The function of TGF- β 1 in ICUAW and the characterization of Sfrp2, a TGF- β 1 target, in
skeletal muscle atrophy
Dissertation
zur Erlangung des akademischen Grades doctor rerum naturalium (Dr. rer. nat.)
im Fach Biologie eingereicht an der
Lebenswissenschaftliche Fakultät an der Humboldt-Universität zu Berlin
von
M. Sc. Xiaoxi Zhu
Präsident der Humboldt-Universität zu Berlin Prof. Dr. Jan-Hendrik Olbertz
Dekan der Lebenswissenschaftliche Fakultät Prof. Dr. Richard Lucius
Gutachter/in: 1. Prof. Dr. rer. nat. Michael Bader 2. PD Dr. med. Jens Fielitz
3. Prof. Dr. rer. nat. Harald Saumweber Tag der mündlichen Prüfung: 18.12.2014
Zusammenfassung
Zusammenfassung
Transforming growth factor beta 1 (TGF-beta1) ist ein multifunktionales Zytokin, welches eine Rolle in der Sepsis und in der Sepsis-induzierten Myopathie spielen könnte. Weiterhin könnten erhöhte TGF-beta1-Level zur Muskelschwäche, die mit der Intensivpflege assoziiert ist (engl. intensiv care unit-acquired weakness, ICUAW), beitragen. Der TGF-beta1- Signalweg wurde in Skelettmuskelbiopsien von ICUAW-Patienten heraufreguliert. Secreted frizzled related protein 2 (SFRP2) wurde in einer Gen-Set-Anreicherungsanalyse als das am höchsten regulierte Gen identifiziert. Im Mausmodell führten Sepsis und Hunger zu einer verringerten Sfrp2-Expression, während dies in der Denervation-induzierten Skelettmuskelatrophie nicht festzustellen war. In differenzierten C2C12-Myotuben führte TGF-beta1 zu einer verringerten Sfrp2-mRNA- und Proteinexpression. Luciferase-Assays deuteten auf eine TGF-beta1-abhängige Herunterregulation von Sfrp2 hin, welche auf Promoterebene durch mögliche negative regulatorische Elemente im Sfrp2-Promoter vermittelt wurde. Weiterhin wurde eine TGF-beta1 induzierte Muskelatrophie durch transkriptionelle Repression der myosin heavy chain Gene beobachtet. Im Gegensatz dazu veränderte TGF-beta1 nicht den proteasomalen Abbau muskulärer Proteine. Die Genexpression von Tripartite motif containing 63 und F-box only protein 32 war hingegen leicht herunterreguliert. TGF-beta1-induzierte Atrophie in differenzierten C2C12-Myotuben wurde teilweise durch rekombinantes Sfrp2 aufgehoben. Weiterhin wurde eine direkte physikalische Interaktion zwischen Sfrp2 und TGF-beta1 gefunden, welche diesen Effekt verursacht haben könnte. Zusammengefasst lässt sich feststellen, dass der TGF-beta1- Signalweg eine wichtige Rolle in der ICUAW durch Inhibition der myosin heavy chain Expression spielt. TGF-beta1-abhängige Herunterregulation von Sfrp2 könnte zu einer Feedback-Antwort, die das Ausmaß der Atrophie durch TGF-beta1 verstärkt, führen.
Schlagworte:
Intensivstation erworbene Muskelschwäche, Skelettmuskelatrophie,
Transforming growth factor beta 1 (TGF-beta1), Secreted frizzled related protein 2 (Sfrp2)
Abstract
Abstract
Transforming growth factor beta 1 (TGF-beta1) is a multifunctional cytokine that may play a role in sepsis and in sepsis-induced myopathy. Our group speculated that increased TGF-beta1 could contribute to intensive care (ICU)-acquired weakness (ICUAW), a catastrophic muscle disease in critically ill patients. We found that TGF-beta1 signaling in skeletal muscle biopsies of ICUAW patients was upregulated. Secreted frizzled related protein 2 (SFRP2) was the most regulated gene identified by gene set enrichment analysis (GSEA). I then studied the regulation and function of SFRP2 in different skeletal muscle atrophy models. In three mouse models, downregulated Sfrp2 expression was observed in sepsis and starvation, but not in denervation-induced skeletal muscle atrophy. In differentiated C2C12 myotubes, TGF-beta1 downregulated Sfrp2 expression on both mRNA and protein levels. Luciferase assays suggested that TGF-beta1-dependent downregulation of Sfrp2 was mediated at the promoter level through possible negative regulatory elements in the Sfrp2 promoter. I also observed that TGF-beta1-induced muscle atrophy was accompanied by transcriptional repression of myosin heavy chain genes. In contrast, TGF-beta1 did not increase proteasomal degradation of muscular proteins since gene expression of Tripartite motif containing 63 (Trim63) and F-box only protein (Fbxo32) was not upregulated; instead, they were slightly downregulated. TGF- beta1-induced differentiated C2C12 myotube atrophy was partially reversed by recombinant Sfrp2. This inhibitory effect could have resulted from direct interaction between Sfrp2 and TGF-beta1, since I found a physical interaction between these two proteins. Taken together, TGF-beta1 signaling pathway could play an important role in ICUAW via inhibition of myosin heavy chain expression. TGF-beta1-dependent downregulation of Sfrp2 may establish a feedback loop augmenting the atrophic effect of TGF-beta1.
Keywords:
Intensive care unit (ICU)-acquired weakness (ICUAW), Skeletal muscle atrophy, Transforming growth factor beta 1 (TGF-beta1), Secreted frizzled related protein 2 (Sfrp2)
Table of content
Table of Content
Zusammenfassung ... 2
Abstract ... 3
List of abbreviations ... 8
1 Introduction ... 14
1.1 Anatomy and physiology of skeletal muscle ... 14
1.2 Intensive care unit-acquired weakness (ICUAW) ... 19
1.2.1 Definition of ICUAW and its consequences ... 19
1.2.2 Risk factors for ICUAW ... 19
1.2.3 Mouse models for studying ICUAW ... 21
1.3 Skeletal muscle atrophy ... 21
1.3.1 Skeletal muscle atrophy in ICUAW ... 22
1.3.2 Molecular mechanisms involved in skeletal muscle atrophy ... 23
1.3.2.1 TGF-β pathway ... 23
1.3.2.2 Proteolytic pathways and protein degradation ... 28
1.3.2.3 Functions of Akt signaling pathway in regulating protein synthesis and degradation ... 29
1.4 Secreted Frizzled Related Proteins (SFRPs) ... 31
1.4.1 SFRP family ... 31
1.4.2 Functions of Sfrp2 ... 31
1.4.3 Mechanisms of Sfrp2 downregulation ... 32
1.5 Aim of this study ... 34
2 Materials and methods ... 35
2.1 Materials ... 35
2.1.1 Primers ... 35
2.1.2 Antibodies ... 38
2.1.3 Equipment ... 39
2.1.4 Reagents ... 42
2.1.5 Patient samples ... 48
2.1.6 Animals ... 49
2.1.7 Cell lines ... 49
2.1.8 Plasmids ... 49
2.2 Methods ... 50
2.2.1 Microarray analysis of patient samples ... 50
2.2.2 Animal experiments ... 50
2.2.2.1 Sepsis-induced skeletal muscle atrophy ... 50
2.2.2.2 Starvation-induced skeletal muscle atrophy ... 51
Table of content
2.2.2.3 Denervation-induced skeletal muscle atrophy ... 51
2.2.3 Molecular biological methods ... 51
2.2.3.1 RNA isolation ... 51
2.2.3.2 Complementary DNA (cDNA) synthesis ... 52
2.2.3.3 Quantitative real-time polymerase chain reaction (qRT-PCR) 52 2.2.4 Protein chemical methods ... 54
2.2.4.1 Protein isolation ... 54
2.2.4.2 Total protein determination ... 54
2.2.4.3 Sodium dodecylsulphate polyacrylamide gel electrophoresis .. 55
2.2.4.4 Western blot ... 55
2.2.5 Histology ... 56
2.2.5.1 Cryosectioning of muscle tissues ... 56
2.2.5.2 Hematoxylin and eosin staining ... 56
2.2.5.3 Metachromatic Myosin ATPase staining ... 57
2.2.5.4 Immunofluorescence ... 59
2.2.5.5 Microscopic documentation ... 60
