Cellular and Molecular Mechanisms Underlying Congenital Myopathy-Related Weakness

Digital Comprehensive Summaries of Uppsala Dissertations from the Faculty of Medicine 977

Cellular and Molecular Mechanisms Underlying Congenital Myopathy-related Weakness

JOHAN LINDQVIST

ACTA UNIVERSITATIS UPSALIENSIS ISSN 1651-6206 ISBN 978-91-554-8894-9 UPPSALA urn:nbn:se:uu:diva-219460 2014 Dissertation presented at Uppsala University to be publicly examined in Hedstrandssalen, Akademiska sjukhuset, ing 70, b.v., Uppsala, Wednesday, 16 April 2014 at 09:15 for the degree of Doctor of Philosophy (Faculty of Medicine). The examination will be conducted in English. Faculty examiner: Professor Anders Arner (Karolinska institutet).

Abstract Lindqvist, J. 2014. Cellular and Molecular Mechanisms Underlying Congenital Myopathy-related Weakness. Digital Comprehensive Summaries of Uppsala Dissertations from the Faculty of Medicine 977. 45 pp. Uppsala: Acta Universitatis Upsaliensis. ISBN 978-91-554-8894-9.

Congenital myopathies are a rare and heterogeneous group of diseases. They are primarily characterised by skeletal muscle weakness and disease-specific pathological features. They harshly limit ordinary life and in severe cases, these myopathies are associated with early death of the affected individuals. The congenital myopathies investigated in this thesis are nemaline myopathy and myofibrillar myopathy. These diseases are usually caused by missense mutations in genes encoding myofibrillar proteins, but the exact mechanisms by which the point mutations in these proteins cause the overall weakness remain mysterious. Hence, in this thesis two different nemaline myopathy-causing actin mutations and one myofibrillar myopathy-causing myosin-mutation found in both human patients and mouse models were used to investigate the cascades of molecular and cellular events leading to weakness. I performed a broad range of functional and structural experiments including skinned muscle fibre mechanics, small-angle X-ray scattering as well as immunoblotting and histochemical techniques. Interestingly, according to my results, point mutations in myosin and actin differently modify myosin binding to actin, cross-bridge formation and muscle fibre force production revealing divergent mechanisms, that is, gain versus loss of function (papers I, II and IV). In addition, one point mutation in actin appears to have muscle-specific effects. The presence of that mutant protein in respiratory muscles, i.e. diaphragm, has indeed more damaging consequences on myofibrillar structure than in limb muscles complexifying the pathophysiological mechanisms (paper II). As numerous atrophic muscle fibres can be seen in congenital myopathies, I also considered this phenomenon as a contributing factor to weakness and characterised the underlying causes in presence of one actin mutation. My results highlighted a direct muscle-specific up-regulation of the ubiquitin-proteasome system (paper III). All together, my research work demonstrates that mutation- and muscle-specific mechanisms trigger the muscle weakness in congenital myopathies. This gives important insights into the pathophysiology of congenital myopathies and will undoubtedly help in designing future therapies.

Keywords: skeletal muscle, skeletal muscle contraction, atrophy, nemaline myopathy, myofibrillar myopathy, myosin, actin

Johan Lindqvist, Department of Neuroscience, Clinical Neurophysiology, Akademiska sjukhuset, Uppsala University, SE-75185 Uppsala, Sweden.

ISSN 1651-6206 ISBN 978-91-554-8894-9 urn:nbn:se:uu:diva-219460 (http://urn.kb.se/resolve?urn=urn:nbn:se:uu:diva-219460)

To my family

List of Papers

This thesis is based on the following papers, which are referred to in the text by their Roman numerals.

I Lindqvist J., Penisson-Besnier I., Iwamoto, H., Li M., Yagi N., Ochala J. (2012). A myopathy-related actin mutation increases contractile function. Acta Neuropathol 123(5): 739- 746 II Lindqvist J., Cheng A.J., Renaud G., Hardeman E.C., Ochala J. (2013). Distinct underlying mechanisms of limb and respiratory muscle fiber weaknesses in nemaline myopathy. J Neuro- pathol Exp Neurol 72(6): 472-481 III Lindqvist J., Hardeman E.C., Ochala J. (2013). Muscle- specific up-regulation of the ubiquitin-proteasome pathway in a mouse model of nemaline myopathy (in manuscript) IV Lindqvist J., Iwamoto H., Blanco G., Ochala J. (2013). The fraction of strongly bound cross-bridges is increased in mice that carry the myopathy-linked myosin heavy chain mutation MYH4(L342Q). Dis Model Mech 6(3): 834-840

Reprints were made with permission from the respective publishers.

Related papers

Kurapati R, McKenna C, Lindqvist J., Williams D, Simon M, LeProust E, Baker J, Cheeseman M, Carroll N, Denny P, Laval S, Lochmüller H, Ochala J, Blanco G. (2012) Myofibrillar myopathy caused by a mutation in the motor domain of mouse MyHC IIb. Hum Mol Genet 21(8):1706-24

Contents

Introduction ...... 11 Skeletal muscle ...... 11 Sarcomeres and myofilaments ...... 11 Thin filaments ...... 13 Thick filaments ...... 15 Contraction ...... 16 Muscle protein synthesis and degradation ...... 18 Protein synthesis ...... 18 Protein breakdown ...... 19 Congenital myopathies ...... 21 Nemaline myopathy ...... 21 Myofibrillar myopathy ...... 22 Aims of present investigation ...... 23 Material & Methods ...... 24 Human patients (Paper I) ...... 24 Animals (Paper II, III and IV) ...... 24 Muscle isolation (Paper I, II, III, IV) ...... 25 Contractile measurements of single fibres (Paper I, II and IV) ...... 25 In vitro motility assay (Paper I) ...... 26 Histochemistry (Paper I and III) ...... 26 Myosin heavy chain isoform expression (Paper I, II, III and IV) ...... 26 Myosin, actin and total protein quantification (Paper I, II, III and IV) ...... 27 Electron-microscopy (Paper I, II and IV) ...... 27 Immunoblotting (paper III) ...... 27 Small-angle X-ray Scattering (paper I and IV) ...... 27 Actomyosin modelling (Paper I and II) ...... 28 Intact fibre preparation and measurements (Paper II) ...... 28 Statistics (Paper I, II, III and IV) ...... 28 Results and discussion ...... 29 Paper I ...... 29 Paper II ...... 30 Paper III ...... 30 Paper IV ...... 31

Conclusion ...... 33 Future therapies ...... 35 RNA interference ...... 35 Inhibition of myostatin ...... 35 Improving efficiency of contraction ...... 36 Acknowledgments...... 37 References ...... 39

Abbreviations

4E-BP1 Eukaryotic translation initiation factor 4E binding protein ADP Adenosine diphosphate ATP Adenosine triphosphate CSA Cross-sectional area DNA Deoxyribonucleic acid dsRNA Double-stranded RNA EDL Extensor digitorum longus EGTA ethylene glycol tetraacetic acid eIF2B Eukaryotic initiation factor 2B FDB Flexor digitorum brevis FoxO Forkhead box O GSK3β Glycogen synthase kinase 3β IGF Insulin-like Growth Factor IRS Insulin receptor substrate KBTBD13 Kelch repeat and BTB (POZ) domain containing 13 MM Myosin meridional mRNA messenger RNA mTOR Mammalian Target of Rapamycin mTORC1 and 2 mTOR complex 1 and 2 MuRF1 Muscle-specific Ring Finger protein1 MFM Myofibrillar myopathy MyHC Myosin Heavy Chain MyLC Myosin Light Chain NM Nemaline myopathy PCR Polymerase Chain Reaction PDK1 Protein-dependent Kinase 1 PI3K Phosphoinositide 3-kinase PIP3 Phosphatidylinositol triphosphate PKB Protein Kinase B RISC RNA-induced silencing complex RNA Ribonucleic acid RNAi RNA interference rRNA Ribosomal RNA RTK Receptor Tyrosine Kinase

SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis siRNA Small interfering RNA SMAD Composite name of Sma and MAD proteins SMYD SET and MYND domain Tm Tropomyosin TnC Troponin C TnI Troponin I TnT Troponin T TRiC TCP-1 ring complex tRNA Transfer RNA UPS Ubiquitin-proteasome system UV Ultraviolet

Introduction

Skeletal muscle The skeletal muscle is the largest organ in the human body, constituting 40- 45 % of the body weight. In the human body, there are over 600 anatomi- cally defined skeletal muscle groups and most skeletal muscles are attached to bones and transverse over joints. They are responsible for breathing, movement and posture, but they also fulfil important functions as secretory organs and storage for energy and amino acids1-3. The body contains different muscles with different properties to meet their functional demands. The skeletal muscle system is composed of hun- dreds of motor units, which are characterised by motor neurons and clusters of muscle fibres having different biochemical and mechanical properties. The muscle fibres can be divided into slow- and fast-twitch fibres in regard to their contractility4. In addition, skeletal muscle has very high plasticity to be able to adapt to different functional stimuli and demands, such as endur- ance and resistance exercise, immobilisation, denervation and disease conditions. The adaptations usually take place at the cell level, i.e. muscle fibre level, and involve changes in fibre size and/or muscle fibre type5, 6. Muscle fibres are multinucleated cells that are derived from the fusion of differentiated single muscle cells, called myotubes. The origin of the myotubes is the myocytes that are formed from the dermomyotome, the dorsal epithelial part of the mesodermal somites during embryogenesis3, 7.

Sarcomeres and myofilaments The most abundant structures inside a muscle fibre are the parallel myofi- brils that longitudinally span the entire fibre. Each myofibril has a diameter of 1-2 µm and is separated from each other by mitochondria, sarcoplasmic reticulum and transverse tubular systems8. The myofibril is made up of re- petitive units called sarcomeres9. The sarcomere has a length of 2-3 µm and is composed of interdigitating thin and thick filaments, which are arranged in a hexagonal pattern. It is responsible for contraction and force production10. The lateral boundaries of the sarcomeres are the Z-discs, which are the thin filament anchoring network that runs perpendicular to the fibre axis11. Light microscopy-studies of skeletal muscle have revealed the characteristic stri-

11 ated pattern, with alternating dark and light bands. This striated pattern arises from the precise protein arrangement of thin and thick filaments that are crucial for proper skeletal muscle function. The dark bands are built up by the thick filaments with overlapping thin filaments. This area is called the A- band. The light non-overlapping region of the thin filaments is denoted I- band. In the middle of the light I-band, the thin Z-disc is observed, while in the central part of the A-band, the brighter H-zone is found. The H-zone only contains the thick filaments. The parallel thick filaments are held together by the M-line found in the middle of the H-zone8, 11, 12 (see figure 1). The actin part of the filament joins the Z-disc with the barded end through interaction with α-actinin and CapZ13, 14.

Figure 1. Top: Basic organisation of sarcomeres with myosin-containing thick and actin-containing thin filaments. The thin filaments are cross-linked with α-actinin and anchored in the Z-disc. The thick filaments are centrally located in the sarcomere. The myosin heads, or cross-bridges, on the thick filament interact with actin during activation. Bottom: Electron microscopy photograph reveal the ultrastructure of sarcomeres. © 2008 Ottenheijm et al. Respiratory Research 2008 9:1215 under Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0)

The cell membrane of the muscle fibre is called sarcolemma and has a central role in the excitation-contraction coupling16. Furthermore, the sarcomeres are attached to the sarcolemma and the extracellular basement membrane through the costameres and Z-disc, which transfer the produced force and shortening to the muscle cell membrane10. It is also important for avoiding contraction-induced damage17.

12 Thin filaments The basic structural unit in the thin filament consists of actin, tropomyosin and troponin. Troponin and tropomyosin are the main regulatory proteins that control actin-myosin interactions, while actin is the main component of the thin filaments11. The ratio between actin monomers, tropomyosin and troponin complex is 7:1:118. Tropomyosin and troponin starts to bind to the thin filament 40-60 nm from the edge of the Z-disc13.

Figure 2. Structure of the actin monomer and α-helical actin filament. Published 2014-02-10 at Genetics Home Reference website, a service by U.S: National Library of Medicine. National Library of Medicine (US). Genetics Home Reference [Inter- net]. Bethesda (MD): The Library; 2014 Feb 10. [Illustration] Gene therapy using an adenovirus vector; [cited 2014 Feb 19]; [about 1 screen]. Available from: http://ghr.nlm.nih.gov/handbook/illustrations/actin

Actin Actin is a ubiquitous and highly evolutionary conserved protein. The human genome has six actin genes that encode six functional actin isoforms. Four isoforms are found in skeletal, cardiac or smooth muscle and three of them are present in skeletal muscle during development. During development the expression of actin isoforms change, in the following order, from cytoskeletal γ-actin to cardiac α-actin and finally to skeletal α-actin. Most important

13 are ACTA1 and ACTC1 genes, respectively encoding skeletal muscle α-actin and its cardiac counterpart19-21. A single polypeptide chain of 375 amino acids forms the mature actin monomer22. The molecular mass is 42 kDa23, 24. In the eukaryotic cell actin exists in two forms, globular G-actin and filamentous F-actin. G-actin spon- taneously forms double-stranded helical filaments under physiological salt concentration25. X-ray diffraction together with the crystal structure of G- actin has given important structural information about actin monomers and filaments. The actin monomer looks like a rectangular prism with the dimen- sions 55 Å x 55 Å x 35 Å. It has two domains, the inner and outer, that tradi- tionally are divided into four subdomains, named subdomain 1-426. The inner and outer domains are separated by a cleft, which holds an ATP or ADP together with a Ca2+ or Mg2+ ions26, 27. In the filamentous form the inner domains of actin monomers are located close to the filament axis, while the smaller subdomains 1 and 2 are positioned on the outside and are available for interaction with myosin. When G-actin assemblies into F-actin, the domains of the actin monomers twist in relation to each other helping further elongation and function of the filament. Each actin monomer is connected to four other actin monomers. These are the preceding and following actin monomers in the same long pitch helix and two actins on the other strand. The complete structure of the actin filament is two twisted α-helices of actin monomers and the two actin strands has a cross-over repeat of about 36 nm (Figure 2)13, 26-28. Functional active actin requires a special folding pathway. This pathway starts with the chaperone prefoldin (GimC) that transfers the actin polypeptide from the ribosomes to cytosolic chaperonins, which helps the actin polypeptide to fold properly into an active monomer. These monomers po- lymerise into a filament, where TCP-1 Ring Complex (TRiC) together with other chaperones plays an important part29, 30.

Tropomyosin Tropomyosin (Tm) binds F-actin and troponin. The tropomyosin molecule is ~40 nm long and lies along the actin filament end to end in a cable-like fash- ion. The structure of tropomyosin is two α-helices that form an elongated coiled-coil with hydrophobic residues and salt bridges holding the helices together. Each tropomyosin molecule binds seven actin monomers and it is electrostatically bound to the actin filament at two points. It serves an important function together with the troponin complex to modulate myosin’s ac- cess to the actin filament31, 32. In mammals, four different tropomyosin genes exist, which express around forty different isoforms due to alternative promotors and mRNA processing. The TPM1 and TPM2 genes are expressed in skeletal muscle and translate into α- and β-tropomyosin respectively33. In fast muscle from rabbit it has been estimated that tropomyosin represent 3% of the total protein34.

