I MOLECULAR MECHANISMS of NF-Κb REGULATION OF

MOLECULAR MECHANISMS OF NF-κB REGULATION OF SKELETAL

MYOGENESIS

DISSERTATION

Presented in Partial Fulfillment of the Requirements for

the Degree Doctor of Philosophy in the Graduate

School of The Ohio State University

Nadine A. Bakkar, M.S.

The Ohio State University 2008

Dissertation Committee: Approved by

Denis Guttridge, PhD, Advisor

Jill Rafael-Fortney, PhD ______

Gustavo Leone, PhD Advisor

Michael Ostrowski, PhD Graduate Program in Molecular, Cellular

and Developmental Biology

ABSTRACT

NF-κB is a ubiquitous transcription factor involved in the regulation of innate immunity, cellular survival, proliferation, as well as differentiation. Its deregulation is associated with various diseases, and have thus been the target of developing therapeutic strategies. Skeletal muscle diseases are one field where this transcription factor is receiving recent attention, owing to its implication in muscular dystrophy, wasting and regeneration. In this dissertation, we focused on NF-κB regulation of myogenic differentiation, in an attempt to further understand the complex ways this transcription factor follows to regulate muscle development and extrapolate it to disease.

In chapter 2, we focused on Myostatin (Mstn) a potent negative regulator of myogenesis that can inhibit myoblast proliferation and suppress synthesis of MyoD. NF-κB is similarly able to promote myoblast growth and induce loss of MyoD message. Given the similarities of these phenotypes, we examined potential Mstn and NF-κB signaling crosstalks in myoblasts and differentiated myotubes. Results show that Mstn does not activate NF-κB, nor does activated

NF-κB induce Mstn expression. Furthermore, Mstn inhibition of differentiation can still occur in cells devoid of NF-κB activity. Such findings were confirmed in proliferating muscle precursors as well as mature muscle fibers and thus highlight the intrinsic differences between those two signaling pathways in the regulation of skeletal myogenesis,

In chapter 3, we examined NF-κB regulation of skeletal myogenesis using genetic

knockout models. Previous studies had attempted to investigate such a regulation, yet results

remained perplexing, with both pro- and anti-myogenic roles of NF-κB documented. Using

primary myoblasts and muscles devoid of NF-κB classical pathway components p65 or IKKβ,

we show that this canonical signaling is a negative regulator of myogenesis and gets

downregulated during differentiation. On the other hand, NF-κB alternative signaling, mediated

by IKKα activating the RelB/p52 complex is turned on and regulates mitochondrial biogenesis.

Such a pathway is hence involved in the energy production and subsequent maintenance of

newly formed myotubes. Consequently, our findings help to resolve the conundrum of NF-κB signaling in myogenesis by showing the existence of two opposing NF-κB pathways that

function at temporally distinct stages of differentiation: classical signals inhibit premature

myogenesis while alternative pathway activation regulate energy production and maintenance of

nascent myofibers.

Collectively, results presented in this dissertation highlight the various branches through

which NF-κB signals to regulate skeletal myogenesis, emphasizing the need to take this complex

regulation into account in clinical strategies aimed to modulate its activity.

iii

Dedicated to my family and friends.

ACKNOWLEDGMENTS

First, I would like to thank my advisor, Denis Guttridge. I think I was pretty lucky in

having you as a benchmate/lab partner during the first year that I joined sharing experiment tips

as well as restaurant reviews, political discussions, french lessons, or independent movie

recommendations. And although you later moved to your office, I am still able to chat with you

about anything. More importantly, you’ve taught me to think outside “the box” and to keep an

open mind and question known dogmas. And although I hate to admit it in public to my friends,

I do enjoy getting new data to step into your office and discuss them with you. You’ve helped

me grow as a scientist while making sure I keep a healthy balance between scientific career and personal life.

To past and present members of the Guttridge lab, it’s been a real pleasure working with you guys as well as organizing floor parties and going out. You made me feel like I belong to a group and we have shared more than I could ever describe, from nerdy science talks over lunch to teaching me american slang and shopping . Thank you for making the work atmosphere so enjoyable. Special thanks go to Erin, Jay and Mike for being above all my first American friends; Jingxin, Kate, and Huating for their constant guidance; Jen, Tara, Wei, Jeff, Swarnali,

Lori and Erik for all the good times and the laughs in the lab and outside.

I would also like to thank Drs Jill Rafael-Fortney, Gustavo Leone and Michael Ostrowski for taking time out of their busy schedules to serve on my committee, as well as their helpful advice.

To all my friends for always being there and making Columbus my home away from home; thank you.. Ihab, I cannot thank you enough for your unconditional and constant support;

Sama, Joe, Danielle, Sleiman, Mirna, Fadi, thank you for always being there to share the good times and the bad ones...Francisco, thank you for all the laughs, coffee breaks and Spanish lessons…Myrna and Lyne, even though you are not physically around, thank you for being

“there”…To all my friends all over the world, Hilda, Wassim, Nesrine, Hisham, Mazen to name a few, thank you for being my friends…

And finally, I cannot but thank my mother Nawal, my father Ali and my brother Walid for believing in me, supporting me in every possible way, and always ALWAYS being proud of me.

VITA

May 8th, 1978………………………………Born- Beirut, Lebanon

June 1999…………………………………..Bachelor of Science in Biology The American University of Beirut Beirut, Lebanon

June 2001 ……………………………...... Master of Science in Biology The American University of Beirut Beirut, Lebanon

Sept 2001-Aug 2002……………………….Research Associate, Dept of Pediatrics The American University of Beirut Beirut, Lebanon

Sept 2002-Present……………………… PhD Candidate, Molecular Cellular and Developmental Biology Graduate Program, The Ohio State University, Ohio, USA

PUBLICATIONS

Bakkar N, Wang J, Ladner KJ, Wang H, Dahlman JM, Carathers M, Acharyya S, Rudnicki MA, Hollenbach AD, Guttridge DC. IKK/NF-kappaB regulates skeletal myogenesis via a signaling switch to inhibit differentiation and promote mitochondrial biogenesis. J Cell Biol. 2008 Feb 25;180(4):787-802.

Wang H, Hertlein E, Bakkar N, Sun H, Acharyya S, Wang J, Carathers M, Davuluri R, Guttridge DC.NF-kappaB regulation of YY1 inhibits skeletal myogenesis through transcriptional silencing of myofibrillar genes. Mol Cell Biol. 2007 Jun;27(12):4374-87.

vii

Acharyya S, Villalta SA, Bakkar N, Bupha-Intr T, Janssen PM, Carathers M, Li ZW, Beg AA, Ghosh S, Sahenk Z, Weinstein M, Gardner KL, Rafael-Fortney JA, Karin M, Tidball JG, Baldwin AS, Guttridge DC. Interplay of IKK/NF-kappaB signaling in macrophages and myofibers promotes muscle degeneration in Duchenne muscular dystrophy. J Clin Invest. 2007 Apr;117(4):889-901.

Bakkar N, Wackerhage H, Guttridge DC. Myostatin and NF-κB regulate skeletal myogenesis through distinct signaling pathways. Signal Trasduction. 2005 4: 202-210.

Hertlein E, Wang J, Ladner KJ, Bakkar N, Guttridge DC. RelA/p65 regulation of IkappaBbeta. Mol Cell Biol. 2005 Jun;25(12):4956-68.

Mikati MA, Holmes GL, Werner S, Bakkar N, Carmant L, Liu Z, Stafstrom CE. Effects of nimodipine on the behavioral sequalae of experimental status epilepticus in prepubescent rats. Epilepsy Behav. 2004 Apr;5(2):168-74.

Mikati MA, Shamseddine A, Sabban M, Dbaibo G, Kurdi R, Abi Habib R, and Bakkar N. Time course of changes in apoptotic signal transduction factors during and after experimental status epilepticus. AES Proceedings. Epilepsia 2002. Vol 43 s7:1-375. Abstract 1.044

Gali-Muhtasib H, Bakkar N. Modulating cell cycle: current applications and prospects for future drug development. Curr Cancer Drug Targets. 2002 Dec;2(4):309-36.