2.2.5.6 Measurement of myocyte cross sectional area ... 60
2.2.6 Generation of expression plasmids ... 60
2.2.6.1 Polymerase chain reaction ... 60
2.2.6.2 Restriction digest and ligation of DNA ... 61
2.2.6.3 Sequencing ... 61
2.2.7 Cell biological methods ... 61
2.2.7.1 Cell culture ... 61
2.2.7.2 Treatment of cells ... 62
2.2.7.3 Transient transfection of cDNA expression plasmids ... 62
2.2.8 Bisulfite sequencing PCR (BSP) ... 62
2.2.9 Methylation specific PCR (MSP) ... 64
2.2.10 Luciferase assay ... 64
2.2.11 Co-immunoprecipitation ... 65
2.2.12 Statistics ... 65
3 Results ... 66
3.1 Regulation of gene expression in skeletal muscle of ICUAW patients ... 66
3.1.1 Expression of atrophy genes was increased in skeletal muscle of ICUAW patients. ... 66
3.1.2 TGF-β1 signaling was activated in ICUAW patients. ... 66
3.2 TGF-β1-induced skeletal muscle atrophy ... 70
3.2.1 TGF-β1 treatment resulted in atrophy of C2C12 myotubes. ... 70
3.2.2 Transcriptional regulation of gene expression in TGF-β1-treated C2C12 myotubes ... 70
Table of content
3.3 Regulation and function of Secreted frizzled related protein 2 (Sfrp2) in
skeletal muscle atrophy ... 72
3.3.1 Transcriptional regulation of SFRP family genes in ICUAW patients ... 72
3.3.2 Downregulation of Sfrp2 by TGF-β1 and dexamethasone in vitro ... 74
3.3.3 Regulation of Sfrp2 in mouse models of skeletal muscle atrophy ... 76
3.3.3.1 Sepsis ... 76
3.3.3.2 Starvation ... 81
3.3.3.3 Denervation ... 85
3.3.4 Mechanism of downregulation of Sfrp2 in muscle atrophy ... 91
3.3.4.1 TGF-β1 decreased expression of Sfrp2 promoter luciferase constructs. ... 91
3.3.4.2 Downregulation of Sfrp2 in muscle atrophy in vivo and in vitro was not due to promoter hypermethylation. ... 93
3.3.5 Fiber-type-specific expression of Sfrp2 in mouse skeletal muscles ... 96
3.3.6 Sfrp2 inhibited TGF-β1-incuded atrophy in C2C12 myotubes. .. 98
3.3.6.1 Sfrp2 inhibited TGF-β1-incuded atrophy in C2C12 myotubes.98 3.3.6.2 Physical interaction between Sfrp2 and Tgf-β1 ... 99
4 Discussion ... 102
4.1 TGF-β1 signaling in skeletal muscle atrophy ... 102
4.1.1 TGF-β1 signaling was activated in skeletal muscle of ICUAW patients ... 102
4.1.2 TGF-β1 induced skeletal muscle atrophy in vitro ... 104
4.2 Regulation of a TGF-β1 signaling target gene Sfrp2 in skeletal muscle atrophy ... 105
4.2.1 TGF-β1 signaling downregulated Sfrp2 in skeletal muscle atrophy ... 105
4.2.2 Regulation of Sfrp2 in mouse models of skeletal muscle atrophy ... 106
4.2.3 Mechanism of downregulation of Sfrp2 in skeletal muscle atrophy ... 107
4.3 Characterization of Sfrp2 and its function in TGF-β1-induced skeletal muscle atrophy ... 108
4.3.1 Fiber-type-specific expression of Sfrp2 ... 108
4.3.2 Sfrp2 rescued TGF-β1-induced atrophy in vitro ... 109
4.3.3 Physical interaction between Sfrp2 and Tgf-β1 ... 110
Table of content
4.4 Hypothesis and outlook ... 111
5 Literature ... 113
Acknowledgement ... 123
Declaration ... 124
List of publication ... 125
List of abbreviations
List of abbreviations
µ 4eBP1 AA ActRIIB ANXA2 Akt ATCC ATP BDNF BDPC BMP bp BSP CASP3 CIM CINM CIP CK CLP
micro
4e binding protein 1 Amino acids
Activin receptor IIB Annexin2
V-Akt Murine Thymoma Viral Oncogene Homolog American Type Culture Collection
Adenosine triphosphate
Brain-derived neurotrophic factor
Bisulfite sequencing Data Presentation and Compilation Bone morphogenetic protein
base pairs
Bisulfite Sequencing PCR Caspase 3
Critical-illness myopathy Critical illness neuromyopathy Critical illness polyneuropathy Creatine kinase
Caecal ligation and puncture
List of abbreviations
CNS COPD CpG CRD CS DAPI DCN DMEM E1 E2 E214k
E3 eIF ELISA Fbxo FGF FoxO Fzl g
GAPDH
Conserved non-coding sequence
Chronic obstructive pulmonary disease Cytosine-phosphate-guanine dinucleotide Cysteine rich domain
Citrate synthase
4',6-diamidino-2-phenylindole Decorin
Modified Eagle's Medium Ubiquitin-protein ligases Ubiquitin-conjugating enzymes 14-kDa Ubiquitin-conjugating enzyme Ubiquitin-activating enzymes
Eukaryotic initiation factor
Enzyme-linked immunosorbent assay F-box only protein
Fibroblast growth factor Forkhead box o protein Frizzled
gram
Glyceraldehyde-3-phosphate dehydrogenase
List of abbreviations
GDF GSEA GSK3-β GTP h H&E HDAC H-LDH Hs ICU ICUAW IGF IL IR IRS l
LAMA2 m
MAP2K1 MAPK
Growth/differentiation factor Gene set enrichment analysis Glycogen synthase kinase 3 beta Guanosine-5'-triphosphate hour/(s)
Hematoxyline & eosin
Heart type lactate dehydrogenase Histone deacetylase
Homo sapiens
Intensive care unit
Intensive care unit-acquired weakness Insulin like growth factor
Interleukin Insulin receptor
Insulin receptor substrate liter
Laminin, alpha 2 mili
Mitogen-activated protein kinase kinase 1 Mitogen-activated protein kinase
List of abbreviations
M-CK MCSA MEF Mef2A min M-LDH Mm Mrf4 MSC MSP mTORC MuRF MW myc MyCH Myf5 MyLC MyoD n NF-κB
Muscle type creatine kinase Myocyte cross sectional area Mouse embryonic fibroblast Myocyte enhancer factor 2A minute/(s)
Muscle type lactate dehydrogenase Mus musculus
Myogenic regulatory factor 4 Mesenchymal stem cell Methylation-Specific PCR
Mammalian target of rapamycin complex Muscle RING finger protein
Molecular weight
V-myc avian myelocytomatosis viral oncogene homolog Myosin heavy chain
Myogenic factor 5 Myosin light chain Myogenic differentiation nano
Nuclear factor kappa-light-chain-enhancer of activated B cells
List of abbreviations
NMES NTR Pax3 PBS PCOLCE PDB PDK1 PEI PGC1α PI3K PIP3 qRT-PCR Raf Ras Rho RIPA RUNX S6K1 SAA1 SDH
Neuromuscular electrical stimulation Netrin-Related
Paired box 3
Phosphate buffered saline
Procollagen C-proteinase enhancer protein Protein Data Bank
Phosphoinositide-dependent kinase 1 Polyethylenimine
Peroxisome proliferator-activated receptor gamma coactivator 1- alpha Phosphatidylinositol-3-kinase
Phosphoinositide-3,4,5-triphosphate
Quantitative real-time polymerase chain reaction Rapidly accelerated fibrosarcoma
Rat sarcoma Rhodopsin
Radioimmunoprecipitation assay buffer Runt domain transcription factor Ribosomal protein S6 kinase Serum amyloid A1
Succinate dehydrogenase
List of abbreviations
SDS-PAGE SEM SFRP Smad SOFA SPARC TGF TGFβR TIE TIMP TNF-α Trim TSS TWEAK UPS UTR Vg1 WNT
Sodium dodecyl sulfate polyacrylamide gel electrophoresis Structural Equation Modeling
Secreted Frizzled Related Protein SMA/MAD-related
Sequential Organ Failure Assessment Secreted protein, acidic, cysteine-rich Transforming growth factor
TGF-β receptor
TGF-β inhibitory element
Tissue inhibitors of metalloproteinase Tumor necrosis factor-alpha
Tripartite motif containing Transcriptional start site
TNF-related weak inducer of apoptosis Ubiquitin-proteasome system
Untranslated region Vegetalising factor-1
Wingless-Type MMTV Integration Site Family
Introduction
1 Introduction
1.1 Anatomy and physiology of skeletal muscle
In mammals, skeletal muscle accounts for more than 40 % of the body mass and is responsible for all voluntary movement (Mcnally et al., 2006) (Wang and Pessin, 2013).