14 Troponin Troponin is a complex composed of three subunits; the Ca2+ binding troponin C (TnC), the inhibitory troponin I (TnI) and the tropomyosin-binding troponin T (TnT)18. The troponin complex covers ~2/3 of the tropomyosin length31. Studies of the TnC crystal structure have revealed that its N- terminal part is responsible for the regulatory Ca2+ binding of TnC, while the C-terminal domain is important for interaction with the other proteins in the Troponin complex. Binding of Ca2+ to these sites cause conformational changes in the N-terminal domain that enables strong interaction between TnC and TnI18, 35. The C-terminal element of TnI inhibits actin through two regions in the molecule. The primary role of TnT is to function as a link between the TnC, TnI, Tm and actin, but it also augments the inhibition effect of TnI as well as the activity of actomyosin ATPase18, 31. In the absence of Ca2+, TnI is strongly bound to actin and weakly bound to TnC, while the presence of Ca2+ greatly increases the TnC’s affinity for TnI causing actin- TnI dissociation36. This process is of crucial importance for proper regulation of skeletal muscle contraction.

Thick filaments The thick filaments are multi-protein complexes with myosin as the most important protein. 200-400 myosin molecules form each of the thick fila- ments19. The proper function of the thick filaments depends on correct folding of the myosin molecule and assembly of the thick filaments, which requires a number of chaperones, including the SMYD1 and UNC4529, 37, 38.

Myosin Myosin is a motor protein that converts chemical energy into mechanical work. It is the only known actin-based motor protein. The class II myosin includes both muscle myosin and non-muscle myosin. The muscle myosin can be further divided into smooth and sarcomeric myosin39-41. Until now, eleven different sarcomeric myosin heavy chain (MyHC) genes have been identified in mammalian genomes, which are believed to be the total number of MyHC genes present in mammals42. The most common MyHCs in skeletal muscles are the slow MyHC (type I) and the three fast MyHCs (type IIa, IIx and IIb). Type IIa is the slowest and type IIb is the fastest with IIx as an intermediate of the fast isoforms43. The genes that encode MyHCs are located on chromosomes 17 and 11 in humans and mice respectively. See table 1 for genes and their corresponding MyHC isoform44-47. Myosin II is composed of two heavy chains and four light chains. The molecular mass of the two heavy chains is roughly 220 kDa and the complete assembled myosin molecule weighs around 520 kDa48. The long C- terminal portions of the heavy chains dimerise and form an α-helical coiled-

15 coil tail. The tails from many myosin dimers pack together to form the back- bone of the thick filaments49. The N-terminal part of the myosin heavy chain has the structure of a large globular head, which is able to bind actin and hydrolyse ATP50. After enzymatic cleavage, this part is known as subfragment 1 (S1). S1 contains the first 843 amino acid residues of the heavy chain49. A long α-helix extends from the thick globular part of S1. This ex- tension is known as the lever arm, while the globular head constitutes the motor domain of myosin51. Two myosin light chains (MyLCs), one regulatory and one essential, are also attached to the S1. The MyLCs modulate myosin activity and stiffen the lever arm48, 49.

Table 1. Skeletal and cardiac muscle myosin heavy chain genes with corresponding protein and location of expression4, 44-46. Gene Protein Human expression MYH6 Α-cardiac MyHC Atria MYH7 MyHC I Slow type I fibre and ventricles MYH2 MyHC IIa Fast type IIa fibres MYH1 MyHC IIx Fast type IIx fibres MYH4 MyHC IIb Not expressed MYH3 Embryonic MyHC Fetal period and muscle regeneration MYH8 Perinatal MyHC Fetal period and muscle regeneration MYH13 Extraocular MyHC Extraocular muscles

The motor domain can be further divided into an actin-binding region, a nucleotide-binding pocket and a converter region. The actin-binding region consists partly of the upper and lower 50-k subdomains. These subdomains are separated by a cleft that runs from the actin-binding part to the nucleotide-binding pocket52.

Contraction Skeletal muscle contraction involves a cascade of events starting at the neuromuscular junction by the release of acetylcholine into the synaptic cleft and binding to the acetylcholine receptor, followed by propagation of an action potential along the muscle fibre membrane, activating dihydropyri- dine receptors which initiates the release of Ca2+ from the sarcoplasmic reticulum notably by ryanodine receptors53. The released Ca2+ binds to TnC, which causes the Troponin complex together with Tm to slightly shift position along the thin filament. This shift reveals myosin head attachment sites on the actin filament. The opening of these attachment sites activates the myosin heads to go from a weak binding state to a strong binding state, which will start conformational changes in the myosin head and lever arm swing. The transition to the strong binding state

16 pushes tropomyosin aside, which allows exposure of more myosin binding sites on other actin monomers and attachment of more myosin heads54, 55. This cycling movement induces a sliding motion of the thin and thick filaments in opposite directions resulting in contraction i.e. shortening of the muscle48, 49 (See Figure 3).

Figure 3. Thin filament movement and actomyosin interaction during contraction. The initial binding of myosin head to skeletal α-actin is weak and transient [1]. The release of Pi induces a stronger myosin-actin interaction as well as a tilting of the converter and light-chain-binding regions [2]. At the end of the tilting, the release of ADP provokes an even stronger myosin-actin interaction [3]. This conformation ends rapidly when ATP binds to the myosin head. Therefore, the myosin head de- taches and the converter and light-chain-binding portions return to their initial orientation [4]. ATP is then hydrolysed [5] resulting in myosin head rebinding [1]. ©Julien Ochala with permission.

17 From biochemical and structural studies, it is known that Troponin-Tm- actin filaments can exist in three different states; blocked, closed and open. When Ca2+ is absent all myosin-binding sites on the actin filament are cov- ered by the Troponin-Tm-complex, corresponding to the blocked state. The closed state occurs when some of the myosin-binding sites are available due to presence of Ca2+ and the shift of the Troponin-Tm complex. Attachment of myosin heads to these sites forces exposure of all myosin binding sites and allows the transition from the weakly to strongly bound cross-bridges. This is the open state18. The ATPase activity of myosin causes ATP to be hydrolysed into ADP and inorganic phosphate, Pi, which exist in a myosin-ADP.Pi-complex. In the weak binding state, the pre-power stroke state, myosin binds to actin with parts of the upper and lower 50k domains. These domains are separated by a cleft that closes on transition to the strong binding state56. The closure leads to a large rotation of the converter subdomain of the motor domain, which is connected to the lever arm. This rotation builds up strain and strain release cause a ~10 nm axial move of the lever arm and sliding of the thin filament. In addition, closure of the cleft allows the Pi and later on the ADP to escape from the nucleotide binding pocket. However, the precise mechanism for this is still unclear, but it results in the power stroke and the myosin rigor state52, 54. ATP is rapidly rebound to the nucleotide-binding pocket, which leads to opening of the 50k cleft and dissociation of myosin and actin, so a new contraction cycle can start49, 50. During isometric contraction in physiological conditions, it has been estimated that the fraction of myosin heads attached to actin is 30-40%57. The force developed by muscle fibres depends on the number of myosin heads attached to actin and the amount of them that are in the strong binding state. The produced force is temperature-dependent, due to that the amount of myosin heads in the strong binding state depend on temperature58.