FIELD OF STUDY

Major Field: Molecular, Cellular and Developmental Biology

viii

TABLE OF CONTENTS

ABSTRACT...... ii

ACKNOWLEDGMENTS ...... v

VITA...... vii

LIST OF FIGURES ...... xi

LIST OF ABBREVIATIONS...... xiii

CHAPTER 1 ...... 1

INTRODUCTION ...... 1 1.1 NF-κB Signaling...... 1 NF-κB and IκB Family Members...... 1 The IKK Complex...... 4 Upstream Activators of NF-κB...... 5 1.2 Classical and Alternative NF-κB Signaling...... 6 1.3 NF-κB and cellular differentiation...... 8 Osteoclastogenesis ...... 8 Hematopoiesis...... 9 1.4 Skeletal Muscle Differentiation ...... 10 Description and Regulation...... 10 Myostatin in Myogenesis...... 12 1.5 NF-κB in Myogenesis and Muscle Regeneration...... 13 NF-κB as a Myogenic Activator...... 14 NF-κB as a Myogenic Inhibitor ...... 15 ix

Model for NF-κB Regulation of Myogenic Differentiation ...... 17 NF-κB Regulation of Muscle Regeneration ...... 18 1.6 Mitochondrial Biogenesis ...... 20 Mitochondria: An Overview ...... 20 Mitochondrial Biogenesis ...... 22 Mitochondrial Biogenesis in Skeletal Muscle ...... 24 CHAPTER 2 ...... 29

MYOSTATIN AND NF-κB REGULATE SKELETAL MYOGENESIS THROUGH DISTINCT SIGNALING PATHWAYS ...... 29 2.1 Introduction...... 29 2.2 Materials and Methods...... 32 2.3 Results...... 33 2.4 Discussion...... 36 CHAPTER 3 ...... 44

IKK/NF-κB REGULATES SKELETAL MYOGENESIS VIA A SIGNALING SWITCH.. 44 3.1 Introduction...... 44 3.2 Results...... 47 3.3 Discussion...... 59 3.4 Materials and Methods...... 64 REFERENCES ...... 81

LIST OF FIGURES

Figure 1.1: NF-κB and IκB family members...... 26

Figure 1.2: NF-κB signaling pathways...... 27

Figure 1.3: Mitochondrial biogenesis...... 28

Figure 2.1: Mstn causes a minor activation of NF-κB DNA binding activity in differentiating myoblasts...... 39

Figure 2.2: Mstn does not regulate NF-κB transcriptional activity...... 40

Figure 2.3: Endogenous activity of NF-κB is not required for Mstn inhibition of myogenesis... 41

Figure 2.4: Mstn causes a transient increase in NF-κB DNA binding activity in myotubes...... 42

Figure 2.5: TNFα activation of NF-κB does not induce Mstn expression C2C12 myoblasts...... 43

Figure 3.1: Loss of p65 enhances myogenic activity in MEFs...... 68

Figure 3.2: Loss of p65 accelerates the myogenic program in MEFs...... 69

Figure 3.3: Loss of p65 enhances differentiation of primary myoblasts...... 70

Figure 3.4: Myogenesis is enhanced in p65 deficient mice...... 71

Figure 3.5: p65 regulation of myogenesis occurs through multiple mechanisms...... 72

Figure 3.6: IKK signaling is temporally regulated and functionally distinct in myogenesis...... 73

Figure 3.7: IKKα regulates myotube maintenance...... 74

Figure 3.8: IKKα regulates mitochondrial biogenesis...... 75

Figure 3.9: IKKα controls mitochondrial structure...... 76

Figure 3.10: A model for IKK/NF-κB signaling and function in skeletal myogenesis...... 77

Figure 3.11: Lack of p65 leads to increased fiber numbers independent of age, fiber type or muscle atrophy...... 78

Figure 3.12: Conditional deletion of IKKβ in muscles...... 79

Figure 3.13: Mitochondrial inhibitors lead to myotube cell death...... 80

xii

LIST OF ABBREVIATIONS

ATP……………………………….………….adenosine triphosphate

BAFF…………………………….…………...B-cell activating factor

CBP...…………………………….…………..CREB binding protein

CSF…………………………………………...Colony stimulating factor

DNA………………………………………….deoxyribonucleic acid

ds….………………………………………….double stranded

EMSA..……………………………………....electrophoretic mobility shift assay

HAT…………………………….………….....histone acetylase

HDAC…………………………..……………histone deacetylase

HLH………………………………………….helix-loop-helix

IκB…………………………………………...inhibitor of kappa B

IKK…………………………………………..IkB kinase

IGF……………………………………………Insulin-like growth factor

IL...…………………………………………...interleukin

LPS……………………………………….…..lipopolysaccharide

LT-βR…………………………….…………..lymphotoxin beta receptor

MEF…………………………….…………....mouse embryo fibroblast

Mstn…………………………………………..myostatin

xiii

MyHC.…………………………….………....myosin heavy chain

NEMO…………………………….………….NF-κB essential modulator

NES…………………………….………….... nuclear export signal

NF-κB………………………………………..nuclear factor kappa B

NIK…………………………………………..NF-κB inducing kinase

NLS…………………………………………..nuclear localization signal

PCR……………………………………….….polymerase chain reaction

PKA……………………………………….…protein kinase A

PDTC……………………………………….. pyrrolidine dithiocarbamate

RHD………………………………………….rel homology domain

RIP……………………………………………receptor interacting protein

RNA………………………………………….ribonucleic acid

SUMO.……………………………………….small ubiquitin-related modifier

TLR…………………………………………..Toll-like receptor

TRAF…………………………………………TNF receptor associated factor

TRADD……………………………………….TNF receptor associated death domain

YY1………………………………………….Yinyang 1

xiv

CHAPTER 1

INTRODUCTION

1.1 NF-κB Signaling

First identified as a transcription factor important for the activation of κ light chain genes in B cells, NF-κB is now recognized as a ubiquitously expressed factor involved in the regulation of a wide array of pathways such as cell survival, proliferation and differentiation, as well as immune responses [1, 2]. This family of proteins includes several members, and can be activated by a variety of signals, adding to the complexity of its regulation.

The past decade has shown an increased interest in the role of NF-κB in the regulation of skeletal muscle differentiation, driven by outstanding findings in the fields of stem cell biology as well as muscular dystrophy and muscle diseases. We have attempted in this literature review to cover recent studies pertaining to the role of NF-κB in myogenesis, as well as its involvement in muscle regeneration.

NF-κB and IκB Family Members

NF-κB is a family of dimeric evolutionary conserved proteins including five members: RelA/p65, RelB, c-Rel, NF-κB1/p50 (and its precursor p105), and NF-

κB2/p52 (and its precursor p100). These proteins share a 300 amino acid Rel homology

domain (RHD) composed of two immunoglobulin-like repeats that is responsible for

DNA-binding, dimerization, nuclear translocation, as well as binding to the NF-κB inhibitors, the IκB family members (Figure 1.1). NF-κB members can bind with

different affinities to sites bearing the consensus sequence GGGRNNYYCC, where R is a purine, Y is a pyrimidine and N is any base [3]. The degenerate nature of this binding sequence and the diverse binding preferences of the NF-κB dimers leads to the recruitment of various coactivators and corepressors and results in the expression of a wide variety of target genes [4]. Transcription of these target genes is further regulated through post-translational modifications of NF-κB that affect its interaction with

transcriptional modulators. p65 has been shown to be phosphorylated on different sites

such as Ser276, Ser529 and Ser536 by various kinases including Protein kinase A (PKA),

Mitogen and stress activated protein kinase 1 (MSK1), Tank bindink kinase 1 (TBK1) or

Casein kinase 2 (CK2 {Campbell, 2004 #922; O'Shea, 2008 #921}). Such

phosphorylation events mainly function to increase p65 transcriptional activity either by

enhancing binding to coactivator proteins and the transcriptional machinery or increased

nuclear localization and stability, although phosphorylation of sites such as Ser468 can be

inhibitory {Buss, 2004 #923}. p65 has been reported to undergo other postranslational

modifications such as acetylation at Lys122, Lys 218, and Lys310 by p300/CBP and

other proteins with HAT activity to enhance its activity and ubiquitination by SOCS-1 ubiquitin ligase between residues 220 and 335 targeting p65 for proteasomal degradation

{Ryo, 2003 #924}[3]. NF-κB proteins can exist as homo or hetero-dimers that function

mostly as transcriptional activators although the p50/p50 and p52/p52 complexes are

essentially repressors of gene activation [5]. Such a distinction is because both p50 and p52 lack a transactivation domain. They are instead characterized in their precursor

forms (p105 and p100, respectively) by the presence of ankyrin repeats and thus are also

considered as members of the ΙκΒ family (Figure 1.1).

The IκB family of NF-κB inhibitors include several members namely IκBα,

IκBβ, IκBγ, IκBε, IκBζ as well as Bcl-3 [3]. IκBα, β, ε and the precursor proteins p100

and p105 are characterized by a core of 6 or more ankyrin repeats that allow them to

interact with the RHD of NF-κB members [6]. Such interaction with NF-κB dimers

masks their nuclear localization signal (NLS), retaining the complex in the cytoplasm of

unstimulated cells. It is noteworthy that the p65/p50/IκBα complex has been shown to

shuttle between the cytoplasm and the nucleus, driven by the NLS of p50 and the nuclear

export sequence of both IκBα and p65 [1]. Nevertheless, it is the degradation of IκB

proteins that alters this dynamic balance of nuclear/cytoplasmic localization, favoring

nuclear entry of NF-κB dimers. This degradation event is induced upon phosphorylation

of specific serine residues on the IκB proteins by the activated IκB kinase complex

(IKK), namely Ser32 and Ser36 of IκBα and Ser19 and Ser23 of IκBβ. Such

phosphorylation results in K48-linked polyubiquitination by the SCF βTcCP E3 ubiquitin ligase complex on Lys21 and Lys22 of IκBα, an ATP-dependent event that rapidly targets these proteins for lysosomal degradation [7]. Bcl-3 and IκBζ are inducibly

expressed atypical IκB proteins that regulate NF-κB function by a distinct mechanism.

IκBζ (also known as MAIL) localizes to the nucleus indicating that it regulates nuclear

NF-κB activity rather than its translocation from the cytoplasm [8]. Its expression is barely detectable in resting cells and is strongly induced upon NF-κB activation where it associates primarily with p50 homodimers to positively and negatively regulate NF-κB target genes. Bcl-3 on the other hand is a unique IκB family member since it contains a transactivation domain. It associates with p50 homodimers and stabilizes this transcriptionally inert complex hence negatively regulating NF-κB gene expression [9].

Conversely, Bcl-3 binds p52 homodimers to transactivate the expression of genes such as

cyclin D1 and p53 [10, 11].