Every individual muscle fiber (or so-called myofibers) is formed after fusing of a number of differentiated mono-nucleated myoblasts. Multiple nuclei are located at the periphery of each muscle fiber. Mature muscle fibers do not have the ability to divide. Therefore, new myofibers can only be formed from undifferentiated satellite cells (muscle stem cells) for example after muscle injury. Every bundle of muscle fibers is bounded by a connective tissue sheath, called perimysium. Perimysium is then continuous with a dense connective tissue, the epimysium.
At the end of each skeletal muscle, there is a myotendinous junction, which transmits force longitudinally along the long axis of the muscle. Motor neurons innervate skeletal muscles through axons and are responsible for transmitting signals from the central nervous system to the muscle fibers (Figure 1) (Mcnally et al., 2006) (Scime et al., 2009).
Individual myofibrils are composed of sarcomeres, which are the basic units of muscle contraction. Each sarcomere includes two A bands containing thick (myosin) filaments, two I bands containing thin (actin filament), and two Z lines made up of actin binding proteins. The spaces were thick and thin filaments overlap is called cross-bridge. These cross-bridges between actin and myosin filaments generate muscle contraction (Figure 2A) (Widmaier et al., 2004) (Mcnally et al., 2006). The myosin molecule contains ATP binding sites that also serve as ATPase providing energy for muscle contraction. Each myosin molecule contains two myosin heavy chains (MyHCs) and two regulatory myosin light chains (MyLC). MyHCs form ATP- and actin-binding sites (Figure 2B) (Widmaier et al., 2004) (Mcnally et al., 2006).
Heterogeneity is a fundamental feature of skeletal muscle. Mammalian skeletal muscle is composed of four types of muscle fibers defined by their different MyHC compositions. There is one type of slow-twitch (type I), and three types of fast-twitch myofibers (type IIa, type IIx/IId, and type IIb) (Table 1) (Schiaffino and Reggiani, 2011) (Wang and Pessin, 2013) (Ciciliot et al., 2013). In contrast to rodents MyHC-IIb is not expressed in human muscles.
Therefore, there are only three types of muscle fibers in human. Different myofibers have distinct metabolic activities. For example, fast-twitch fibers (type IIx and type IIb) are characterized by glycolytic metabolism and high ATPase activity. They have higher contraction speed and are not resistant to fatigue. In the contrary, the slow-twitch type I fibers
Introduction
use oxidative metabolism and thus contain a large amount of oxidative enzymes and myoglobins. They have slower contraction speed and are resistant to fatigue. Type IIa fibers have a metabolic and physiological profile which is in between the very fast-twitch and the slow-twitch fibers (Table 1) (Schiaffino and Reggiani, 2011) (Ciciliot et al., 2013) (Wang and Pessin, 2013). The systematic distribution of muscle fiber types in four major skeletal muscles varies (Schiaffino and Reggiani, 2011). For example, among the four major hind limb skeletal muscles of mouse, M. soleus contains almost equal amounts of type I and type IIa fibers; M.
extensor digitorum longus, M. tibialis anterior and M. gastrocnemius muscles mainly contain type IIb fibers and only few type IIa fibers; M. gastrocnemius also contains a few type I fibers (Augusto et al., 2004).
Another important feature of skeletal muscle is its function as a secretory organ. Skeletal muscle produces different kinds of cytokines and peptides and releases/secretes them into the circulation. These signal molecules are called myokines (Pedersen and Febbraio, 2012).
Important for this thesis, myokines are expressed in a fiber-type-specific pattern (Plomgaard et al., 2005). For example, immunohistochemistry studies on human skeletal muscle biopsies demonstrated that tumor necrosis factor-alpha (TNF-α) and interleukin-18 (IL-18) were solely expressed in slow fibers, whereas interleukin (IL)-6 expression was mainly found in fast fibers (Plomgaard et al., 2005). This indicated that different myokines play diverse roles in regulating normal muscle physiology. Myokines have either autocrine or paracrine effects on muscle tissue (Table 2) (Plomgaard et al., 2005) (Pedersen and Febbraio, 2012) (Aoi and Sakuma, 2013). Moreover, myokines also target other organs, such as adipose tissue, liver, pancreas, heart, bones, and brain in an endocrine manner (Table 2) (Pedersen and Febbraio, 2012) (Aoi and Sakuma, 2013). Altered myokine expression may explain association between inflammation or disuse-induced muscle atrophy and many chronic diseases including diabetes, cardiovascular disease and cancer (Pedersen and Febbraio, 2012) (Aoi and Sakuma, 2013). Effects of myokines have been largely underestimated and studies of regulation and function of myokines are limited. Recently, we reported that systemic inflammation led to inflammatory response and acute phase response directly in myocytes of critically ill patients and in mice with polymicrobial sepsis. More specifically, muscle-derived IL-6 and serum amyloid A1 (SAA1) were increased in skeletal muscle of patients with high risk of critical- illness myopathy (CIM) a severe form of intensive care unit (ICU)-acquired weakness (ICUAW) (Langhans et al., 2014). In this study, I was also interested in studying regulation and function of myokines in ICUAW.
Introduction
16
Figure 1: Anatomy of skeletal muscle. Skeletal muscle is composed of bundles of multi-nucleated muscle fibers, satellite cells, motor neurons, blood vessels, and connective tissues. Adapted from (Scime et al., 2009).
A www.mhhe.com/widmaier9 271
Sliding-Filament Mechanism
When force generation produces shortening of a skele- tal muscle fiber, the overlapping thick and thin filaments in each sarcomere move past each other, propelled by movements of the cross-bridges. During this shortening of the sarcomeres, there is no change in the lengths of either the thick or thin filaments (Figure 9–8). This is known as the sliding-filament mechanism of muscle contraction.
During shortening, each myosin cross-bridge at- tached to a thin filament actin molecule moves in an arc much like an oar on a boat. This swiveling motion of many cross-bridges forces the thin filaments attached to successive Z lines toward the center of the sarcomere,
movement of a thin filament relative to a thick filament.
As long as a muscle fiber remains activated, however, each cross-bridge repeats its swiveling motion many times, resulting in large displacements of the filaments.
Thus, the ability of a muscle fiber to generate force and movement depends on the interaction of the contractile proteins actin and myosin.