Muscle protein synthesis and degradation The size and function of skeletal muscle is controlled by anabolic and cata- bolic signals that regulate the rates of protein synthesis and degradation, which determines the net protein balance. When there is a surplus of protein synthesis, muscle hypertrophy occurs, while excess protein degradation causes muscle atrophy.

Protein synthesis A very important pathway that regulates skeletal muscle growth is the insulin-like growth factor 1-AKT/protein kinase B (IGF1-Akt/PKB) pathway. AKT signalling is initiated by the binding of IGF1 or insulin to the IGF1

18 receptor, a receptor tyrosine kinase (RTK) that is autophosphorylated, which generates docking sites for insulin receptor substrate (IRS). IRS is also phosphorylated by the IGF1 receptor. The phosphorylated IRS recruits and forms a docking site for phosphoinositide 3-kinase (PI3K). Phosphatidy- linositol (3, 4, 5)-triphosphate (PIP3) is generated by PI3K and functions as an anchor for the two kinases PDK1 and AKT. This leads to phosphorylation and activation of AKT. AKT, in turn, phosphorylates a number of different targets, including mTOR, FoxO and GSK3β59-63. mTOR is found in two complexes, mTORC1 and mTORC2. Today, mTORC1 is believed to be made up of at least 6 proteins while mTORC2 consists of seven proteins. Growth factors, stress, energy status, oxygen and amino acids affect the activity of mTORC1. mTORC1 directly phosphorylates two proteins that promote protein synthesis, namely the eukaryotic translation initiation factor 4E binding protein (4E-BP1) and S6 kinase 1 (S6K1)64. S6K1 determines muscle fibre size65, 66. mTORC1 is also important for the transcription of genes regulating mitochondrial metabolism, rRNAs and tRNAs61. Regulation of mTORC2 is more mysterious with its insensitivity to nutrients, but it seems to respond to growth factors through PI3K. It is important for cell survival and proliferation64. GSK3β modulate protein synthesis through inhibiting the translation initi- ating eIF-2B protein. Phosphorylation of GSK3β by AKT stops this inhibition, which leads to muscle hypertrophy as well as enhanced myoblast dif- ferentiation62, 67.

Protein breakdown Muscle atrophy occurs when protein synthesis is either decreased and/or the protein degradation is increased68, 69. Denervation, unloading, immobilisation, bed rest, various diseases (e.g. cancer or diabetes), fasting, sepsis or disuse can cause skeletal muscle protein degradation and consequently muscle atrophy70-74. Three important degradation pathways are the ubiquitin- proteasome system (UPS), the autophagy-lysosomal pathway and the calpain system. It has been suggested that the UPS targets short-lived or abnormally folded proteins, while the autophagy-lysosomal pathway selectively targets long-lived macromolecules and organelles75. Numerous studies indicate that the UPS is the major regulator of protein degradation in skeletal muscles during various conditions76, 77.

Ubiquitin-proteasome system Ubiquitin is a polypeptide consisting of 76 amino acid residues. A polymer of ubiquitin attached to a protein marks that protein for degradation by the 26S proteasome78. Polyubiquitination of proteins depends on a sequential action mechanism with activating enzyme (E1), ubiquitin-conjugating enzymes (E2) and ubiquitin-protein ligases (E3)76. E1 activates the C-terminal

19 of ubiquitin making it ready for transfer. The E2 ubiquitin-conjugating enzymes transiently carry the activated ubiquitin until an E3 ligase transfer the activated ubiquitin from the E2-enzyme to the targeted protein78. The polyubiquitin chains on the degradation-targeted protein are then recognised by the regulatory subunit of the proteasome79. After recognition, the proteins are unfolded and transported into the proteasome by an ATP-dependent mechanism. The 26S proteasome is built up by the 20S catalytic core and two 19S regulatory caps. Four heptameric rings makes up the 20S catalytic core. The rings are responsible for both structure and chymotrypsin-, trypsin- and caspase-like activity. This activity cleaves the tagged proteins into short oligopeptides that are further degraded by tripeptidyl-peptidase II and other exopeptidases77. Two important E3 ligases in skeletal muscle are MuRF1 and Atrogin-180. MuRF1 and Atrogin-1 are only expressed in skeletal muscle and their expression is increased during atrophic conditions81. During such conditions, the activity of AKT is decreased. It, in turn, promotes dephosporylation of FoxO family members and their translocation to the cell nucleus, which leads to an up-regulated expression of various proteolytic systems, including multiple atrophy-related genes (atrogenes) MuRF1 and Atrogin-162, 68. In addition, cytokines, hormones and myostatin can affect the UPS76.

Autophagy-lysosomal pathway The autophagy-lysosomal protein degradation pathway uses lysosomes for degradation and recycling of cellular components. Macro-autophagy occurs when a distinct region of the cytoplasm with organelles, protein complexes and cytosol is sequestered within a double-membrane cytosolic vesicle. This vesicle is called an autophagosome, which ultimately fuses with the lysosome creating the autolysosome. The inner membrane of the autolysosome is broken down and its cargo degrade by hydrolases and other enzymes82. In contrast, the lysosome itself invaginates and engulfs a portion of the cytoplasm in micro-autophagy. During this process the lysosome goes through complex structural rearrangements to create an autolysosome-like vesicle that is degraded together with its cargo75. In chaperone-mediated autophagy, proteins with a specific pentapeptide sequence are, during nutrient deprivation, recognised by cytosolic chaperones and delivered to the surface of the lysosome, where the protein is transferred across the lysosomal membrane and then is degraded inside the ly- sosome83. The debris that is created by the degradation is later transported out of the lysosome into the cytoplasm and reused by the cell82. Interestingly, apoptosis can inhibit the autophagy programme by caspase-mediated diges- tion of autophagy proteins84.

20 The calpain system A third proteolytic system in skeletal muscles is the calpains. They are intra- cellular Ca2+-activated cysteine proteases. It is suggested that calpains function as initiators of myofibrillar degradation, because they are believed to cleave the proteins that are responsible for anchoring the thick and thin filament proteins in the sarcomeres. The release of these proteins from the sarcomeres makes the contractile proteins available for sequential degradation by the UPS and the autophagy-lysosomal pathway. The calpains are regulated by Ca2+ binding, which activates the calpains through protein rearrangements that lead to formation of the active catalytic site. In addition to Ca2+ regulation, calpastatin exists. It is an endogenous protein inhibitor of calpains85-87. Calpains are expressed in all vertebrate cells and three isoforms (1-3) of calpain are found in skeletal muscles86.

Congenital myopathies Congenital myopathies are a rare and heterogeneous group of genetic diseases that usually lead to skeletal muscle weakness and hypotonia. These diseases are in most cases present from birth or have an early onset and have distinct morphological characteristics in common. The diseases are caused by a single mutation in a wide range of genes; however there are complex relationships between the mutation and the type of disease88, 89. Nemaline myopathy (NM) and myofibrillar myopathy (MFM) are the two congenital myopathies investigated in this thesis.