The IKK Complex

The IKK complex, sometimes referred to as the IKK signalosome contains two

kinases IKKα/IKK1/CHUK and IKKβ/IKK2, as well as several copies of a regulatory

subunit IKKγ/NEMO. IKKα and IKKβ are homologous particularly in their catalytic

region (65% homology) and contain helix-loop-helix domains (HLH), while IKKγ is

distinct, smaller and characterized by coiled-coil, leucine zipper and Zn finger-like

domains [12]. IKKα and IKKβ dimerize through their leucine zipper domain, and

although these kinases can homodimerize, heterodimers are highly favored and are more

catalytically efficient [13]. They both bind IKKγ through their C-terminal NEMO-

binding domain (NBD), though with different affinities [14]. Activation of the IKK

complex is mediated by IKKγ oligomerization and interaction with upstream signaling

adapters and the subsequent phosphorylation of T-loop serines of at least one of the IKK

subunits, through the action of an upstream kinase or by transautophosphorylation [15].

Interestingly, K63-linked ubiquitination, as well as SUMOylation and phosphorylation of

IKKγ have been described in response to various upstream NF-κB activators [16].

Activation of IKK is a transient event that is terminated by deubiquitination of IKKγ

through the action of the NF-κB target gene A20, and the Cylindromatosis (CYLD)

deubiquitinases [17]. This signaling shutdown is also mediated through

dephosphorylation of IKK T-loop serines by the protein phosphatase 2A (PP2A).

Prototypical IKK substrates include IκBα and IκBβ, with IKKβ being more

efficient than IKKα in phosphorylating the IκB family members [17]. Another substrate

for IKKβ is Ser536 and 468 of p65, resulting in the enhanced transactivation potential of

this NF-κB subunit [18]. IKKα on the other hand phosphorylates p100 on several serine

residues leading to its limited processing into the p52 subunit as well as docking of the

NF-κB inducing kinase (NIK) [19]. Interstingly, and similarly to IKKβ, IKKα can also

phosphorylate p65 on Ser 536 in response to IL-1 signaling, enhancing its transactivation

potential {Buss, 2004 #920}. Other IKKα targets include transcriptional cofactors such

as CBP and SMRT [20, 21], chromatin such as histone H3 [22], as well as mitotic

regulators such as Aurora A [23].

Upstream Activators of NF-κB

NF-κB is activated by a variety of upstream signals including bacterial products,

inflammatory cytokines, oxidative stress, and mitogens. Such signals are then channeled

through intracellular adapter proteins that allow for specific receptor-induced signaling events, starting with IKK activation and culminating with NF-κB dimer nuclear

translocation. TNF receptor associated factors (TRAF) are a family of such adapter

proteins critical for NF-κB signaling pathways. They are characterized by a conserved

TRAF domain important for homo and heterodimerization as well as interaction with

surface receptors [24]. This family of 6 members may function as E3 ubiquitin ligases.

They get recruited to the cytoplamic portion of receptors upon ligand-induced receptor

oligomerization through interactions with TRADD (in the case of TRAF2 and TNFα

signaling), or MyD88 and the IRAK kinase (for TRAF6 in response to IL1 [3]) and result

in the assembly of multiprotein signaling complexes.

The receptor interacting proteins (RIPs) are another family of adapter proteins

that can recruit the IKK complex through binding to IKKγ [25]. These are

serine/threonine kinases that, in the case of RIP1 associate with TRADD via its Death

Domain (DD), and TRAFs1, 2 and 3 via its intermediate domain [26]. Their kinase

activity is dispensable in some signaling pathways and needed for others, and the various

RIP members exert non-redundant functions. Nevertheless, they act as scaffolds and

adapters for IKK activation and RIP 1 is essential for NF-κB activation via TNFα, TLR3

and 4 [25].

1.2 Classical and Alternative NF-κB Signaling

Targeted disruption of the different IKKs has shown that IKKβ and IKKγ are

essential for p65/p50 activation via IκBα phosphorylation [27], while IKKα is largely

dispensable. Such a signaling pathway, mediated by IKKβ and IKKγ activation leading

to IκB degradation and NF-κB activation was thus named classical or canonical signaling

(Figure 1.2). It occurs in response to upstream signals such as proinflammatory

cytokines and viruses. Conversely, pathways signaling through IKKα are known as

alternative or non-canonical and are based on the fact that p52 is stored in cells in its

p100 precursor form. Whereas the processing of p105 to p50 is constitutive, processing of p100 into its p52 subunit is a tightly regulated process induced by signals involved in

B-cell maturation and lymphoid organogenesis namely B-cell activating factor (BAFF),

CD40 ligand, and lymphotoxin β [28, 29]. Alternative NF-κB signaling does not require

IKKβ, IKKγ or RIPs, it rather proceeds through the activation of the NF-κB inducing

kinase NIK and is negatively regulated by TRAF3 [30, 31]. NIK activation leads to p100

phosphorylation on two sites Ser 866 and 870, an event required for the recruitment and

docking of IKKα on p100 [19]. Once recruited, IKKα can now phosphorylate p100 on

both C and N terminal serines, resulting in the recruitment of SCF βTcCP E3 ligase,

polyubiquitination and subsequent processing into p52 [32]. Such processing can occur

at the posttranslational level, constitutively (in the case of some lymphomas), as well as

by a cotranslational mechanism [31].

p100 is mainly cytosolic in unstimulated cell bound to the RelB subunit of NF-

κB, and its processing releases the RelB/p52, a complex poorly sequestered by other IκB

members. More recent data implicates the RelB/p52 dimer in the regulation of NF-κB

target genes distinct from classically regulated promoters and reports variations in the

conscensus NF-κB binding site to favor one complex versus the other [33, 34]. 7

1.3 NF-κB and cellular differentiation

Osteoclastogenesis

Aside from its more commonly accepted roles as regulator of innate immunity

and cell survival, NF-κB is also prominent in orchestrating cellular differentiation. One such κB-regulated differentiation system is that of osteoclast formation or osteoclastogenesis. Osteoclasts are multinucleated cells responsible for bone resorption,

and are histologically characterized by a ruffled border necessary for their attachment to

the bone surface. They differentiate from hemopoietic stem cells into the common

macrophage/osteoclast precursor cells in response to M-CSF [35]. The subsequent

differentiation step into multinucleated osteoclasts is characterized by cell-cell fusion and is controlled by various signals such as RANKL and NF-κB. The last step of maturation

results in bone-resorbing active osteoclasts, a phenomenon dependent on c-Src and

carbonic anhydrase II among others [35]. Generation of p50/p52 double knockout had early on established the importance of NF-κB signaling in osteoclastogenesis. Although

mice deficient in p50 or p52 alone have no bone disorders, lack of both subunits resulted

in severe osteopetrosis [36, 37]. In addition, these two subunits were found to be

dispensable for the determination of the RANK-expressing osteoclasts precursor lineage

[38], yet they are essential for their terminal differentiation and bone resorption activities

[39]. Mechanistically, such an effect was mediated by p50 and p52 through activating c-

Fos and NFATc1 expression in response to RANKL and TNFα [40]. Further attempts to

implicate the classical or alternative NF-κB signaling pathways in osteoclastogenesis are

8 still unclear. On the one hand, NIK-/- mice do not exhibit osteopetrosis despite increased p100 levels and failure to differentiate in vitro [41]. In addition, IKKα was found to be required in vitro but not in vivo for osteoclastogenesis, while IKKβ turned out to be a critical mediator of osteoclast survival and bone resorption activities, arguing against a role for the alternative pathway in this differentiation system [42, 43]. More recent evidence however points to RelB being the transcriptionally active NF-κB subunit signaling downstream of NIK that is required for osteoclast differentiation and inflammatory osteolysis in vivo [44], although this same group also provided evidence in a different study for p65 protecting against JNK-mediated cell death during differentiation [45]. Thus the precise role of the two NF-κB signaling pathways and the function of the various subunits in osteoclastogenesis is still unclear and requires further investigation.

Hematopoiesis

Hematopoiesis involves several proliferation and differentiation steps whereby a pluripotent hematopoietic stem cell differentiates into common myeloid and common lymphoid progenitor cells [46]. Myeloid progenitors can now give rise to osteoclasts, macrophages, erythrocytes, granulocytes and dendritic cells, while lymphoid progenitors further differentiate into B and T cells. These processes are regulated by several signaling pathways such as the JAK/STAT, MAPK and PI(3) kinase pathways [47]. NF-

κB is an essential regulator of these various stages of innate and adaptive immunity. It affects proliferation, apoptosis and differentiation of B-cells in response to extracellular

signals [48]. For example, B-cells lacking p65, c-Rel, p50, or the double knockouts p65/p50 and c-Rel/p50 are unresponsive to LPS or mitogenic stimuli, and show decreased survival in response to signals such as α-IgM [48-50].

Hematopoietic stem cells that lack both p65 and p50, or both p65 and c-Rel, or

IKKβ do not generate any lymphocytes [29, 51, 52]. Such an effect might be due to the

protective role of NF-κB against excessive TNF-α signaling [53]. Deletion of p65 or c-

Rel does not affect myelopoiesis presumably due to compensatory effect from the other

NF-κB members. However, deletion of both p65 and c-Rel affects common myeloid

progenitors resulting in impaired erythropoiesis, macrophage apoptosis, aberrant expansion of granulocytes and reduced colony-forming units progenitors [51]. Further studies also showed that p65, RelB, and p50 are required for the development of dendritic cells, while c-Rel instead affects their maturation and survival [54].