An actin molecule is a globular protein composed of a single polypeptide that polymerizes with other actins to form two intertwined helical chains (Figure 9–10). These chains make up the core of a thin fila- ment. Each actin molecule contains a binding site for myosin. The myosin molecule, on the other hand, is composed of two large polypeptide heavy chainsand four smaller light chains. These polypeptides com-
Muscle fiber Myofibril
I band A band
Z line Z line Sarcomere
Myofibril
Z line Z line
Cross-bridge
Thick (myosin) filament Thin (actin) filament
FIGURE 9 – 4
Arrangement of filaments in a skeletal muscle fiber that produces the striated banding pattern.
Introduction
B
Figure 2: Composition of skeletal muscle fiber and thick filaments. (A) Structure of a myofiber. Most of the cytoplasm of a muscle fiber is filled with myofibrils. Sarcomere, the basic unit of myofibril is composed with thick and thin filaments, along with interconnecting proteins between filaments. In the thin filament: yellow, actin; blue=tropomyosin; pink=troponin complex. (B) Structure of a myosin molecule. The thick filament (upper panel) is composed of myosin molecules orienting in opposite directions. The two globular heads of each myosin molecule extending from the sides of a thick filament form a cross-bridge. A myosin molecule (lower panel) is made up of two myosin heavy chains (MyHCs) and two each of regulatory myosin light chains (MyLC). Each myosin globular head contains one ATP binding site and one actin binding site. Adapted from (Widmaier et al., 2004, with permission from McGrawHill Education).
www.mhhe.com/widmaier9 275
Thick filament
Cross-bridge
ATP binding site ATP
binding Light chains site
Heavy chains
Actin binding sites
Cross- bridge (a)
(b)
Myosin
FIGURE 9–11
(a) The heavy chains of myosin molecules form the core of a thick filament. The myosin molecules are oriented in opposite directions in either half of a thick filament. (b) Structure of a myosin molecule. The two globular heads of each myosin molecule extend from the sides of a thick filament, forming a cross-bridge.
the cross-bridge to a thin filament, (2) movement of the cross-bridge, producing tension in the thin filament, (3) detachment of the cross-bridge from the thin filament, and (4) energizing the cross-bridge so that it can again attach to a thin filament and repeat the cycle. Each cross-bridge undergoes its own cycle of movement independently of the other cross-bridges. At any instant during contraction only a portion of the cross-bridges are attached to the thin filaments and producing tension, while others are in a detached portion of their cycle.
The chemical and physical events during the four steps of the cross-bridge cycle are illustrated in Figure 9–12. In a resting muscle fiber the cytoplasmic calcium concentration is low, and the myosin cross-bridges (M) cannot bind to actin (A). The cross-bridges, however, are in an energized state produced by the splitting of ATP, and the hydrolysis products (ADP and inorganic phos- phate) are still bound to myosin. This storage of energy in myosin is analogous to the storage of potential en- ergy in a stretched spring.
Cross-bridge cycling is initiated by calcium entry into the cytoplasm (by a mechanism that will be de- scribed shortly). The cycle begins with the binding of an energized myosin cross-bridge to a thin filament actin molecule (step 1):
Step 1 A!M"ADP"Pi88nA"M"ADP"Pi
Actin binding
The binding of energized myosin to actin triggers the release of the strained conformation of the energized
bridge, which produces the movement of the bound cross-bridge (sometimes called the power stroke) and the release of Piand ADP (step2):
Step 2 A"M"ADP"Pi88nA"M!ADP!Pi
Cross-bridge movement
This sequence of energy storage and release by myosin is analogous to the operation of a mousetrap: Energy is stored in the trap by cocking the spring (ATP hydroly- sis) and released after springing the trap (binding to actin).
During the cross-bridge movement, myosin is bound very firmly to actin, and this linkage must be bro- ken in order to allow the cross-bridge to be re-energized and repeat the cycle. The binding of a new molecule of ATP to myosin breaks the link between actin and myosin (step 3):
Step 3 A"M!ATP88nA!M"ATP
Cross-bridge dissociation from actin
The dissociation of actin and myosin by ATP is an ex- ample of allosteric regulation of protein activity. The binding of ATP at one site on myosin decreases myosin’s affinity for actin bound at another site. Note that ATP is not split in this step; that is, it is not acting as an en- ergy source but only as an allosteric modulator of the myosin head that weakens the binding of myosin to actin.
Introduction
Type I Type IIa Type IIx/IId Type IIb
Gene Myh7 Myh2 Myh1 Myh4
Protein MyHC-I MyHC-IIa MyHC-IIx/IId MyHC-IIb
Color Red Red White White
Contractile speed Slow Fast Fast Fast
Fatigue resistant High High Low Low
Metabolism Oxidative Oxidative Glycolytic Glycolytic
SDH activity High High Low Low
Mitochondria and myoglobin
content; CS activity High High Low Low
ATPase activiy Low Low High High
Mitochondrial CK and H-LDH High High Low Low
M-CK; M-LDH Low Low High High
Table 1: Physiological characters and metabolic activities of different fiber types in mouse skeletal muscle.
One type of slow-twitch fiber (type I) and three types of fast-twitch fibers (type IIa, type IIx/IId, and type IIb) exist in mouse skeletal muscle. Skeletal muscle fiber subtypes differ in enzyme expression, have distinct physiological characteristics and metabolic activity (Wang and Pessin, 2013). SDH, succinate dehydrogenase;
CS, citrate synthase; CK, creatine kinase; H-LDH, heart type lactate dehydrogenase; M-CK, muscle type creatine kinase; M-LDH, muscle type lactate dehydrogenase. Based on (Schiaffino and Reggiani, 2011) (Ciciliot et al., 2013) (Wang and Pessin, 2013).
Myokines Function
myostatin, IL-4, IL-6, IL-7, IL-15 Muscle hypertrophy
IL-6 Inflammation, insulin sensitivity
IL-6, BDNF Adipose tissue oxidation
IGF-1, FGF-2 Osteogenesis
IL-6, Irisin, myonectin Lipid metabolism
SPARC, Oncostatin M Anti-tumor defence, pancreas function
Table 2: Different muscular and systemic functions of myokines. IL, interleukin; BDNF, brain-derived neurotrophic factor; IGF-1, insulin like growth factor 1; FGF-2, fibroblast growth factor 2; SPARC, secreted protein, acidic, cysteine-rich. Based on (Pedersen and Febbraio, 2012) (Aoi and Sakuma, 2013).
Introduction
1.2 Intensive care unit-acquired weakness (ICUAW) 1.2.1 Definition of ICUAW and its consequences
During their stay in intensive care unit (ICU), critically ill patients often develop a significant loss of muscle weight leading to weakness (Saxena and Hodgson, 2012). These neuromuscular disorders are sub-classified into critical-illness myopathy (CIM), critical illness polyneuropathy (CIP), and critical illness neuromyopathy (CINM) (Appleton and Kinsella, 2012). CINM is a co-existing situation of CIM and CIP. CIP is diagnosed by electrophysiological testing (Appleton and Kinsella, 2012) whereas muscle biopsies are commonly used to confirm CIM (Schweickert, 2007). Depending on the ICU setting, method of assessment, patient population and time point of evaluation, the prevalence of ICUAW may vary. In general, 25% to 58% of ICU patients mechanically ventilated for more than one week develop ICUAW (Nordon-Craft et al., 2012). In certain ICU sub-groups, such as patients with sepsis, this percentage reaches 50% to 100% (Llano-Diez, 2012). However, the pathogenesis of ICUAW is unknown. It is further unclear why some patients develop ICUAW and others do not. Finally, how general inflammation and sepsis contribute to CIM is also uncertain.
Thanks to the developments in the treatment of critically ill patients, nowadays after initial stages of respiratory failure, circulatory shock, and severe infections many patients survive their critical illness, (Schefold et al., 2010). However, ICUAW prolongs ICU stay, increases morbidity and mortality of surviving patients (Sidiras et al., 2013). Approximately 45% of patients with ICUAW die during their stay in hospital and a further 20% mortality is expected in the first year after hospital discharge (Appleton and Kinsella, 2012) (Saxena and Hodgson, 2012). 28% of ICUAW patients develop severe disabilities such as quadriparesis, quadriplegia, or paraplegia during long-term follow-up (Schweickert, 2007). Impairment of muscle function may persist for up five years after hospital discharge and largely decreases the quality-of-life of survivors (Llano-Diez, 2012).