Nemaline myopathy Nemaline myopathy is the most common non-dystrophic congenital myopathy with an incidence of 1:50 00090. NM is characterised by generalised skeletal muscle weakness, hypotonia and the presence of rod-like protein accumulations, i.e. nemaline bodies (rods) in fibres. The nemaline bodies stain red with modified Gomori trichrome stain. Predominance of type I fibres is also a common feature. Usually, there is no cardiac involvement91-93. Six clinical categories of NM exist: (1) severe congenital form, with patients lacking spontaneous movement or respiration at birth, or have frac- tures or severe contractures at birth; (2) intermediate congenital form includes spontaneous moving and breathing newborns, which fail to achieve ambulation or respiratory independence; (3) typical congenital form with patients displaying typical distribution of muscle weakness, delay in reach- ing ambulation milestones and a slowly progressive or non-progressive course; (4) mild NM with childhood onset; (5) adult onset NM; (6) other forms of NM with unusual associated features. The categories have multiple pathological and clinical overlaps94. NM is caused by recessive, dominant or

21 de novo autosomal mutations in eight genes mainly encoding thin filament proteins. Thus, TPM2 (β-tropomyosin), TPM3 (slow α-tropomyosin), TNNT1 (slow skeletal muscle troponin T), NEB (nebulin), CFL2 (cofilin-2), KBTBD13 (Kelch repeat and BTB domain-containing protein 13), KLHL40 (Kelch-like family member 40) and ACTA1 (skeletal α-actin) have been identified as causative genes91, 92. Mutations in nebulin are responsible for over 50 % of all cases of nemaline myopathy, while ACTA1 mutations cause up to 25% of the cases and over half of the severe cases. Today, over 200 disease-causing mutations have been identified in ACTA191, 93. No curative treatment exists for NM today, but a small clinical study and animal experiments have indicated that dietary supplementation of amino acids can alleviate the pathology and weakness that are associated with NM93, 95, 96.

Myofibrillar myopathy Myofibrillar myopathies constitute another rare group of adverse congenital myopathies. MFMs are defined by skeletal muscle weakness that initially starts in the lower extremities, and at later stages affect all limbs, respiratory and masticatory muscles97. The characteristic features of MFMs are the presence of protein aggregates and disintegration of the myofibrillar networks. Typically, the disintegration starts at the Z-disc. In addition, histological studies of patients have revealed fibres with abnormal size, large vesicular nuclei, necrotic and regenerating patterns as well as increased fibrosis. In these altered fibres, amorphous, granular or hyaline deposits with variations in size, shape and colour can also be observed. MFMs are related to mutations in DES (desmin), CRYAB (αB-crystalline), MYOT (myotilin), ZASP (Z- disc associated protein), FLNC (filamin C) BAG3 and TTN (titin)98, 99. Nev- ertheless, a large number of cases remains with no identified genetic cause 97, but candidate genes exist including the genes encoding the motor protein myosin.

22 Aims of present investigation

Most of the mutations leading to NM or MFM are typically missense mutations that change a single DNA nucleotide, which results in the substitution of one amino acid in the protein. An exchange of an amino acid residue affects the physicochemical properties of the protein and due to the strong structure-function relationship in skeletal muscle this affects the muscle function. The mechanisms by which such minor changes result in contractile dysfunction and weakness remain in many cases unknown. Therefore the aim of this thesis is to give insights into these deregulations (especially when thin and thick filament proteins are mutated) in order to design potential therapeutic interventions. Thus, I performed studies using various models (patient biopsy samples and transgenic animal models) and approaches.

Specific aims:  Is general skeletal muscle weakness associated with a decreased force production in single skeletal muscle fibres?  How does a single amino acid substitution cause changes in the contraction mechanism?  Do different mutations deregulate the contraction mechanism in the same way?  Investigate underlying molecular pathways in atrophic muscle fibres.

23 Material & Methods

Human patients (Paper I) An open muscle biopsy was performed under local anaesthesia from the deltoid muscle of a 46-year old male patient diagnosed with NM. Control biopsies were obtained by using a percutaneous conchrotome from tibialis anterior and vastus lateralis muscles of seven healthy age-matched subjects. A local ethics committee at Uppsala University approved the protocol and the experiments were carried out according to the guidelines of the Declara- tion of Helsinki.

Animals (Paper II, III and IV) Two different animal models were used in this thesis; one model for NM and one for MFM. The MFM-model was created during an N-ethyl, N- nitrosourea (ENU) induced mutagenesis screen set up for the identification of recessive mutant alleles in the C57BL6/J mouse strain. The model developed hindlimb paralysis by postnatal day 13 and a genetic screening local- ised a mutation in exon 11 of the MYH4 gene. Next generation sequencing had revealed a nucleotide mutation that lead to a leucine to glutamine exchange in the motor protein myosin heavy chain IIb (L342Q mutation). Only the skeletal muscles displayed pathological changes in the model100. To study the molecular consequences of NM and compare it to human patient, a mouse model of severe NM was also used. It resembled the clinical features of NM and it had a H40Y-substitution skeletal muscle α-actin. This model was generated by inserting a knock-in sequence of ACTA1 that codes for the H40Y-mutation into R129/Sv embryonic stem cells. The embryonic cells were introduced into blastocysts that were placed in pseudo-pregnant fe- males95. Mice were killed by cervical dislocation during isoflurane sedation and the hearts were removed. Different muscles, including extensor digitorum longus (EDL), tibialis anterior (TA), soleus (SOL), flexor digitorum brevis (FDB), gastrocnemius and diaphragm, were dissected from the animals in paper II and III. In paper IV, the fast hindlimb skeletal muscles EDL and TA were excised from two- to three-months old WT male and gender- and age- matched heterozygous transgenic animals that expressed the MYH4 L342Q mutation. All procedures involving animal care and handling were per-

24 formed according to institutional guidelines and were reviewed and approved by the local Uppsala Ethical Committee on Animal Research. Presence of the H40Y-substitution-causing ACTA1 mutation was determined by agarose gel electrophoresis. First the DNA from ear biopsies was purified and amplified in a standard PCR-reaction. Samples from the PCR- reaction were loaded on a 2%-agarose gel and gel bands were observed by UV-light.

Muscle isolation (Paper I, II, III, IV) Some of the biopsies and one of the excised muscles were frozen in liquid nitrogen-chilled liquid-propane and stored at – 160 °C for further analyses. The contralateral muscles were placed in ice-cold relaxing solution (in mM: 100 KCl, 20 imidazole, 7 MgCl2, 2 EGTA, 4 ATP, pH 7.0 at 4°C). Bundles of ~50 fibres were dissected free and tied to glass capillaries at slightly stretched lengths with surgical silk. They were treated with ‘skinning’ solution (relaxing solution containing glycerol 50:50 v/v) for 24 h at 4 °C. After 24 h they were transferred to – 20 °C. Within two weeks, the bundles were sucrose-treated, working as a cryo-protection, and snap-frozen in liquid- chilled liquid-propane for long-term storage at – 160°C101.

Contractile measurements of single fibres (Paper I, II and IV) On the day of experiment, bundles were desucrosed and single fibres were isolated. A 1-2 millimetre fibre segment was tightly fixed between two connectors. One of connectors was attached to a force transducer (model 400A, Aurora Scientific) and the other connector was joined with a lever arm system (model 308B, Aurora Scientific)102, 103. The apparatus was mounted on the stage of an inverted microscope (model IX70, Olympus). The sarcomere length was set to 2.50-2.60 µm (optimal mouse sarcomere length) in paper II and IV, while in paper I the sarcomere length was set to 2.65-2.75 µm (optimal human sarcomere length) by adjusting the overall length of the fibre segment. It was controlled during the experiment by a high-speed video analysis system (model 901A HVSL, Aurora Scientific). The diameter of an attached fibre was measured through the microscope at 320x magnification and the depth was assessed through the vertical displacement of the microscope nosepiece while focusing on the top and bottom of parts of the fibre’s sarcolemma with the focusing control working as a micrometer. This was done at three locations along the length of each fibre and the values aver- aged. The fibre’s CSA was calculated from the depth and diameter by as- suming an elliptical shape. It was corrected for the 20% swelling that occurs during the skinning process103. Mechanical recordings were done at 15°C and included force (normalized to CSA) and time measurements after various length steps at numerous [Ca2+] as previously described104-106.