1.4 Skeletal Muscle Differentiation

Description and Regulation

Skeletal muscles in the body derive from a subdivision of the paraxial mesoderm

called the somites. Muscle progenitors within the maturing somite then become confined

to the dorsolateral region called the dermomyotome, migrate to the limb buds and

differentiate to form muscle fibers [55]. Migrating somites are characterized by the expression of Pax3 and Pax7, two members of the paired homeodomain transcription factors that need to be downregulated for the myogenic program to initiate. A small

fraction of these cells marked by the expression of Pax7 generates the satellite cells that

10 then reside between the basal lamina and the sarcolemma of myofibers [56]. Skeletal muscle differentiation is controlled by members of the basic-helix-loop-helix transcription factors, including MyoD, myogenin, Mrf4, and Myf5. Such factors are necessary and sufficient for the determination of the myogenic lineage, with Myf5 and

MyoD expression preceding that of Myogenin [57]. MyoD and Myogenin are expressed during skeletal muscle differentiation, while Mrf4 is present in terminally differentiated cells [58]. MyoD forms heterodimers with the E-protein subfamily and binds to a consensus sequence termed E-box present in the regulatory regions of many skeletal muscle genes including its own and that of the Mef2 transcriptional regulator [59, 60]. It also initiates chromatin remodeling through recruitment of HATs and the SWI/SNF complex [61, 62]. It is noteworthy that although MyoD and Myf5 are expressed in undifferentiated muscle precursors or myoblasts, they are transcriptionally inactive, bound to the Id protein [63]. MyoD activity is also kept silent on the enhancer of skeletal muscle genes in myoblasts, sequestered in a complex containing the Sir2 histone deacetylase as well as the pCAF acetyltransferase [64, 65]. Another inhibitory complex found on the promoter of myogenic genes comprises the polycomb repressor YY1, the

Ezh2 methyltransferase and HDAC1 [66, 67]. During myogenic differentiation, this complex gets replaced by the transcriptionally active MyoD/SRF complex allowing the expression of these contractile proteins.

Myogenesis can be positively or negatively regulated by a variety of factors and signaling pathways. p38MAPK is one such regulator that activates differentiation by recruiting the SWI/Snf complex to muscle promoters, phosphorylating the E47 E-box

protein to promote its association with MyoD and/or phosphorylating MEF2A, C and D

factors [68, 69]. It was also shown to phosphorylate MRF4 to inhibit its activity and to

antagonize the JNK proliferation-promoting pathway [70, 71]. Another pro-myogenic

signaling pathway is the PI(3)K/Akt pathway that gets activated in response to

insulin/IGF-1 [72]. It signals through its effectors mTOR and p70S6K to stimulate

protein synthesis and promote myotube hypertrophy [73]. Negative regulators of

myogenesis include fibroblast growth factors FGFs, mainly FGF6 [74, 75], TGF-β [76]

and the oncogenes Ha-ras, E1a and c-fos [77-79]. In addition, myostatin (Mstn) is

another repressor of skeletal myogenesis that acts by suppressing the activity of MyoD, and its absence is associated with the double muscled phenotype in cattle, mice and even

humans [72, 80, 81].

Myostatin in Myogenesis

Myostatin (Mstn) was originally identified as a member of the TGF-β family that

results in a hypermuscular phenotype in cattle, mice or even humans when inactivated

[81, 82]. It was thus recognized as a negative regulator of myogenesis. Both developing

and adult skeletal muscles express Mstn [83], and studies on cultured myoblast cells

showed that Mstn localized predominantly to myotube nuclei [84]. Transgenic animals

expressing a muscle-specific Mstn inhibitor result in muscle mass increase resulting from

both fiber hypertrophy and hyperplasia [85]. More recently, postnatal knockout of Mstn

showed increased muscle mass and improved muscle strength [86]. In addition, short-

term inhibition of Mstn in aged mice enhanced muscle regeneration and activated satellite

cell activation in a sarcopenia model [87].

Mechanistically, Mstn was found to inhibit myoblast proliferation and DNA

synthesis, to arrest muscle cells in the G1 phase of the cell cycle and to inhibit the

recruitment of p300 to the cyclin D1 promoter resulting in its silencing [88-91]. Mstn

also inhibits myogenesis by suppressing the synthesis of the transcription factors MyoD,

Myf5 and myogenin in differentiated myotubes [91-93]. Furthermore, myogenesis is blocked by Mstn through its ability to phosphorylate MyoD, causing a loss in

transcription factor DNA binding activity [93]. More recently, Mstn was implicated in

the control of satellite cell self-renewal through regulation of Pax7 [94]. It is thus now

becoming clear that Mstn can act as a potent negative regulator of skeletal muscle mass during development, and several attempts at inhibiting its expression or function are currently being investigated for maintaining muscle mass during diseases.

1.5 NF-κB in Myogenesis and Muscle Regeneration

Early studies using the C2C12 myoblasts line had revealed the p65 and p50

subunits of NF-κB to be abundantly expressed [95]. Further studies then reported c-Rel to be present at low levels in skeletal muscles [96]. Similarly RelB, p100/p52 and Bcl-3 were also described in adult muscles, suggesting a role for all NF-κB members in muscle formation and/or function [97]. It is noteworthy that neither p50-/- nor c-Rel-/- skeletal muscles showed differences in their fiber morphology suggesting that these subunits are not required during muscle development, results that were also confirmed in MyoD-

13 converted murine embryonic fibroblasts (MEF) [96, 98]. Nevertheless, and given the role of NF-κB in various cellular differentiation models as well as diseases, many laboratories have focused on this signaling pathway in myogenesis. Findings are perplexing though, with both pro-myogenic and anti-myogenic roles of NF-κB being described.

NF-κB as a Myogenic Activator

NF-κB activation during C2C12 myogenesis has been described in response to p38 MAPK signaling or activation [99, 100]. Using electrophoretic mobility gel shift assays (EMSA), Baeza-Raja et al. showed increased NF-κB DNA binding during differentiation, a process dependent on p38 activation. The authors further went on to provide evidence that both p38 and NF-κB activation are required for IL-6 production during myogenesis. Likewise, a recent study by De Alvaro et al. reported enhanced NF-

κB DNA binding in response to overexpression of mutant Ras deficient in Raf activation, a process that was also preceded by p38 MAPK signaling. Further studies subsequently reported an activation of NF-κB by EMSA analysis during C2C12 differentiation in response to insulin/IGF-II stimulation [101-105]. Mechanistically, this activation of NF-

κB was mediated by PI(3)K signaling in response to IGF-II, leading to decreased IκBα levels [105] and was found to restore differentiation of Ras-transformed cells [103]. In addition, IGF-II mediated NF-κB induction was accompanied by NIK and IKK activation, and was found to be specifically dependent on the kinase potential of the

IKKα subunit of the IKK complex, thus implicating the alternative pathway in

myogenesis [102].

Inhibitors of myogenesis were found to block differentiation at least partly by abrogating NF-κB activity. Specifically, pyrrolidine dithiocarbamate (PDTC) and the

proteosomal inhibitor Lactacystin block NF-κB activation in L6 rat myoblasts, thus

hindering cellular fusion and expression of muscle-specific proteins [106, 107].

Additionally, three-dimensional (3D)-clinorotation, a simulated-model of microgravity

was also shown to inhibit differentiation of these same cells through preventing IκB

ubiquitination and subsequent NF-κB activation [108]. More recently, we have used

MyoD-converted MEF deleted for RelB, p52 or even their upstream regulator IKKα to

examine the involvement of NF-κB in myogenesis [98]. Surprisingly, lack of these

subunits abrogated differentiation, suggesting that these NF-κB subunits are actually

required for myogenesis. However further analysis in primary non-transformed cell lines

failed to prove that such NF-κB alternative complex is required for expression of

myofibrillar genes, highlighting the limitations of the use of immortalized cell lines as a

genetic model. Nevertheless, our studies show the requirement for the NF-κB alternative

dimer in the formation of metabolically active myotubes capable of energy production,

thus underscoring its role as a myogenic activator.

NF-κB as a Myogenic Inhibitor

While the above-mentioned studies provided evidence for NF-κB as a pro- myogenic factor that gets activated during differentiation, a compelling number of other studies rather showed that this factor is a negative regulator of myogenesis. Using similar EMSA analysis in the C2C12 myoblast cell line, NF-κB DNA binding activity was found to decrease following onset of differentiation [95, 109-112]. Such a decrease occurred within 12h in differentiation medium and correlated with downregulated levels of the NF-κB target gene IκBα thus indicating decreased transcriptional activity [95,

109]. Additionally, inhibition of NF-κB signaling through expression of the dominant negative IκBα-superrepressor (SR) mutant accelerates myogenesis, increasing myogenin expression as well as myotube formation [95]. Conversely, activators of NF-κB such as

TNF-α, the TNF family member TWEAK, IL1-β, or the RIP homologue RIP2 strongly inhibit myogenesis [113-117]. Furthermore, glutathione depletion and cyclic mechanical strain impair myogenic differentiation through sustained activation of NF-κB [118, 119].