1.2.2 Risk factors for ICUAW
ICUAW has been attributed to multiple pathologies and its pathogenesis may vary from different patients (Deem, 2006). Several clinical observational studies have identified risk factors predisposing for ICUAW. It is generally agreed that systemic inflammation, nutritional deficiency, muscle inactivity, multiple organ failure, hyperglycemia, and application of corticosteroids and neuromuscular blockers are important contributors for ICUAW (Schefold
Introduction
et al., 2010) (Nordon-Craft et al., 2012). Here I am introducing the three most common and critical risk factors: sepsis, fasting, and immobilization.
● Sepsis
Sepsis is a leading cause of death and a pivotal risk factor for ICU patients. In the early stage of sepsis, cytokines TNF-α, interleukin IL-6 and IL-1 and complement factors (C3a and C5a) are released. In later stages anti-inflammatory cytokines (IL-10 and transforming growth factor beta-1, TGF-β1) are upregulated. Cytokines are involved in stimulation of the Ubiquitin-proteasome system (UPS) and proteolysis that lead to increased protein degradation. Moreover, decreased insulin sensitivity is often observed in sepsis patients. This may lead to deactivation of insulin-like growth factor-1 (IGF-1) signaling, which results in decreased protein synthesis and contributes to the loss of muscle weight in ICUAW (Schefold et al., 2010) (Deem, 2006).
● Fasting
Fasting is also considered as a major inducer of skeletal muscle atrophy of ICU patients who are not able to have normal diet. The mechanism of fasting-induced skeletal muscle atrophy is complicated. First of all, nutritional deficiency contributes to decreased muscle protein synthesis (Schefold et al., 2010). Secondly, fasting increases the UPS activity increasing protein degradation (Dehoux et al., 2004) (Paul et al., 2012). Moreover, fasting increases serum levels of corticosteroids, which also leads to enhanced UPS mediated protein degradation (Clarke et al., 2007) (Menconi et al., 2007).
● Immobilization
Complete or nearly complete limb immobilization contributes to loss of muscle weight and strength in ICU patients being mechanically ventilated (de Jonghe et al., 2009) (Bloch et al., 2011). It also causes a switch from slow, fatigue-resistant slow to fast fibers. Since fast fibers are more sensitive to inflammation-induced atrophy, this switch caused by immobilization renders the muscle more sensitive to inflammation-induced atrophy (Wang and Pessin, 2013).
Other mechanisms for immobilization-induced muscle weakness are elevated insulin resistance and increased apoptosis (Bloch et al., 2011).
Introduction
So far no medical therapy is shown to improve the recovery from ICUAW. Therefore, identification of ICUAW at early disease stages and prevention of disease development are important for ICUAW management (Appleton and Kinsella, 2012) (Puthucheary and Hart, 2009). Electrophysiological testing helps to differentiate ICUAW patients from patients having simple muscle degeneration (Nordon-Craft et al., 2012). New techniques, such as neuromuscular electrical stimulation (NMES) and cycle ergometry might provide new hope for early rehabilitation of affected patients (Stevens et al., 2009). Understanding the precise molecular mechanisms of ICUAW will provide new aspects and methods to intervene with disease progression. Therefore, different mouse models based on major risk factors have been established to study molecular mechanisms of ICUAW in vivo (Bloch et al., 2011). Here I will introduce these mouse models used in our group for investigating the different aspects of ICUAW.
1.2.3 Mouse models for studying ICUAW
Caecal ligation and puncture (CLP), exogenous administration of toxin (such as lipopolysaccharide) and pathogens (like bacteria) are commonly used to induce sepsis in mice (Buras et al., 2005). The CLP model is considered the gold standard for sepsis studies and was applied in our research group. It generates an intraabdominal (peritoneal) infection with mixed bacteria and provides an inflammatory source of necrotic tissue (Buras et al., 2005).
For starvation studies, we deprive food from mice for up to 48 h. For mimicking muscle disuse during immobilization a denervation mouse model was established. Ligation of the sciatic nerve results in disuse of the lower hind limb skeletal muscles in the operated mouse leg. Compared with the other two models, denervation induced atrophy is designed to study mechanisms of long-term skeletal muscle atrophy.
1.3 Skeletal muscle atrophy
Loss of skeletal muscle mass in ICUAW results from an imbalance between muscle protein synthesis and degradation (Batt et al., 2013). The underlying mechanism is multifactorial.
Upregulated muscle protein breakdown is mainly associated with the Ubiquitin-proteasome system (UPS), which is activated by many signalling molecules including TGF-β family members and the pro-inflammatory cytokine TNF-α in ICUAW. The downregulation of muscle protein synthesis is throught to be linked to impaired IGF-1/ V-Akt Murine Thymoma Viral Oncogene Homolog (Akt) signaling cascade (Bloch et al., 2011). Molecular pathways involved in skeletal muscle atrophy in ICUAW are introduced in this chapter.
Introduction
1.3.1 Skeletal muscle atrophy in ICUAW
Fiber-specific skeletal muscle atrophy is found in ICUAW patients, because skeletal muscle fiber subtypes have different sensitivities to atrophy signaling pathways under different pathophysiologic conditions (Figure 3) (Schefold et al., 2010) (Wang and Pessin, 2013). Slow (type I) fiber-specific atrophy and slow to fast (type II) fiber transition are mainly associated with muscle disuse (denervation and immobilization) (Wang and Pessin, 2013). Additionally, during the progressive reduction of exercise capacity, skeletal muscle dysfunction in chronic obstructive pulmonary disease (COPD) patients was identified as a result of slow to fast fiber-type shift (Gosker et al., 2002). This result also supported that slow fibers were more sensitive to atrophy induced by disuse of skeletal muscle. In contrast, fast fiber-specific atrophy is typically found in systemic diseases (i.e. sepsis, diabetes, cachexia, and chronic heart failure). This fiber type-specific muscle atrophy is found to be driven by proteins of the TGF-β family, Peroxisome proliferator-activated receptor gamma coactivator 1- alpha (PGC1α), Forkhead box o protein (FoxO) family, autophagy inhibition, and Nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB) signaling. In previous studies it was shown that TGF-β1 regulates atrophy of fast fibers (Mendias et al., 2012). Interestingly, overexpression of the TGF-β family member, myostatin, led to muscle atrophy an primarily of fast fibers (Wang and Pessin, 2013). Immunohistochemistry studies of muscle biopsies from CIM and CINM patients’ patients demonstrated a predominant atrophy of fast fibers during early critical illness (Schefold et al., 2010) (Bierbrauer et al., 2012). This indicated that sepsis was a more important atrophy risk factor compared with immobilization in the early stage of ICUAW. Skeletal muscle atrophy in the later stage of ICUAW might be more correlated to muscle disuse.
Figure 3: Metachromatic staining of histological sections from vastus lateralis muscle of a CIM (a subtype of ICUAW) and an ICU patient without CIM. Fiber-type-specific-stained patients’ skeletal muscle biopsies suggeste that fast (type II) fibers are more sensitive to sepsis-induced ICUAW. Compared with ICU control patients, sizes of type IIa and IIb fibers are much smaller in ICUAW patients. Atrophy in slow (type I) fibers are
Introduction
not as significant as in fast fibers. Slow (type I) fibers are in the darkest blue; fast (type IIa and IIb/IIx) fibers are in lighter blue. Adapted from (Schefold et al., 2010).
1.3.2 Molecular mechanisms involved in skeletal muscle atrophy 1.3.2.1 TGF-β pathway
The TGF-β family consists of more than thirty structurally related secreted polypeptide growth factors, which regulate a wide spectrum of cellular processes including cell proliferation, differentiation, motility, adhesion, and death (Massagué, 1998) (Zhang, 2008).