25 After contractile recordings, the fibre was placed in urea buffer (120 g urea, 38 g thiourea, 70 ml H2O, 25 g mixed bed resin, 2.89 g dithiothreitol, 1.51 g Trizma base, 7.5 g SDS, 0.004% bromophenol blue) in -80 °C for determination of the MyHC isoform.

In vitro motility assay (Paper I) Fluorescent-labelled unregulated actin filaments from rabbit skeletal muscle was tested together with extracted myosin from patients and control subjects in a flow cell and placed on an inverted epifluorescense microscope. The movement of the actin-filaments were recorded by an image-intensified SIT camera (SIT 66; DAGE-MTIvideos). Motility speed and force index was calculated. Force index is the negative linear relationship between the rela- tive fraction of moving filaments and α-actinin concentration107.

Histochemistry (Paper I and III) Six micrometer thin cryo-sections were cut perpendicular to the longitudinal axis of the muscle fibres with a cryostat at -20°C. Sections were stained with haematoxylin and eosin for cross-sectional area (CSA) measurements, according to standard procedures. The cross-sectional area (CSA) of 100-150 muscle fibres, regardless of fibre types, was measured semi-automatically with the aid of an imaging application (Simple PCI 6.0 Hamamatsu Corpora- tion, USA). In addition, rhodamine-phalloidin (Molecular probes) staining of cryo-sections was performed and analysed by confocal microscopy with a PlanNeofluar 20x/0.5 objective in a Zeiss LSM 510 Meta.

Myosin heavy chain isoform expression (Paper I, II, III and IV) Expression of MyHC isoforms in single fibres was determined by 6% SDS- PAGE. The acrylamide concentration was 4% (w/v) in the stacking gel and 6% (w/v) in the running gel with 30% glycerol included in the gel matrix. The gels were silver-stained, scanned and quantified in a soft laser densitometer (Personal Densitometer SI, Molecular Dynamics)102. Cross-sections were dissolved into 100 µl of urea buffer after centrifuga- tion and heating (90°C for 2 minutes) and samples were loaded on 6% SDS- PAGE. The gels were then stained with Coomassie blue. The gels were subsequently scanned to determine the fibre type proportion102.

26 Myosin, actin and total protein quantification (Paper I, II, III and IV) Cross-sections (10 µm) were dissolved into 100 µl of urea buffer after cen- trifugation and heating (90°C for 2 minutes) and a volume of 4 μl was loaded on 12% SDS-PAGE. The gels were stained with Coomassie blue and subsequently scanned to determine the MyHC to actin ratio. Total protein content was, in paper II and III, determined by using Pierce 660 Protein Assay (Thermo Scientific Inc) according to manufacturer’s instructions. The absorbance of the samples were measured and related to a standard curve.

Electron-microscopy (Paper I, II and IV) EDL and diaphragm longitudinal sections were slightly stretched and fixed in 2.5% (vol/vol) glutaraldehyde for 48 hours. Then the samples were incu- bated in 1% (wt/vol) OsO4 for one hour, dehydrated with ethanol and em- bedded into plastic. Samples were examined in a Joel-1200EX electron microscope. Micrographs were taken from randomly selected fields at constant calibrated magnifications108.

Immunoblotting (paper III) Specific proteins were identified and their expression quantified by western blotting. 5-10 µg of total protein, depending on protein of interest, was loaded on a 12% polyacrylamide-gel. 10 and 5 µl ladder was loaded for orientation and size control. The following proteins were investigated: MuRF1, atrogin-1, S6 ribosomal protein, p-S6 ribosomal protein Ser240/244, mTOR , p-mTOR, AKT and p-AKT. α-actinin and β-tubulin were used for normalisa- tion. Immunoblots were quantified by Odyssey® Imaging Systems (LI- COR® Biosciences UK Ltd) were used for two-colour infrared fluorescent and chemiluminence detection according to manufacturer’s instructions. The intensity volumes of each protein signal were normalised to intensity volumes of α-actinin or β-tubulin depending on protein size of protein of interest.

Small-angle X-ray Scattering (paper I and IV) Arrays of ~30 single membrane-permeabilised muscle fibres were fixed between two ceramic chips and stored in skinning solution at -20°C until experiment. On the day of the experiment, the arrays were thoroughly washed with pre-activating solution and the chips were then placed in a specimen chamber. The sarcomere length of the fibres was adjusted to 2.65 µm in paper I and 2.50 µm in paper IV. X-ray diffraction patterns were re-

27 corded at 15°C in pre-activating solution and activating solution (pCa 4.5). 10-20 diffraction patterns were recorded for each array in both solutions and background scattering was subtracted. The intensities of the reflections were calculated109-111.

Actomyosin modelling (Paper I and II) A model of actin and myosin interaction in rigor state was used with and without insertion of a mutation in actin56. It was visualised with UCSF Chi- mera112 and Swiss PBD viewer. Energy computations in absence and presence of the actin mutation of interest were done in vacuo with the GRO- MOS96 implementation of Swiss PDB viewer.

Intact fibre preparation and measurements (Paper II) Intact single muscle fibres were dissected free from whole FDB. Each fibre with tendons was mounted between a force transducer (Akers 801) and an adjustable holder and placed in Tyrode solution (121 mmol/L NaCl, 5.0 mmol/L KCl, 1.8 mmol/L CaCl2, 0.5 mmol/L MgCl2, 0.4 mmol/L NaH2PO4, 24.0 mmol/L NaHCO3, 0.1 mmol/L EDTA, 5.5 mmol/L glucose, 95% O2/5% CO2 and pH 7.4). Fibre length was set to give maximum tetanic force. Contraction was induced by electrical stimulation through platinum 2+ electrodes. Force-[Ca ]i relationships were established by tetani stimulations at different frequencies and measurements of myoplasmic free [Ca2+] through the fluorescent Ca2+ indicator, Indo-1 (Invitrogen).

Statistics (Paper I, II, III and IV) Excel (Microsoft) and Sigma Plot (Systat Software Inc.) were used to gener- ate descriptive statistics. Data are presented as means ± standard error of the means. Where appropriate, either unpaired Student’s unpaired T-test or analyses of variance followed by post hoc Tukey tests were used for comparison between wild-type (WT) and mutant groups. If the data was not normally dis- turbed, the non-parametric Mann-Whitney rank-sum test was performed. Otherwise regressions were performed and relationships were considered significant at P<0.05.

28 Results and discussion

Paper I In the first paper, we studied muscle biopsies from a middle-aged male diagnosed with NM and suffering from muscle weakness. Previous genetic test- ing has revealed a mutation in exon 7 of the ACTA1 gene that leads to a F352S amino acid substitution. The patient had, as often seen in NM, a predominance of slow type I fibres, therefore most of the comparisons of contractile properties were restricted to MyHC type I fibres. The force production was increased by 80 % at maximal Ca2+ activation (pCa 4.5) in skinned fibres from the patient compared to healthy controls. In line with this result, stiffness measurements revealed increased stiffness under both active and rigor conditions, suggesting that the strain per individual myosin cross- bridge is greatly increased (a gain of function). One potential explanation is the alteration of non-bonded interactions between myosin and actin in presence of the F352S mutation as shown with computer modelling. To gain further molecular insights, small-angle x-ray scattering was performed. Surprisingly, during contraction, a slight decrease in the ratio I1,1/I1,0 was observed in fibres from the patient compared to healthy controls, proving a small decrease in the number of myosin cross-bridges attached to the actin filaments. Analysis of cross-bridge cycling kinetics confirmed this statement. Indeed, ktr was decreased whilst V0 was maintained, implying that myosin attachment rate is slowed down, reducing the time spent by each myosin head in the strong binding state. Altogether, during contraction, the F352S actin mutation is likely to slow down the myosin attachment rate and reduce the number of actual cross- bridges but, at the same time, stiffen and favour the actomyosin interaction by creating new bonds between these two proteins, ultimately increasing fibre force production. These findings are quite surprising and cannot explain the muscle weakness in the patient. Other processes, such as the neural drive, excitation-contraction coupling or Ca2+ handling, are on the other hand, potentially altered and responsible for the skeletal muscle weakness. Designing a mouse model carrying such mutation might help to answer this missing part of knowledge.