We have now used MEFs as well as primary myoblasts and histological muscle fiber analysis from mice lacking the classical p65 subunit to confirm the anti-myogenic role of this NF-κB family member [98]. Such analysis revealed that classical NF-κB signaling is a potent inhibitor of differentiation, and its absence results in enhanced myogenesis and myotube formation both in vitro and in vivo. Mechanistically, NF-κB has been found to regulate cyclin D1 expression, keeping myoblasts proliferating, and thus inhibiting their differentiation [95]. Binding of NF-κB on cyclin D1 decreases during myogenesis, therefore allowing the transition to the differentiation stage. Furthermore, NF-κB can

suppress synthesis of MyoD through binding to a destabilization element in the MyoD

transcript, particularly in response to TNF-α and TWEAK signaling [113, 116, 120].

More recently, NF-κB was shown to inhibit myogenesis through binding to the

YinYang1 (YY1) transcriptional repressor, resulting in transcriptional silencing of

myofibrillar genes [67]. Interestingly, NF-κB-mediated YY1 expression decreases

during differentiation, allowing derepression of myogenic genes such as Troponin I2 and

providing another level of myogenic regulation by NF-κB [67].

Model for NF-κB Regulation of Myogenic Differentiation

It is now clear that NF-κB regulation of myogenesis is an intricate process,

rendered even more complex by the various roles that the different NF-κB subunits play.

Nevertheless, since deletion of p65 the classical NF-κB subunit and its upstream

regulator IKKβ results in the same phenotype of enhanced myogenesis in MyoD-

converted fibroblasts, primary myoblasts as well as various skeletal muscles, one can

strongly conclude that classical NF-κB signaling is a negative regulator of myogenic

differentiation [98]. Such inhibition can be achieved through transcriptional regulation of

the cell cycle regulator cyclin D1, as well as the transcriptional repressor YY1. Other

mechanisms include MyoD stabilization as well as some yet unidentified mechanisms.

On the other hand, and although data from IKKα-/-, as well as p52 and RelB-/-

immortalized MEFs remains puzzling, preliminary data from primary IKKα null MEFs

as well as myoblasts suggest that this kinase is not required for expression of myofibrillar

genes, although litter variability has been observed [98, and unpublished observations].

Alternative NF-κB signaling mediated by IKKα, RelB and p52 is however necessary for

mitochondrial biogenesis and subsequent ATP production, two requirements for a

metabolically active contractile myotube [98]. Consequently, NF-κB signaling appears

to be a bipartite process that can through its classical subunits negatively regulate

myogenesis to prevent premature differentiation in myoblasts. Onset of differentiation is accompanied by a shutdown of classical signaling and the activation of the alternative dimer needed for mitochondrial biogenesis and energy production. Similarity in the

DNA binding sites of the classical and alternative NF-κB dimers might have thus been to

blame in the seemingly contradicting literature about the role of this signaling pathway in

myogenesis. Use of more specific p65 vs RelB EMSA probes in the future should allow

better dissection of these two signaling pathways in biological processes.

NF-κB Regulation of Muscle Regeneration

Adult skeletal muscles are stable tissues that undergo little turnover. However, in

response to severe injury such as exercise, muscle damage or degenerative muscle

diseases, they can undergo complete regeneration thanks to their residents muscle

precursor, the satellite cells. This process starts with a phase of degeneration

characterized by muscle fiber necrosis and inflammatory cells infiltration and is followed

by the activation of muscle repair [121, 122]. Satellite cells are mitotically and

metabolically quiescent in the adult, however they can be activated at the site of muscle

injury by microenvironment-secreted growth factors such as FGF, TGF-β and LIF [122,

123]. These cells then start proliferating and undergo rounds of cellular division and

differentiation and initiate expression of Myf5 and MyoD. At this stage they are called

adult myoblasts and undergo the various stages of myogenesis to fuse into injured fibers.

It is noteworthy that some of these activated cells also proliferate to restore the quiescent

satellite cell population [56, 124].

Being a regulator of myogenesis per se, NF-κB has also been found to modulate

muscle regeneration both in response to damage and in degenerative muscle diseases. In

a cardiotoxin injury model, lack of p65 from 4 week old mice was accompanied by

increased numbers of centrally located nuclei, a hallmark of muscle regeneration [67].

Similarly, mice lacking the upstream activator of NF-κB, namely IKKβ specifically in

skeletal muscles showed enhanced regeneration as revealed by increased sizes of repaired

fibers although no differences in the extent of injury or central nucleation were noted

[125]. Mechanistically, Mourkioti et al. observed increased numbers of centrally located

myonuclei per regenerated fiber in IKKβ-deleted muscles. Furthermore, these muscles

accumulated less fibrotic tissue and exhibited an earlier clearance of inflammatory

infiltrates, correlating with enhanced muscle regeneration. Similarly, and using another

model of muscle-specific IKKβ deletion in the Duchenne muscular dystrophy injury

model, Acharyya et al. reported central nucleation and embryonic myosin heavy chain

positive fibers confirming enhanced muscle regeneration [126]. The authors linked the

repair process to increased numbers of muscle progenitors, namely the CD34+/Sca-1- population and the Pax7-positive satellite cells. Taken together, these studies strongly support that disruption of classical NF-κB signaling enhances regenerative myogenesis

19 and conversely that this pathway negatively regulates adult muscle differentiation. It is noteworthy that the specific inactivation of IKKβ in skeletal muscles excludes the involvement of other cell types such as fibroblasts in the regeneration process, explaining discrepancies from other studies [127, 128]. These latter studies inhibited TNF-α signaling using a genetic model and observed compromised muscle regeneration, yet no links to NF-κB signaling were drawn. Conversely, muscle-specific overexpression of

IGF-1 accelerated muscle regeneration in response to local injury by decreasing fibrosis and the inflammatory response, possibly through inhibiting NF-κB [129]. It is noteworthy that use of the general NF-κB inhibitors curcumin and pyrrolidine dithiocarbamate (PDTC) in dystrophic mice or freeze-injured muscles increased expression of biochemical markers associated with muscle regeneration [130, 131].

However, both compounds are associated with non-specific effects and can activate various other signaling pathways; and so genetic knockout models remain the preferred tool to address the involvement of the NF-κB pathway in skeletal myogenesis.

1.6 Mitochondrial Biogenesis

Mitochondria: An Overview

Mitochondria are small membrane-bound organelles found in most cell types.

They range between 1 and 10um in size and their numbers per cell can vary between zero in red blood cells, to 50 in fibroblasts, all the way to a thousand in a cardiac myocyte.

They function in the regulation of cellular energy supplies (mainly ATP production), apoptosis, as well as mediating cellular signaling events [132]. Mitochondria have

20 several distinct compartments; the outer membrane allowing free diffusion of molecules less than 5000 KDa, the intermembrane space, the inner membrane home to the several members of the electron transport chain and several ion channels, and finally the matrix space [133]. The matrix is the space enclosed by the inner membrane and contains enzymes involved in fatty acid β-oxidation and citric acid cycle, mitochondrial tRNAs and ribosomes as well as 2-10 copies of the mitochondrial genome. Mitochondria possess their own genome in the form of a circular DNA that encodes 22 tRNAs and 13 polypeptides that are subunits of the oxidative phosphorylation machinery. This genome is maternally inherited and highly conserved, with a single promoter for all 13 genes transcribed as a polycistronic transcript [134]. The remaining 78 subunits of the respiratory chain are encoded by the nuclear genome and need to be imported into the mitochondria. Additionally, the protein machinery responsible for transcription, translation and replication of mtDNA is also nuclear-encoded. Such proteins encoded in the nucleus need to be translocated across both mitochondrial outer and inner membranes through the action of TOMs (translocase of the outer membrane) and TIMs (translocase of the inner membrane) [132]. Mitochondria cannot be made de novo, they rather form by a process of fission from existing organelles whereby new proteins are added to preexisting subcompartments before they separate into new mitochondria. The fission process is facilitated by the dynamin-related protein Drp1 that utilizes GTP hydrolysis to constrict and eventually pinch off the new organelle [135].

Diseases and disorders associated with mitchondria are often neurological, but can also include myopathies, encephalomyopathies or cardiopathies [136]. They can be

caused by mutations in mitochondrial DNA such as Leber’s hereditary optic neuropathy,

Pearson’s syndrome or myoclonic epilepsy with ragged red fibers and follow maternal

inheritance. Other mitochondrial diseases are attributed to defects in nuclear genes

leading to dysfunction of mitochondrial proteins. These include Friedreich's ataxia

characterized by progressive damage to the nervous system, and Wilson’s disease or hepatolenticular degeneration [136]. Other diseases are not directly linked to mitochondria, yet are associated with mitochondrial dysfunction, namely Alzheimer’s and Parkinson’s disease, epilepsy and stroke [137].