Based on sequence comparisons between their bioactive domains, the TGF-β family can be further grouped into the following subfamilies: TGF-β-, bone morphogenetic protein 2 (BMP2)-, BMP3- , BMP5- , growth/differentiation factor-5 (GDF-5)- , Vegetalising factor-1 (Vg1)- , activin-subfamilly (Massagué, 1998). Several TGF-β family members participate in regulation of skeletal muscle atrophy and fibrosis (Bloch et al., 2011) (MacDonald and Cohn, 2012). A bioinformatic meta-analysis of signaling events integrating multiple expression datasets revealed that signaling events in different muscle atrophy models, such as wasting, fasting, denervation; unloading and ageing, involve major networks including TGF-β pathway, NF-κB pathway, and apoptosis. An activation of the TGF-β pathway played a central role in different models of muscle atrophy, especially in the short-term muscle response to atrophy (within 14 days) (Calura et al., 2008).
● Canonical and non-canonical TGF-β pathways
TGF-β family members utilize a multitude of intracellular signal pathways. SMA/MAD- related (Smad) transcription factors (Smad2, Smad3) are very important downstream signal transducers in the TGF-β1 signal transduction pathway. Based on whether Smad2 and Smad3 are involved in TGF-β downstream signal transduction, TGF-β signaling is categorized as canonical and non-canonical pathway (Zhang, 2008).
In canonical TGF-β signaling transduction, ligand binding to type I and type II TGF-β receptors leads to phosphorylation and activation of Smad2/3 (Figure 4). In vivo experiments showed that overexpression of constantly activated type I and type II TGF-β receptors induced mouse skeletal muscle atrophy and proteasomal degradation-related gene F-box only protein 32 (Fbxo32) expression. This effect could be blocked by knockdown of Smad2 and Smad3. These results indicated that activation of canonical TGF-β signaling could lead to atrophy of adult myofibers (Sartori et al., 2009). Activated Smad2/3 translocate from the cytoplasm into nucleus and bind to regulatory elements of target gene promoters, and regulate
Introduction
gene expression together with other transcription factors, co-activators, and co-repressors (Figure 4). The runt domain transcription factor (RUNX) family members are among a few transcription factors which interact and co-function with Smads (Ito and Miyazono, 2003). Its strong induction in muscle shortly after denervation implied its function in regulating muscle atrophy. Indeed, muscle-specific inactivation of Runx1 led to more severe skeletal muscle atrophy indicated by myofibrillar disorganization, autophagy, and structual defects in denervated mice. Upregulation of atrophy genes Fbxo32 and Tripartite motif containing 63 (Trim63) in knockout mice suggested Runx1 was an important transcription co-factor that suppressed protein degradation pathways in muscle upon activation of canonical TGF-β signaling (Wang et al., 2005).
Non-canonical TGF-β pathways regulate a variety of signaling events independently of activation of Smad2 and Smad3 proteins. These non-Smad pathways include cross-talks with many other pathways such as: mitogen-activated protein kinase (MAPK) pathways, Rhodopsin (Rho)-like GTPase signaling pathways, and phosphatidylinositol-3-kinase/Akt pathways (Zhang, 2008).
Importantly, TGF-β family members regulate skeletal muscle atrophy through both canonical and non-canonical TGF-β pathways as introduced in following.
● Functions of TGF-β family members in muscle atrophy
TGF-β1 is the funding member of the TGF-β subfamily and induces skeletal muscle atrophy.
Muscle-specific overexpression of TGF-β1 induced skeletal muscle atrophy in mice (Narola et al., 2013). In addition, Mendias et al. reported that the in vivo injection of TGF-β1 decreased muscle fiber size and reduced maximum isomertric force generation of mouse skeletal muscle (Mendias et al., 2012). Clinical studies suggested that inhibition of TGF-β1 attenuates muscle atrophy. For example, compounds blunting TGF-β signaling through direct interacting with TGF-β1 or TGF-β receptors decreased skeletal muscle fibrosis, improved muscle regeneration, and restored muscle function. TGF-β1 neutralizing antibody prevented muscle atrophy and improved muscle regeneration (Burks and Cohn, 2011). Ki26894, an inhibitor of TGF-β receptor I (TGFβRI) increased muscle weight and strength in wild type mice, and restored muscle atrophy in a transgenic mouse model of dystrophy by promoting differentiation of myoblast (Ohsaw et al., 2012). Unfortunately, in vitro studies of TGF-β1- induced skeletal muscle atrophy are limited and its underlying molecular mechanism is far
Introduction
from being understood. In my study, using differentiated C2C12 myotubes as an in vitro model, I demonstrated that TGF-β1 directly induced muscle atrophy.
Another evidence supporting TGF-β1’s function in skeletal muscle atrophy is the correlation between upregulated TGF-β1 expression and muscle atrophy induced in dystrophy patients and denervation mouse model (Bernasconi et al., 1995) (Bernasconi et al., 1999) (Ishitobi et al., 2000) (Gosselin et al., 2004) (Ozawa et al., 2013). TGF-β1 serum levels were shown to be increased critically ill patients with sepsis (Schulte et al., 2013). Therefore, it is likely that TGF-β1 contributes to skeletal muscle atrophy in ICUAW developed in critically ill patients.
The microarray study of ICUAW patients’ samples performed by our collaborators provided a novel aspect of activated TGF-β1 signaling in the disease process.
Among all TGF-β family members, myostatin (also known as GDF-8) is the best characterized skeletal muscle atrophy inducer. Myostatin is known as a negative regulator of differentiation and growth of muscle tissue. It triggers downstream events via both canonical and non-canonical TGF-β signaling pathways. Activation of Smad2/3 and Smad4 upon myostatin binding to its receptor (activin receptor IIB; ActRIIB) downregulated expression of genes involved in myogenesis, such as Myogenic differentiation (MyoD), Myogenic factor 5 (Myf5), and Myogenin (Elkina et al., 2011). Moreover, myostatin suppressed muscle differentiation by activating MAPK1/3 via the Rat sarcoma (Ras)/ Rapidly accelerated fibrosarcoma (Raf)/ Mitogen-activated protein kinase kinase 1 (MAP2K1) pathway and inhibiting the Akt/ Mammalian target of rapamycin (mTOR) pathway, which promoted protein synthesis (Elkina et al., 2011). Administration of myostatin resulted in a loss of body weight, and reduced number and size of myotubes in mouse skeletal muscle in vitro. The mechanisms involved downregulation of MyoD and Paired box 3 (pax3), and upregulation of atrophy-related genes (Fbxo32, Trim63, and E214k) in vivo through AKT-FoxO1 pathway.
Also in vitro, Trendelenburg el al. showed inhibition of myoblast differentiation and decreased myotube size after treatment of myostatin through reducing Smad2/3-dependent Akt/TORC1/p70S6K signaling. This observation was consistent in in vitro culture of human skeletal myotubes as well. Administration of lower dose of myostatin decreased expression of differentiation-related genes (myogenin and MyoD) and activity of creatine kinase (CK), and reduced expression of atrophy genes (FBXO32 and TRIM63) in differentiated human skeletal myotubes (Trendelenburg et al., 2009).
Introduction
In contrast to functions of TGF-β1 and myostatin in inducing muscle atrophy, BMP family members and activated BMP signaling are involved in prevention of muscle wasting through a non-canonical TGF-β signaling pathway (Sartori et al., 2013) (Winbanks et al., 2013).
Inhibition of BMP signaling caused muscle atrophy, and abolished the hypertrophic phenotype of myostatin-deficient mice through activating of Smad1/5/8 and downregulating of Fbox30 (Sartori et al., 2013). In another independent study, activation of BMP signaling was detected by increased phosphorylation of Smad1/5. Usage of BMP signaling inhibitor Smad6 exacerbated denervation-induced skeletal muscle atrophy by restoring expression of histone deacetylase 4 (HDAC4) and activation of proteolysis (Figure 5) (Winbanks et al., 2013).