29 Paper II To further elucidate whether gain of function is also true in the presence of other disease-causing actin mutations, in the second paper, we studied limb muscle samples from a recently developed mouse model of severe NM (ACTA1 gene mutation that results in a H40Y amino acid substitution). In addition, we investigated whether the respiratory muscles, i.e. diaphragm muscles, were as badly affected. On the contrary to the F352S-actin mutation, in the presence of the H40Y mutation, the force production and active stiffness were decreased at maximal activation (pCa 4.5) in EDL and diaphragm skinned fibres from transgenic mice compared to wild-type animals, indicating a loss of function. However, rigor stiffness was only reduced in the diaphragm muscle. As the fraction of strongly bound cross-bridges (fxb), calculated using the following equation113; / / was only different in EDL, one may suggest that distinct mechanisms underlie limb and respiratory weakness. In EDL, the H40Y mutation functions as a ‘poison peptide’. It directly alters non-bonded and electrostatic interactions within and between actin monomers, disturbing the actin-actin interface, probably by changing actin flexibility during contraction and decreasing the number of attached myosin cross-bridges. Overall it induces a disrupted fibre force production, contributing to weakness. In the diaphragm, morphological and electron microscopy-based investigations revealed significantly large areas with sarcomeric disarray. These areas appear to be mainly responsible for the decreased fibre force production. Mechanical recordings from intact muscle fibres confirmed that the weakness originates from perturbations at the sarcomeric and myofilament levels. Altogether, these findings are of great importance, demonstrating that actin point mutations can lead to a loss versus gain of function and may have muscle-specific consequences. Therefore, these results have to be considered when designing future therapies.

Paper III Besides a change in function, disease-causing actin mutations are also known to induce muscle fibre atrophy, which can obviously contribute to the weakness. In the third paper, we decided to explore the unknown underlying mechanisms using the same mouse model as in paper II. Intriguingly, muscle fibre atrophy was only seen in the soleus which is considered as a slow-twitch muscle. On the other hand, a striking preferential myosin loss was observed in fibres from the fast-twitch tibialis anterior.

30 Immunoblotting was applied to define the molecular determinant of these findings. Interestingly, in both muscles, the protein synthesis rate seemed unaffected by the presence of the H40Y mutation as attested by unchanged levels of S6 ribosomal protein phosphorylation114, 115. However, in the tibialis anterior, the muscle-specific E3 ligase MuRF1 was up-regulated, whilst another muscle-specific E3 ligase, atrogin-1, was down-regulated. In the soleus, on contrary, atrogin-1 was up-regulated. The molecular pathways that regulate the MuRF1 and Atrogin-1 expression in this model remain to be investigated, as we did not find any difference in the AKT-mTOR signalling. In the tibialis anterior, the up-regulation of MuRF1 may well explain the preferential myosin loss and maintained fibre size since MuRF1 is thought to specifically degrade thick filament proteins, including myosin, but not thin filament proteins116, 117. Such up-regulation of MuRF1 and associated loss of myosin is in agreement with studies on critical illness myopathy118, 119. On the other hand, in soleus, a general loss of protein is likely to occur, probably through an Atrogin-1-dependent mechanism, which underlies the fibre atrophy. Other mechanisms may also contribute to the atrophy and the protein loss, such as apoptosis. One study showed that different NM-causing actin mutations can lead to cell death in cultured myotubes120. As muscle cells are multinucleated cells, apoptosis might only affect distinct regions of a muscle cell, causing termination of that region, which could result in an overall atrophic appearance121. Overall, this is the first time a direct link between a nemaline myopathy- causing mutation and the ubiquitin-proteasome system could be established. Another protein that, if mutated, has been associated with NM is KBTBD13. KBTBD13 is a muscle-specific protein and is believed to be part of the ubiquitin-proteasome complex and be involved in the ubiquitination process, but if other NM-associated genes affect the ubiquitin-proteasome system are so far unclear. These results open up potential therapies for NM that specifically targets the ubiquitin-proteasome system and modulate its activity to prevent the observed protein loss122, 123.

Paper IV In the final paper, I studied muscle samples from a recently developed mouse model of MFM (MYH4 gene mutation, which causes a L342Q amino acid replacement in the MyHC IIb protein). This mutation is located in the catalytic domain of the myosin head. Homozygous mice carrying this mutation develop hindlimb paralysis by postnatal day 13 and they need to be euthanised100, therefore we used heterozygous rodents (containing 7% mutant protein). The force production and active stiffness were increased at maximal Ca2+ activation (pCa 4.5) whilst rigor stiffness was maintained in

31 skinned fibres from transgenic mice compared to wild-type animals (a gain of function). This suggests that a higher proportion of myosin heads are attached to actin in the strong-binding state due to a faster myosin attachment rate and a slower detachment rate as ktr was unchanged. To have a further molecular understanding, small-angle x-ray scattering was used. During contraction, an increase in the ratio I1,1/I1,0 was observed in fibres from the transgenic mice compared to wild-type rodents, confirming the increased number of myosin cross-bridges. In addition, the myosin meridional (MM) reflection was recorded and analysed. The 1st MM reflection was increased proving that mutant myosin heads have a different orientation compared to wild-type heads, due to changes in the attachment and detachment rates as explained earlier, which affect the duration of the strong binding state and in the end will be responsible for the increased stiffness and force production. The cross-bridge cycling alterations also give rise to an increased Ca2+ sensitivity, in accordance with a simple analytical model that combines both thin filament activation and strong actomyosin binding124. L342 is a highly evolutionary conserved amino acid residue located close to sites involved in ATP/ADP binding and exchange in the catalytic domain51, 125, 126. The gain of a positive charge, which is the consequence of leucine to glutamine mutation, probably interferes with normal ATP-binding and/or ADP.Pi dissociation, which may explain the cross-bridge cycling alterations. As stated previously, the mutant protein content was only 7% in heterozygous animals100. It implies that the phenotype is likely severely exacer- bated in homozygous animals where all MyHC IIb proteins are mutated, explaining the muscle structure disruption that is associated with MFM and observed in this model100. The main limitation with this model is that the MyHC IIb protein is not expressed in human skeletal muscles. Although the MYH4 gene is present in the human genome, its mRNA has only been found in jaw-closing human muscles127. However the gene’s product, the MyHC IIb protein, has not been detected anywhere in the human body47.