Mitochondrial Biogenesis

Mitochondrial biogenesis involves replicating mitochondrial DNA, along with transcribing the different mitochondrial proteins. It requires the tight coordination of

expression of both mitochondrial and nuclear genomes (Figure 1.3). Mitochondrial

transcription factor A (mtTFA) is required for mitochondrial DNA replication through

binding to a critical upstream region in the promoter and recruiting mitochondrial

polymerase. mtTFA itself is nuclear-encoded through the action of the nuclear

transcription factor 1 (NRF1), thus allowing nucleo-cytoplasmic interaction. Aside from

mtTFA, NRF1 controls many other nuclear genes involved in mitochondrial function and

biogenesis such as cytochrome c [138]. Other regulatory factors implicated in the

expression of respiratory genes include a functional homologue of NRF1 named NRF2,

the ubiquitous transcription factor SP1, and YY1 [138]. More importantly, mitochondrial

biogenesis is regulated by the peroxisome-proliferator-activated receptor coactivator-1

(PGC-1), initially discovered as an interacting partner for PPARγ in brown adipose tissue

[139]. The first member identified was named PGC1-α, while the other two were called

PGC1-β and PRC (PGC-related coactivator). These coactivators have strong transcriptional activity when they dock on a transcription factor, and although they do not have histone acetyltransferase properties, they can bind HATs such as p300 to form a transcriptionnally active complex. PGC-1 proteins can interact with a variety of transcription factors to activate various pathways in different tissues. For example, interaction of PGC-1α or β with nuclear respiratory factor 1 (NRF1) mediates mitochondrial biogenesis, while binding to PPARγ activates genes involved in fatty acid oxidation [140]. Although PGC-1α and β are highly similar in their N, C, and central domains, and are expressed in the same highly oxidative organs, differences in their upstream activators and downstream targets are observed. In brown fat, PGC1-α is highly induced upon cold exposure and turns on the adaptive thermogenic program in complex with PPARγ, while PGC1-β is induced during differentiation and seems to play a fundamental role in regulating brown adipocytes development also in collaboration with PPARγ [141]. PGC1-α and β are however essential for mitochondrial biogenesis

although some compensatory mechanisms between these two family members have been

observed. These coactivators serve as integrators of external stimuli activating

mitochondrial biogenesis in response to cold exposure, exercise, or energy demand [132].

Signaling pathways feeding into PGC-1 regulation of mitochondrial biogenesis include

calcineurin and calcium/calmodulin-dependent protein kinase (CaMK), as well as nitric

oxide/cGMP [132, 138].

Mitochondrial Biogenesis in Skeletal Muscle

Skeletal myogenesis is accompanied by changes in metabolic needs of growing myotubes and subsequent switching from glycolitic metabolism to oxidative phosphorylation as a source of ATP [142]. Remarkably, undifferentiated myoblasts possess only 5-20% of the mitochondrial content of myotubes, hence differentiation is accompanied by increased mitochondrial proliferation [143]. On the molecular level,

NRF2 and NRF1 mRNA levels increase by 2-4 folds, respectively [144]. PGC1-α was not detected in either myoblasts or myotubes, while PGC1-β levels increased by 60%, reaffirming the role of the latter in developmental mitochondrial biogenesis. Myogenic differentiation also resulted in increased citrate synthase and cytochrome oxidase levels and activity.

Contractile activity and exercise can also stimulate mitochondrial biogenesis through increased calcium flux and ATP turnover (Figure 1.3). Elevations in intracellular calcium result in activation of calcineurin and CaMK, leading to increased PGC1-α transcription mediated through MEF-2 and CREB-binding sites, respectively [145]. ATP depletion on the other hand can, through altering the AMP/ATP ratio activate the AMP- activated protein kinase (AMPK). AMPK can in turn induce increased fatty acid oxidation and mitochondrial biogenesis through direct phosphorylation and activation of

PGC1-α [146]. It is noteworthy that PGC1-α has also been found to serve as a target for calcineurin signaling, coactivating MEF2 proteins and driving the formation of slow- twitch muscle fibers [141]. Consistently, muscle specific overexpression of PGC1-α or

β results in increased mitochondrial numbers proteins and switching to oxidative fibers

[147-149]. On the other hand, PGC1-α or β knockout results in mitochondrial

dysfunction, lower respiration rates and increased numbers of glycolytic fibers [150,

151]. Hence, it seems that in skeletal muscle, both mitochondrial biogenesis and slow fiber type specification are regulated by PGC1-α and β, and these transcriptional

coactivators are currently being examined for therapeutic intervention for metabolic

disorders.

κ RELκ NF B anddomain I BNLS FamiliesTAD RelA/p65

cRel

L NFκB RelB Z p105/p50 * p100/p52 * IκBα IκBβ

IκB IκBε IκBγ Bcl3

IκBζ

Ankyrin repeats

Figure 1.1: NF-κB and IκB family members

ALTERNATIVE/NON-CLASSICAL CLASSICAL PATHWAY PATHWAY

BAFF IL-1 β CD40L Lymphotoxin TNFα

NIK

IKK Complex

α γ β

P P P P p50 p100 RelB IκBα p65

p52 RelB p65 p50

p65 p52 RelB p50

Figure 1.2: NF-κB signaling pathways.

Exercise

Calcium Nuclear-encoded mitochondrial genes CaMK α/β PGC1 PGC1α/β high AMP/ATP mtCO1

mtCO3 ? NRF2 mtCO2 p-AMPK PGC1α/β NRF1

Complex Differentiation Assembly mtCO1 Cues PGC1α/β mtTFA NRF1 mtCO2

mtCO3 mtDNA (13 genes) Nuclear-encoded NRF2 mitochondrial genes mtTFA

mtTFA

Nucleus Mitochondria

Figure 1.3: Mitochondrial biogenesis.

CHAPTER 2

MYOSTATIN AND NF-κB REGULATE SKELETAL MYOGENESIS THROUGH

DISTINCT SIGNALING PATHWAYS

2.1 Introduction

NF-κB is a ubiquitously expressed transcription factor involved in many cellular processes that regulate immune responses, cellular proliferation, differentiation, and cell survival [1, 152]. The mammalian NF-κB family consists of five members: NF-κB1

(p50), NF-κB2 (p52), Rel (c-Rel), RelA (p65) and RelB [6]. All of these proteins share a common REL homology domain that is responsible for dimerization, DNA binding, and nuclear localization [6]. However, these Rel proteins differ in their C-terminus such that only p65, RelB and c-Rel possess a transactivation domain. NF-κB proteins can exist as homo- or heterodimers, the most commonly found transcriptional activator being the p65/p50 heterodimer. In unstimulated mammalian cells, NF-κB is predominantly found in the cytoplasm in an inactive state, bound to its inhibitory protein (IκB). Various stimuli such as inflammatory cytokines, bacterial products, double stranded RNA, irradiation, reactive oxygen species or growth factors lead to the activation of NF-κB by inducing the degradation of IκB, thus allowing NF-κB to translocate to the nucleus where it binds to its cognate DNA sequence and stimulates gene expression [153, 154].

Aside from its more commonly accepted role as a regulator of innate immunity, accumulating evidence suggest that NF-κB is also involved in the differentiation and maintenance of skeletal muscle [72]. The dynamics of skeletal myogenesis is controlled by members of the myogenic basic helix-loop-helix (bHLH) transcription factors MyoD,

Myf5, myogenin and MRF4 [57, 155], as well as the myogenic enhancing factors

MEF2A, B, C, and D [156]. These myogenic transcription factors are responsible for the early myogenic commitment (MyoD, Myf5) and/or later downstream differentiation events involving cell cycle arrest, fusion, and expression of contractile genes (MyoD, myogenin, MEF2C, MRF4) [155, 157]. Mechanistically, NF-κB activity can inhibit myogenic differentiation by promoting muscle cell growth through transcriptional regulation of Cyclin D1 [95]. The cytokines TNFα or IL-1β have been shown to inhibit myogenic differentiation through the activation of NF-κB [113, 115]. In myotubes,

TNFα activation of NF-κB downregulates myosin heavy chain expression, a scenario associated with skeletal muscle wasting or cachexia [158, 159]. Furthermore, combinatorial treatment of similar myotube cultures with TNFα and IFNγ was shown to cause a loss in both myosin heavy chain and its upstream regulator MyoD also in an NF-

κB dependent manner [113, 160]. Taken together, these findings show that NF-κB is a pivotal player that mediates the inhibitory effect of inflammatory cytokines on both myogenic differentiation and the pathological muscle degeneration associated with cachexia.

Myostatin (Mstn, also known as growth and differentiation factor-8, GDF-8) is a member of the transforming growth factor-β (TGFβ) family of growth and differentiation

factors that has also been shown to be a potent regulator of skeletal myogenesis [161-

163]. Both developing and adult skeletal muscles express Mstn [164], and studies on cultured myoblast cells showed that Mstn localized predominantly to myotube nuclei

[84]. Furthermore, deletion of the Mstn gene in both mice and cattle showed dramatic

increases in muscle mass and body weight due to muscle fiber hyperplasia and

hypertrophy thus establishing its role as a negative regulator of muscle development

[164-167]. Strikingly, a similar phenotype has very recently been described in a child

exhibiting loss-of-function in both Mstn alleles, making this the first case where a human mutation mimicks the animal phenotype [168]. Mstn is also capable of inducing skeletal muscle degeneration [169]. Several groups have observed upregulation of Mstn levels in humans with conditions of muscle loss, resulting from HIV-infection, and disuse atrophy

[170-172]. Likewise, blockade of Mstn resulted in an increase in muscle mass, size and absolute muscle strength in both normal and dystrophic mice [173-175]. Consistent with these findings, systemically administered Mstn induced muscle and fat loss, in an analogous fashion to what is commonly observed in cachectic patients [176]. Further investigations into the mechanism of action of Mstn showed that it inhibits myoblast proliferation and DNA synthesis, and arrests muscle cells in the G1 phase of the cell cycle [89-91]. This arrest is accompanied by increases in p21 and p53 levels and accumulation of the hypophosphorylated form of retinoblastoma (Rb) protein [91, 92,

177]. In addition to its effects on muscle precursor cell proliferation, Mstn also inhibits myogenesis by suppressing the synthesis of the transcription factors MyoD, Myf5 and myogenin [91-93]. Myogenesis is also blocked by Mstn through its ability to

phosphorylate MyoD, causing a loss in transcription factor DNA binding activity [93].

