Groups 0.000336 1 0.000336 0.981547 0.394853 10.12796 Within Groups 0.001027 3 0.000342
Total 0.001363 4
SUMMARY Groups Count Sum Average Variance Wild-type 3 0.15534 0.05178 2.85E-07 9d/89 3 0.29792 0.099307 0.001435 Fold change Up 1.9 fold Not significant ANOVA Source of Variation SS df MS F P-value F crit Between Groups 0.003388 1 0.003388 4.722132 0.095484 7.708647 Within Groups 0.00287 4 0.000718
Total 0.006258 5
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Cortactin SUMMARY Groups Count Sum Average Variance 1.25E- Wild-type 2 0.0182 0.0091 05 7.32E- Tm5/52 3 0.02952 0.00984 05
ANOVA Source of Variation SS df MS F P-value F crit Between 6.57E- Groups 6.57E-07 1 07 0.0124 0.918367 10.12796 Within Groups 0.000159 3 5.3E-05
Total 0.00016 4
SUMMARY Groups Count Sum Average Variance Wild-type 3 0.003337 0.001112 4.81E-07 9d/89 3 0.002733 0.000911 1.87E-08
ANOVA Source of Variation SS df MS F P-value F crit Between 6.08E- Groups 08 1 6.08E-08 0.243236 0.647728 7.708647 Within Groups 1E-06 4 2.5E-07
1.06E- Total 06 5
Myozenin SUMMARY Groups Count Sum Average Variance Column 1 3 0.2167 0.072233 7.85E-06 Column 2 3 0.4465 0.148833 0.006666
ANOVA Source of Variation SS df MS F P-value F crit Between Groups 0.008801 1 0.008801 2.637589 0.179685 7.708647 Within Groups 0.013348 4 0.003337
Total 0.022149 5
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Kcnma SUMMARY Groups Count Sum Average Variance Wild-type 3 0.2581 0.086033 0.002929 Tm5/52 3 0.10543 0.035143 0.002026
ANOVA Source of Variation SS df MS F P-value F crit Between Groups 0.003885 1 0.003885 1.568096 0.278705 7.708647 Within Groups 0.009909 4 0.002477
Total 0.013794 5
Myosin-8 SUMMARY Groups Count Sum Average Variance Wild-type 3 0.1875 0.0625 0.002932 Tm5/52 3 1.0238 0.341267 0.06061 Fold Change Up 5.46 fold Not Significant ANOVA Source of Variation SS df MS F P-value F crit Between Groups 0.116566 1 0.116566 3.668928 0.127941 7.708647 Within Groups 0.127085 4 0.031771
Total 0.243651 5
Triadin SUMMARY Groups Count Sum Average Variance Wild-type 3 2.339 0.779667 0.018105 9d/89 3 1.7827 0.594233 0.003782
ANOVA Source of Variation SS df MS F P-value F crit Between Groups 0.051578 1 0.051578 4.713 0.095709 7.708647 Within Groups 0.043775 4 0.010944
Total 0.095354 5
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Caveolin-2 SUMMARY Groups Count Sum Average Variance Wild-type 3 0.4021 0.134033 0.000941 9d/89 3 0.2582 0.086067 0.002048
ANOVA Source of Variation SS df MS F P-value F crit Between Groups 0.003451 1 0.003451 2.309387 0.203223 7.708647 Within Groups 0.005978 4 0.001494
Total 0.009429 5
Caveolin-3 SUMMARY Groups Count Sum Average Variance Wild-type 3 0.03803 0.012677 7.39E-06 9d/89 3 0.05085 0.01695 1.73E-05
ANOVA Source of Variation SS df MS F P-value F crit Between 2.74E- Groups 05 1 2.74E-05 2.216749 0.210752 7.708647 4.94E- Within Groups 05 4 1.24E-05
7.68E- Total 05 5
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