32 Conclusion

Congenital myopathies are a poorly understood group of diseases and this thesis aimed to unravel some of the underlying mechanisms. In particular, this thesis has investigated the molecular and cellular defects in skeletal muscle from patients and animals with three different mutations in two genes, ACTA1 and MYH4, associated with the congenital myopathies NM and MFM respectively. The most important finding was that mutations causing congenital myopathies have different weakness-inducing mechanisms and functional alterations (gain versus loss of function). These mechanisms are mutation- specific, as seen in paper I, II and IV, but are also observed in other studies128. The strong structure-function relationship, including electrostatic and polarity-based interactions, is determined by the amino acid sequence and when a substitution occurs, it will affect these properties. Depending on how the mutation changes these properties, the final outcome on the contractility will vary. Surprisingly, the NM-causing H40Y-actin mutation had different contractile defects in limb and respiratory muscles. This particular mutation also had distinct effects on the protein degradation pathways. MuRF1 was up- regulated and atrogin-1 was down-regulated in the fast-twitch tibialis anterior muscle while in the soleus muscle, the expression of MuRF1 was unchanged and a somewhat increased expression of Atrogin-1 was observed. These results hint that the H40Y-substitution severely impairs normal muscle function in many different aspects, but this also opens up potential therapeutic options that can help in improving muscle function in patients. The MFM-related L342Q MyHC IIb mutation gives important information about the contractile defects that possibly could be found in MFM patients, despite the fact that MyHC IIb is not expressed in humans. The MyHC gene family is highly evolutionary conserved from a structural view and the mutated L342 residue is found in all skeletal muscle myosin isoforms. This implies that this transgenic mouse could be a useful model for further studies of MFM45, 100. The fact that congenital myopathies are caused by a vast variety of mutations in several different genes, restricts the number of patients suffering from a specific mutation. This together with my findings of different weakness-inducing mechanisms for NM- and MFM-causing mutations slightly limits the conclusion of this thesis. It also restricts the number of patients

33 that could benefit from a therapy that is based on counteracting phenomena that is caused by a specific mutation. Nevertheless my research work sheds light on different mechanisms that can underlie the muscle weakness associated with NM and MFM and it points to possible therapeutic approaches.

34 Future therapies

NM and MFM are in many cases lethal and no curative treatments exist to- day44, 93. Potential treatments could be based on several molecular mechanisms, such as RNA interference (RNAi), inhibition of myostatin and modu- lation of proteins involved in contraction.

RNA interference Specific degradation of mutant mRNA could potentially treat any disease that is caused by dominant single-allele mutations, which often are the case in congenital myopathies. Removal of the mutant mRNA would prevent production of the malfunctioning protein, so only wild-type mRNA is trans- lated to functional protein. A system that can do this is RNA interference. In RNAi, long double-stranded RNA (dsRNA) transcripts are cleaved by the endoribonuclease Dicer into small interfering RNAs (siRNA). These short sequences are transferred to the RNA-induced silencing complex (RISC) that separates the dsRNAs and releases one of the strands. The RISC now only contains one single-stranded piece of RNA, which allows the RISC to bind, by base pair recognition, an mRNA homologous in sequence to the siRNA. This mRNA is then degraded, which stops expression of the corresponding protein129. Muscle fibre force production is determined by the fibre size and the efficiency of contraction. This implies that there are two ways to improve force production of a muscle fibre. One is to increase muscle fibre size and the other is to improve the efficiency of contraction.

Inhibition of myostatin Myostatin plays a crucial role during developmental and postnatal muscle growth as a negative regulator of muscle size. It is bound by the activin type IIB receptor and upon binding the receptor forms a heterodimer with activin- like kinase, which, in turn, phosphorylates SMAD2 and 3. SMAD2 and 3 then make a complex with SMAD4. This complex regulates genes involved in proliferation and differentiation of muscle precursor cells, protein degradation pathways in mature muscle fibres and inhibits protein synthesis through the mTOR/AKT pathway130. Muscle size could therefore be increased by inhibiting the action of myostatin, which could counteract muscle

35 fibre atrophy seen in NM131. Myostatin inhibition could be achieved by using a soluble activin type IIB receptor that removes circulating endogenous my- ostatin132.

Improving efficiency of contraction One way to improve efficiency of contraction is to modulate the MyLCs that are attached to the MyHC. It is known that the essential (alkali) MyLCs pro- vide fine-tuning of myosin motor activity. In particular, expression of the atrial MyLC1 in hypertrophied ventricles increases its contractility. The underlying mechanism for this increased contractility is likely that the MyLC1 stiffens the neck region of the myosin molecule making it more resilient to distortion, which enhances the lever arm swing133-135. Introduction of the essential atrial MyLC1 gene by adeno-associated viral vectors into skeletal muscles and its following expression could, through the above-mentioned mechanism, improve force production in the treated muscles.

36 Acknowledgments

This work was performed at Clinical Neurophysiology, Department of Neu- roscience, Uppsala University, Uppsala, Sweden in collaboration with research groups at Départment de Neurologie, Centre de reference des mala- dies neuromusculaires, Angers, France; Japan Synchrotron Radiation Re- search Institute, Spring-8, Hyogo, Japan; Department of Physiology and Pharmacology, Karolinska Institutet, Stockholm, Sweden; Neuromuscular and Regenerative Medicine Unit, University of New South Wales, Sydney, Australia and Biology department, University of York, York, United King- dom.

I would like to express my gratitude to the people who directly or indirectly contributed to this thesis. I would especially like to thank the following the persons:

First I want to express my deepest gratitude to my supervisor Julien Ochala, for being the best supervisor one can have. You have always been patient, supportive and excellent at explaining fibre mechanics as well as pushing me to go to different parts of the world.

My co-supervisor, Lars Larsson, for good support and sharing of his vast knowledge in the skeletal muscle field, and providing the lab space and equipment need for performing the work done in this thesis.

To Ann-Marie and Yvette for providing excellent technical assistance and being friendly and cheerful co-workers. Yours supportive work made this thesis a lot easier.

The former members of the lab; Rizwan, Monica, Sudhakar and Guillaume as well as students Marit, Mayank, Rutger and Johan. The kind and cheerful present members of the group; Heba, Nicola, Hannah, Hazem, Meishan, Stefano and Maria. I really enjoyed working together with all of you. You are good co-workers and friends.

A special thanks to my office mate Rebeca for good advises regarding immunoblotting and being a good friend and colleague.

37 The staff at the Clinical Neurophysiology for their warm and welcoming environment and the very nice tradition of Friday breakfasts.

Professor Naoto Yagi and Dr Hiroyuki Iwamoto for giving the opportunity to go Japan and perform the X-ray scattering experiments as well as explor- ing Japan.

Professor Edna Hardeman for donating one of the mice models and making this thesis possible. The staff at Rudbeck and BMC’s animal facilities for expert handling of the mice and excellent technical assistance.

The administrators Inger, Annika, Mona and Gun for taking care of the fi- nancial matters.

A special thank to Anna-Maria Lundin’s foundation for giving me the opportunity to attend many interesting and valuable conferences both close and far away.

All persons working together with me in the Medical PhD-student council during the years. It has been a pleasant and interesting experience working together with all you.

All my friends both here in Uppsala and from my home areas, you have made the struggle a lot easier by getting me out of the laboratory from time to time.

Till sist men inte minst min familj utan ert stöd och uppmuntran hade detta inte varit möjligt och för det är jag evigt tacksam.

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45 Acta Universitatis Upsaliensis Digital Comprehensive Summaries of Uppsala Dissertations from the Faculty of Medicine 977 Editor: The Dean of the Faculty of Medicine

A doctoral dissertation from the Faculty of Medicine, Uppsala University, is usually a summary of a number of papers. A few copies of the complete dissertation are kept at major Swedish research libraries, while the summary alone is distributed internationally through the series Digital Comprehensive Summaries of Uppsala Dissertations from the Faculty of Medicine. (Prior to January, 2005, the series was published under the title “Comprehensive Summaries of Uppsala Dissertations from the Faculty of Medicine”.)

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