Furthermore, Mstn signaling was found to stabilize Smad2/3 phosphorylation, induce

Smad7 expression and increase Smad3-MyoD association again leading to repressed

MyoD transcriptional activity [93, 178].

Given that NF-κB and Mstn have both been found to suppress skeletal muscle differentiation, we explored the possibility in this study that these signaling pathways crosstalked to regulate myogenesis, and to potentially also promote skeletal muscle wasting. Precedence exists for IKK/NF-κB and Mstn/Smad signaling crosstalk.

TGFβ signal has been found to both positively and negatively regulate NF-κB activity

[179-181], while NF-κB can function in a feedback loop to regulate TGFβ/Smad

signaling [182, 183]. Moreover, an NF-κB binding site has been reported in the myostatin

promoter, located –2982 and –2997 relative to the transcription initiation site [184].

Since little is known regarding the extracellular signals that regulate NF-κB in proliferating myoblasts, the following study was performed to examine the potential regulation of NF-κB by Mstn, and to better understand how these signaling pathways cooperate to inhibit skeletal muscle maturation.

2.2 Materials and Methods

Cell Culture and plasmids. C2C12 myoblasts were cultured and differentiated as

previously described [95]. Murine Mstn was purchased from R&D Research

(Minneapolis, MN), while murine TNFα was obtained from Roche Industries

(Indianapolis, IN). Reporter plasmids, 3x-κB-Luc, Tn-I-Luc, AchR-Luc, as well as

32 plasmids expressing MyoD, p65, and IκBαSR were used as previously described [95].

The Gal4-Luc reporter and the CMV-p65TA1 expression plasmid (containing the transactivation domain of human p65 from amino acids 521-551) were generously provided by A. Baldwin (University of North Carolina, Chapel Hill, NC).

EMSA and Transfections. Preparation of nuclear extracts from C2C12 myocytes and

EMSA analysis was performed as previously described [95]. For transfections, cells were plated in triplicate in 12 well dishes overnight. The next day cells were transfected with either reporter plasmids at 250 ng/well, or expression plasmids at 50 ng/well or at concentrations where indicated. Transfection efficiencies were normalized with CMV-

LacZ added at 250 ng/well. Results were reported as mean±standard deviation (SD).

RT-PCR. Total RNA was isolated with Trizol as recommended by the manufacturer

(Invitrogen, CA). Semi-quantitative RT-PCR was performed with 2 µg of RNA and the

Easy Access kit (Promega, WI), using the following Mstn primers that amplify a fragment size of 209 bp from Mstn cDNA: forward primer 5’CCT GAG ACT CAT CAA

ACC CAT G 3’; reverse primer 5’ CCT GGG AAG GTT ACA GCA AGA T 3’.

2.3 Results

To initiate this study we first asked whether Mstn was capable of activating NF-

κB in differentiating C2C12 cells. Electrophoretic mobility shift assays (EMSA) displayed the typical double banding pattern of NF-κB, which by supershift analysis in

33 myoblasts was previously identified to be the p50/p50 homodimer and p50/p65 heterodimer complexes [95]. Results showed that simply switching myoblasts from growth medium (GM) to differentiation medium (DM), in the absence of Mstn, was sufficient to induce a transient activation of NF-κB (Figure 2.1A, compare lanes 1 to 2 or lanes 1 to 6). However, addition of Mstn at increasing concentrations (0-100 ng/ml) for up to 1 h had modest effects on this activity. Likewise, addition of Mstn over a 24h timecourse caused only a minor induction of NF-κB as compared to myoblasts treated with DM alone (Figure 2.1B). As predicted, the addition of Mstn to differentiating

C2C12 myoblasts inhibited myotube conversion (Figure 2.1C) confirming that the lack of any significant induction of NF-κB was not related to Mstn inactivity. Together these results suggest that Mstn does not induce NF-κB nuclear translocation or DNA binding activity in myoblasts undergoing differentiation.

Given that Ras [185] and Akt/protein kinase B [186] have been shown to induce

NF-κB transcriptional activity without increasing NF-κB nuclear translocation or DNA binding [185, 186], we asked whether Mstn could potentially regulate NF-κB activity via a similar mechanism. C2C12 myoblasts were therefore transfected with an NF-κB responsive reporter plasmid and cells were subsequently induced to differentiate in the absence or presence of Mstn. In comparison to TNFα, which is known to stimulate NF-

κB transcriptional activity, no such activation was observed in the presence of Mstn in

C2C12 myoblasts (Figure 2.2A). To confirm this result, transfections were repeated with a p65 expression plasmid containing only the carboxyl transactivation domain fused to

DNA binding domain of GAL4. Again, while TNFα induced p65 transactivation, similar

activation was not observed upon Mstn treatment (Figure 2.2B).

Since NF-κB is constitutively active in proliferating myoblasts [95], we postulated that it is perhaps this portion of NF-κB that may be regulated by Mstn to maintain myoblasts in an undifferentiated state. To test this hypothesis, transfections were performed in C2C12 myoblasts with a myogenic responsive reporter in the presence

or absence of the transdominant inhibitor of NF-κB, IκBαSR (IκBα super-repressor),

which functions to inhibit basally active NF-κB. As expected, Mstn treatment decreased

the activity of the myogenic reporter, yet this regulation was unaltered in the absence of

NF-κB activity due to the expression of IκBαSR (Figure 2.3A). Similar results were

obtained when transfections were performed in 10T1/2 fibroblasts where myogenesis was

driven by a MyoD expression plasmid (Figure 2.3B). These results argue that basal

nuclear activity of NF-κB is not required for Mstn to inhibit muscle differentiation.

Similar to Mstn, constitutive activation of NF-κB in myotubes is associated with

muscle wasting [72]. TNF induced activation of NF-κB has also been shown to be higher

in myotubes compared to proliferating myoblasts, suggesting that mechanisms regulating

NF-κB activity in myotubes are distinct to those in myoblasts [160]. We therefore tested

whether Mstn was capable of activating NF-κB in differentiated myocytes. EMSA

results first showed that unlike the activation of NF-κB that occurred when myoblasts

were switched from GM to DM (Figure 2.1A), in pre-differentiated myotubes, the switch

from 3 day cultured DM to fresh DM (0 ng/ml Mstn) did not induce NF-κB activity

(Figure 2.4A). Secondly, treatment of myotubes with increasing concentrations of Mstn was seen to cause a modest, but reproducible transient increase in NF-κB activity. By 1h however, only myotubes treated with a high dose of Mstn (100 ng/ml) retained significant

NF-κB binding activity, but even at this dose, activity could not be sustained past this 1h time point (Figure 2.4B). In addition, and consistent with this transient increase, we were unable to detect any difference in NF-κB transcriptional activity in myotubes at 6 or 12 hours following Mstn treatment (data not shown).

Finally, because the Mstn promoter has been reported to contain an NF-κB consensus binding site [184], it remained possible that the basal activity of NF-κB detected in proliferating myoblasts functioned to regulate Mstn transcription, and this was possibly one mechanism of signaling crosstalk. To test this hypothesis we treated differentiating C2C12 myoblasts with TNFα to induce NF-κB activity and monitored

Mstn gene expression by semi-quantitative RT-PCR. Under these conditions however, little if any increase in Mstn expression was observed (Figure 2.5), nor did it appear to matter whether NF-κB activation occurred in myoblasts or pre-differentiated myotubes

(data not shown). Taken together, we conclude from these analyses that Mstn inhibition of myogenic differentiation is independent of NF-κB signaling.

2.4 Discussion

Mstn and NF-κB are major players in skeletal muscle homeostasis. Mstn, signaling through Smad 2/3 and Smad7 [178], and NF-κB, regulated by several cytokine signals inhibit myogenesis through similar mechanisms: they regulate components of the 36 cell cycle machinery, and inhibit the synthesis/function of MyoD, a master regulator of myogenesis. Both of these signaling pathways have also been implicated in skeletal muscle wasting. Given these similarities, the NF-κB binding site in the myostatin pomoter [184], and the literature showing crosstalks between TGF-β and NF-κB signaling [181, 182], we hypothesized that Mstn may be an upstream regulator of NF-κB in C2C12 skeletal myogenesis. Our results show that these two pathways, though targeting the same downstream effectors do not cooperate: Mstn does not induce NF-κB activity in differentiating myoblasts or differentiated myotubes, nor does it utilize the basal κB pool to mediate its effect. We also showed that Mstn expression is not regulated by NF-κB transcriptional activity. It is noteworthy that in a recent finding, differentially effects of Mstn on myogenesis was observed which depended on whether this factor was added to cells as a recombinant protein, as was done in this current study, or was overexpressed in cells by gene transfection [187]. Consistent with these findings, expression of MyoD by Mstn was differentially regulated in cases of overexpression versus exogenous treatment [92, 93]. Given these considerations, it remains possible that

NF-κB activity may be regulated by the overexpression of Mstn in myoblasts. However, we’re less likely to favor this scenario given the nature of non-specificity often associated with overexpression systems.

This study highlights the selectivity of signaling during myogenesis, and the importance of having distinct and separate signaling pathways that could be activated in response to different upstream signals. Mstn and NF-κB, like other signaling molecules

AP-1 [188] and delta Notch [189], do converge on MyoD, a master switch of

37 myogenesis, to regulate skeletal differentiation. Mstn was reported to downregulate

MyoD expression, and inhibit its activity through increased Smad3/MyoD binding [93].