Figure 4: Canonical TGF-β signaling. Binding of TGF-β family members (includes members from TGF-β subfamily, activin subfamily, and BMP subfamily), with their corresponding type II receptors (includes TGF-β recptor II, BMP receptor II, and anti-Mullerian hormone recptor) leads to phosphorylation of type I receptors, which then phosphorylate type I receptors (includes TGF-β receptor I, activin receptor IB, BMP recptor IA/IB, and activin recptor-like kinase 1/2/7). Activated type I and type II receptor complexes subsequently phosphorylate receptor-regulated Smads (R-Smads) (Smad2 or Smad3). Activated R-Smads bind to Smad4 and move into the nucleus, where Smad proteins regulate target gene transcription along with other transcription factors. Adapted from (Massagué, 1998).
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Figure 5: Proposed schema of function of BMP signaling in denervation-induced skeletal muscle atrophy.
In skeletal muscles undergoing neurogenic atrophy, expression of BMP13 and BMP14 were upregulated and activated the downstream signaling, which led to an increased Smad1/5/8 complex phosphorylation. Activated Smad1/5/8 complex bound to Smad4, and potentially abolished the inhibitory effect of HDAC4 on atrophy gene expression. Adapted from (Winbanks et al., 2013).
Introduction
● TGF-β1 signaling inhibits gene transcription
Previous studies have implied a role of TGF-β1 in negative regulation of gene transcription in regulating skeletal muscle differentiation, fibrosis, and regeneration. For example, TGF-β1 treatment impaires differentiation of C2C12 cells, and largely reduces expression of myosin heavy chain during differentiation of C2C12 myocytes (Wicik et al., 2010). Microarray analysis of gene expression profiles of TGF-β1-treated C2C12 myoblasts showed that among 502 significantly regulated genes, 436 were downregulated (Wicik et al., 2010). The TGF-β- 1-ihibibted gene transcription is resulted from binding of transcription factors to target gene promoters. The first TGF-β inhibitory element (TIE) (5’-GNNTTGGNGA-3’) was found in the promoter of the collagenase stromelysin, which promoted fibrosis in damaged muscle (Serrano et al., 2011). Moreover, TIE-like elements were also identified in promoter of c-myc (Chen et al., 2001) (Frederick et al., 2004), which inhibited myogenic differentiation in muscle (Miner and Wold, 1991). Mechanism of TGF-β1-inhibited c-myc expression was depend on activation of canonical TGF-β1 signaling (Chen et al., 2001) (Frederick et al., 2004).
In our study, microarray results implied that both TGF-β1 induced and reduced gene expression were equally important in skeletal muscle of ICUAW patients. Since information about TGF-β1-inhibited gene expression in skeletal muscle atrophy is very limited, we were encouraged to identify and characterize TGF-β1-downregulated target genes in muscle atrophy.
1.3.2.2 Proteolytic pathways and protein degradation
Mechanisms of skeletal muscle atrophy involve a significant increase in protein degradation (Glass, 2005). The UPS largely contributes to protein breakdown in muscle. The UPS is composed of ubiquitin-activating enzymes (E1), ubiquitin-conjugating enzymes (E2), and ubiquitin-protein ligases (E3), and the 26S proteasome. Ubiquitin is first activated by E1 in an ATP-dependent manner. Activated ubiquitin is then transferred to the E2. E2 binds to E3, which attaches ubiquitin to substrate proteins. Ubiquitinated proteins are than degraded by the 26S proteasome (Chopard et al., 2009). Risk factors of ICUAW such as muscle inactivity, inflammation, and food deprivation are shown to upregulate the transcription factor forkhead box O (FoxO) which stimulates expression of the E3 ubiquitin ligase tripartite motif containing 63 (Trim63; also named muscle RING finger protein-1, MuRF-1) and the F-box adaptor protein F-box only protein 32 (Fbxo32; also called atrogin-1) (Batt et al., 2013).
Introduction
Upregulation of these two atrophy genes are often used as key indicators for skeletal muscle atrophy (Glass, 2005). But this does not apply to all atrophy models. For example, in a denervation-induced skeletal muscle atrophy mouse model, upregulation of these two genes was not identified (Sartori et al., 2013). This indicated possible functional redundancy of UPS-related E3 ligases genes. It is also possible that other mechanisms such as decreased protein synthesis are more important to muscle atrophy induced by certain stimuli.
1.3.2.3 Functions of Akt signaling pathway in regulating protein synthesis and degradation Decreased protein synthesis is another major cause for skeletal muscle atrophy (Appleton and Kinsella, 2012). Akt is the key signaling pathway regulating muscle weight by influencing protein synthesis. It also represses protein degradation by inhibition of atrophy gene expression (Schiaffino and Mammucari, 2011) (Ochala et al., 2011). Binding of IGF-1 or insulin with insulin receptor (IR) leads to phosphorylation of insulin receptor substrate (IRS).
Decreased IGF-1 serum levels are responsible for impaired Akt signaling in ICUAW (Bloch et al., 2011). Activated IRS recruits and activates phosphatidylinositol-3-kinase (PI3K) which in turn generates phosphoinositide-3,4,5-trisphosphate (PIP3). PIP3 acts as a docking site for phosphoinositide-dependent kinase 1 (PDK1) and Akt. Phosphorylation of Akt at one threonine residue by PDK1 and one serine site by mammalian target of rapamycin complex-2 (mTORC2) leads to its activation. Activated Akt promotes protein synthesis and inhibits protein degradation through three downstream pathways by phosphorylation of mTORC1, Glycogen synthase kinase 3 beta (GSK3-β), and FoxO proteins. The activation of mTORC1 leads to phosphorylation of eukaryotic initiation factor 4e binding protein 1 (4eBP1) and ribosomal protein S6 kinase (S6K1). 4eBP1 is an inhibitor of eukaryotic initiation factor 4e (eIF4e). Phosphorylation of 4eBP1 leads to its inactivation and in turn releases eIF4e and stimulates protein synthesis. Phosphorylated S6K1 further activates ribosomal protein S6, which inducing protein anabolism. Phosphorylation of GSK3-β leads to its inhibition and further activation of translation initiation factor eIF2b, which also stimulates protein synthesis. Activation of Akt signaling pathway inhibits protein degradation by phosphorylating and inhibiting the FoxO transcription factors. FoxO proteins are transcription factors which bind to target promoters via the consensus site TTGTTAC (Chopard et al., 2009). Inactivated FoxO proteins translocate from the nucleus to the cytoplasm and indeed cause decreased transcription of atrophy-associated genes (atrogenes) such as Trim63 and Fbxo32. Moreover, FoxO proteins directly downregulate mTOR (Figure 6) (Schakman et al., 2008) (Schiaffino and Mammucari, 2011) (Llano-Diez, 2012).
Introduction
Figure 6: Akt pathway in regulation of protein synthesis and degradation in skeletal muscle atrophy. IGF- 1 activates Akt pathways and influences on both protein synthesis and protein degradation. Phosphorylation leads to activation of mTORC1 and inhibition of GSK3-β, which both result in stimulated protein synthesis.
Phosphorylation of FoxO proteins by Akt leads to deactivation of these transcription factors, which induce expression of atrogenes (Trim63 and Fbxo32). Based on (Schiaffino and Mammucari, 2011).
IGF1 IGF1 receptor
IRS PI3K PDK1
Akt
mTORC1 GSK3-β FoxO1, FoxO3
Trim63, Fbxo32
Protein synthesis
3 Fb 1 rec
IRS PI3K PDK
C1 GSK3-β
eIF2bF2 mTORC2
β 4eBP1 S6K1
Introduction
1.4 Secreted Frizzled Related Proteins (SFRPs) 1.4.1 SFRP family
Human and mouse Secreted Frizzled Related Protein (SFRP) family comprises five secreted proteins, SFRP1-5 (Cruciat and Niehrs, 2012). They received their name because they have an N-terminal cysteine rich domain (CRD) homologous to the extracellular Wingless-Type MMTV Integration Site Family (Wnt) binding domain of Frizzled receptors. They also share a C-terminal Netrin-Related (NTR) motif. This motif was found in other unrelated proteins, such as Netrin-1, tissue inhibitors of metalloproteinase (TIMPs), complement proteins, and typeI procollagen C-proteinase enhancer proteins (PCOLCEs) (Figure 7) (Esteve and Bovolenta, 2010).
1.4.2 Functions of Sfrp2
From microarray data, we identified significant transcriptional downregulation of SFRP family members (SFRP1, SFRP2, and SFRP4) in muscle biopsies of ICUAW patients implicating a function of thisgene family in the pathophysiology of this disease. Among these three family members, SFRP2 had the strongest fold change in patients and was chosen for further analysis.
Function of Sfrp2 in mature skeletal muscle is largely unknown, whereas its functions in regulating myogenesis, myocardial fibrosis and regeneration have been addressed in previous studies (Descamps et al., 2008) (Kobayashi et al., 2008) (Alfaro et al., 2010) (Snyder et al., 2012). Recombinant Sfrp2 impaired myotube formation of differentiating C2C12 myoblasts and primary satellite cell cultures by decreasing myogenin and myogenic regulatory factor 4 (Mrf4) expression (Descamps et al., 2008). Moreover, WNT3A and WNT5A rescued impaired differentiation of C2C12 cells induced by upregulation of Sfrp2 (Snyder et al., 2012). This indicated that inhibition of myogenesis by Sfrp2 was depending on inhibition of Wnt signaling. As an important regulator of Wnt signaling, Sfrp2 is involved in cardiac regeneration (Alfaro et al., 2010). Sfrp2 was shown to mediate mesenchymal stem cell (MSC) proliferation in vitro and in vivo. Treatment with recombinant Sfrp2 or overexpression of Sfrp2 decreased MSC apoptosis and inhibited both osteogenic and chondrogenic lineage commitment through inhibition of BMP and Wnt signaling. These data indicated that Sfrp2 promoted MSC-driven myocardial and wound repair. However, whether these inhibitory effects involved direct binding between Sfrp2 and BMP/Wnt proteins was not known (Alfaro et al., 2010).
Introduction
Sfrp proteins also function as evolutionarily conserved antagonists of chordinase proteinases in zebrafish, Xenopus and Drosophila melanogaster (Kobayashi et al., 2008). Sizzled, the orthologous Sfrp gene in Xenopus laevis and zebrafish was shown to function as an antagonist of chordin processing by tolloid-like metalloproteinases. Mammalian Sfrp2 had no proteinase activity towards chordin. Instead, Sfrp2 served as a direct binding partner and enhancer of procollagen C proteinase activity of BMP1. In Sfrp2-knockout mice subjected to myocardial infarction, cardiac fibrosis was remarkably reduced (Kobayashi et al., 2008).
Considering the fact that the protein sequences of growth factor domains of TGF-β family members are highly conserved (Shi et al., 2011), I assumed that the three-dimensional structures of TGF-β family members could also be homologous. By comparing crystal structures of human BMP1 and BMP3 (Allendorph et al., 2007), and solution structure of human TGF-β1 (Hinck et al., 1996), I found these three structures to be very similar (Figure 8). Since Sfrp2 can directly interact with BMP1, it is likely that it also binds to TGF-β1 and regulates TGF-β1 signaling. Also very interestingly, function of another SFRP family member in regulating TGF-β1 has been described (Gauger et al., 2011). In immortalized non- malignant mammary epithelial cells, knockdown of SFRP1 led to an increased sensitivity to TGF-β1 signaling indicated by upregulation of TGF-β1 target genes and increased phosphorylation of mitogen-activated protein kinase 1 (MAPK1) and MAPK3 (Gauger et al., 2011). However, it was not known if the inhibitory effect of SFRP1 on TGF-β1 signaling was due to direct binding between these two proteins, or due to secondary effects resulting from alternative SFRP1 signaling events. In my study, I hypothesized that Sfrp2 directly interacts with TGF-β1, and had an inhibitory effect on TGF-β1-induced muscle atrophy.
1.4.3 Mechanisms of Sfrp2 downregulation
Previous studies suggest microRNAs (miRNA) and promoter hypermethylation are mechanisms for Sfrp2 transcriptional repression. In C2C12 myoblasts, a subset of miRNA cluster regulated myocyte enhancer factor 2A (Mef2A) transcription factor repressed expression of Sfrp family members, including Sfrp2. Knockdown of Mef2A led to upregulation of Sfrp1, 2, and 4 and impaired myotube formation (Snyder et al., 2012).
Downregulation of SFRP2 has been observed in several human invasive carcinomas.
Interestingly, SFRP2 promoter hypermethylation seems to occur in the early onset of many tumors, such as breast, gastric, cervix, hepatocellular, pancreatic, lung, ovary, renal, prostate,
Introduction
and colon carcinomas. This suggests a correlation between SFRP2 promoter hypermethylation and its epigenetic silencing in cancers (Esteve and Bovolenta, 2010).
So far, little is known for the regulation of Sfrp2 in muscle diseases. Our group first identified it as a downregulated target gene of activated TGF-β1 signaling in ICUAW. In this study, I tested whether transcriptional regulation led to repression of Sfrp2 by TGF-β1, and whether its downregulation in skeletal muscle atrophy was correlated with its promoter hypermethylation.
Figure 7: Common domain structure of SFRP family members. Signal peptide is shown in orange. CRD, Cysteine rich domain (Blue); NTR, Netrin-related motif (Purple).
A B
C
Figure 8: Three-dimensional structures of human BMP3, BMP6, and TGF-β1. (A) Cristal structure of human BMP3 (Protein Data Bank (PDB) identifier: 2QCQ) (Allendorph et al., 2007) (This figure is downloaded from http://www.ebi.ac.uk). (B) Crystal structure of human BMP6 (PDB identifier: 2QCW) (Allendorph et al.,
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2007) (This figure is downloaded from http://www.ebi.ac.uk). (C) Solution structure of human TGF-β1 (Hinck et al., 1996) (PDB identifier: 1KLA) (This figure is downloaded from http://www.rcsb.org).
1.5 Aim of this study
The cause of ICUAW is multifactorial and remains largely unknown. Sepsis, immobility, administration of corticosteroids and neuromuscular blocking agents are suggested as common risk factors (de Jonghe et al., 2009). But clinical studies attempting to isolate and study effects of individual factors often showed variable and contradictory results (de Jonghe et al., 2009) (Stevens et al., 2009). These might be due to heterogeneity in patients’
conditions, medical treatments, or time points of data collection. A previous microarray study investigating differences in muscular gene expression between critically ill patients with CIM, without CIM and control patients had provided us the first glance at altered signaling pathways in ICUAW, and suggested an important role of TGF-β signaling (Di Giovanni et al., 2004). Compared to previous studies, our microarray study included a more comprehensive collection of ICUAW patient samples to generate a more general and complete view of molecular signaling pathways in this disease. Our data indicate activation of TGF-β1 signaling in ICUAW patients and revealed regulated downstream target genes. Following our microarray study, there are two major aims of my project for studying activated TGF-β1 signaling in skeletal muscle atrophy. The first aim is to investigate the phenotype and the molecular mechanism underlying loss of myosin heavy chain in TGF-β1-induced skeletal muscle atrophy in vitro. Analysis of our microarray data identified SFRP2 as the most regulated TGF-β1 downstream target in ICUAW patients. This result implies that Sfrp2 could be very important in conducting TGF-β1 signaling which leads to muscle atrophy. Therefore, my second aim is to characterize the regulation and function of Sfrp2 in skeletal muscle atrophy.