NF-κB on the other hand has been shown to induce loss of MyoD in differentiating

C2C12 myocytes by a posttranscriptional mechanism [113]. However, although both pathways do converge on MyoD, their upstream regulators are distinct. Mstn expression is regulated by MyoD itself [190], glucocorticoids [184], or follistatin [191], while NF-

κB regulation in skeletal myogenesis is less well characterized. p38/MAPK and Akt signaling have been shown to activate NF-κB [104], however the significance of this activation in myogenesis remains to be completely defined. Taken together, these data show that Mstn and NF-κB signaling do not crosstalk to inhibit skeletal myogenesis, a finding that highlights the specificity of signaling pathways regulating this differentiation process.

0.5h 1hr A B DM DM DM + Mstn Mstn (ng/ml) GM 0 10 50 100 0 10 50 100 time (h) GM .5 1 4 12 24 .5 1 4 12 24

NF-κB

C DM DM + Mstn

free probe

Figure 2.1: Mstn causes a minor activation of NF-κB DNA binding activity in differentiating myoblasts.

A. Proliferating C2C12 cells were induced to differentiate (DM) in the absence or presence of different concentrations of Mstn (0, 10, 50, or 100 ng/ml) for 30 min or 1h. Nuclear extracts were prepared and EMSA was performed. B. Myoblasts were induced to differentiate (DM) in the absence or presence of 20 ng/ml of Mstn. Nuclear extracts were prepared at the indicated times and EMSA was performed. C. C2C12 myoblasts were cultured in differentiation media in the absence or presence of Mstn (50 ng/ml) for 72hrs.

A B 120000 control 50000 control TNF 100000 TNF 40000 Mstn Mstn 80000

30000 RLU

RLU 60000

20000 40000

10000 20000

0 0 6 h 24 h 6 h 24 h

Figure 2.2: Mstn does not regulate NF-κB transcriptional activity.

A. C2C12 myoblasts were transfected with 0.25 µg of a MHC-3xκB-Luc reporter plasmid and the next day cells were subsequently induced to differentiate for up to 24 hours in media containing either 5 ng/ml TNFα or 20 ng/ml Mstn. Cell lysates were collected 6 and 24 hours later and luciferase activity was measured. B. C2C12 myoblasts were co-transfected with 0.25 µg of a Gal4-Luc reporter plasmid with 0.01 µg of a CMV-Gal4-p65TA1 expression plasmid. Next day similar treatments were applied as in (A) and at indicated times, extracts were prepared and luciferase activity was measured.

A C2C12 B control 10T1/2 fibroblasts 30000 control Mstn Mstn 25000 4000000

20000 3000000 RLU 15000 RLU 2000000 10000 1000000 5000

0 0 pCMV IκBα-SR MyoD MyoD+ IκBα-SR

Figure 2.3: Endogenous activity of NF-κB is not required for Mstn inhibition of myogenesis.

A. C2C12 myoblasts were co-transfected with 0.5 µg of the troponin I-Luc reporter plasmid with or without 0.25 µg of the CMV-IκBαSR expression plasmid. 24 hours later, cells were induced to differentiate in the presence or absence of 20 ng/ml Mstn. Cell lysates were prepared and luciferase activity was determined. DNA was standardized by the addition of Bluescript plasmid (Stratagene Inc., La Jolla, CA). B. 10T1/2 fibroblasts were co-transfected with 0.5 µg troponin I- Luc, 0.05 µg CMV-MyoD in the presence or absence of 0.25 µg of the CMV-IκBαSR expression plasmid. Cells were subsequently treated with Mstn under differentiation conditions and after 24h luciferase activity determined.

A 0.5 h 1 h Mstn (ng/ml) DM 0 10 50 100 0 10 50 100

NF-κB

B 0 ng/ml 100 ng/ml

Time (h) DM 0.5 1 4 12 24 48 0.5 1 4 12 24 48

Figure 2.4: Mstn causes a transient increase in NF-κB DNA binding activity in myotubes.

A. C2C12 myoblasts were allowed to differentiate for 3 days, and then myotubes were treated with increasing doses of Mstn (0,10, 50, and 100 ng/ml) in DM for 0.5 and 1 h. Nuclear extracts were prepared and EMSA was performed. B. Myotubes were treated either in DM alone or DM containing 100 ng/ml Mstn for various time points in DM and nuclear extracts were prepared for EMSA analysis.

0 ng/ml 5 ng/ml TNFα Time (h) 0 24 1 4 6 12 24

Mstn

Figure 2.5: TNFα activation of NF-κB does not induce Mstn expression C2C12 myoblasts.

A. Myoblasts were treated with TNFα (5 ng/ml) to activate the NF-κB pathway under differentiation conditions. At indicated time points RNA was extracted and semi-quantitative RT-PCR was performed to detect for Mstn gene expression.

CHAPTER 3

IKK/NF-κB REGULATES SKELETAL MYOGENESIS VIA A SIGNALING

SWITCH

3.1 Introduction

NF-κB is ubiquitously expressed transcription factor and in mammals consists of

five family members, RelA/p65, c-Rel, RelB, p50 (the processed form of p105), and p52

(the processed form of p100) [192]. These subunits contain a DNA binding, protein

dimerization domain, and nuclear localization signal, but only RelA/p65 (from here

referred to as p65), c-Rel, and RelB possess transactivation domains. NF-κB forms homo

and heterodimers with the p50/p65 complex being the most common. In most cells NF-

κB is bound to IκB inhibitor proteins that masks its nuclear signal and sequesters it in the cytoplasm [193].

NF-κB is regulated by a variety of factors, such as inflammatory cytokines that

direct NF-κB by what is now referred to as the classical pathway [1]. This occurs through

stimulation of the IκB kinase (IKK) complex consisting of two catalytic subunits, IKKα

and IKKβ, and a regulatory subunit, IKKγ/NEMO/IKKAP1 [194]. Once activated, IKKβ

phosphorylates IκB proteins targeting them for ubiquitination and proteasomal

degradation. This releases p50/p65 or p50/c-Rel dimers to translocate to the nucleus and bind DNA where they induce gene expression. Mice null for IKKβ, IKKγ, and p65 are 44

embryonic lethal due to massive liver apoptosis, and cells derived from these embryos are unresponsive to classical NF-κB inducers [195-197], demonstrating a signaling link between p65, IKKβ, and IKKγ subunits.

In response to a second set of factors that include CD40L, BAFF, and lymphotoxin β, NF-κB is activated through an alternative pathway independent of IKKγ

[28]. Instead, activation proceeds through the NF-κB inducing kinase (NIK) that

phosphorylates and activates IKKα homodimers, and in turn phosphorylates p100 in

complex with RelB. This leads to ubiquitin-dependent processing of p100 to p52, and

translocation of p52/RelB to the nucleus [29, 198]. BAFF, NIK and p100/p52 knockout

mice have similar phenotypes [27], confirming that these molecules are also part of the

same linear “non-classical” signaling cascade. In addition, the classical and alternative

pathways are thought to regulate distinct genes in response to their various activators [33,

34].

Aside from its more commonly accepted role as a regulator of innate immunity

and cell survival, NF-κB is also prominent in regulating cellular differentiation. In

hematopoietic cells, c-Rel and RelB are essential for B cell lymphopoiesis and T cell

maturation [27, 199]. NF-κB is also required for osteoclastogenesis, and mice lacking

p50 and p52 display severe osteopetrosis [37]. Furthermore, IKKα is important for skin

differentiation as well as skeletal and craniofacial morphogenesis [200-202], a function

thought to be independent of its kinase activity.

Over the past years an increasing number of studies have also implicated NF-κB

in skeletal muscle differentiation, a process regulated by transcription factors MyoD,

Myf5, myogenin, MRF4/Myf6/Herculin, and MEF2A-D [57, 155, 156]. These factors regulate myoblasts to undergo growth arrest and fuse into multinucleated myotubes.

However, in contrast to hematopoiesis, the function of NF-κB in myogenesis is less defined and results have conflicted as to whether NF-κB promotes or inhibits this differentiation process. On the one hand, studies demonstrate that NF-κB DNA binding and transcriptional activities decrease during differentiation [95, 109] and that inhibition of NF-κB via expression of the IκBα-SR mutant accelerates myogenesis [95]. In addition, activators of NF-κB such as TNFα, IL-1β or the RIP homologue RIP2 act as potent inhibitors of differentiation [113, 115, 117], which together support the notion that

NF-κB functions as an inhibitor of myogenesis. NF-κB mediates this regulation through induction of cyclin D1 [95] or by suppressing MyoD synthesis through a destabilization element in the MyoD transcript [113, 120]. More recent data suggest that NF-κB can also inhibit myogenesis by stimulating expression of the Polycomb group protein, YY1 [67].

In contrast, similarly performed studies have reported that NF-κB activity increases during myogenesis in response to insulin-like growth factor (IGF) [103, 105].

IGF activation is mediated in part through the classical pathway causing IκBα degradation and p65 nuclear translocation, although the alternative pathway also appears to be involved since over expression of IKKα or NIK was seen to enhance myogenesis

[102]. In addition, expression of IκBα-SR in L6 rat myoblasts was also found to inhibit terminal differentiation markers, and recently it was also determined that p38 MAPK-

induced myogenesis functions through IL-6 synthesis in an NF-κB dependent manner

[100].

Taken together, these studies show that NF-κB function in skeletal muscle

differentiation remains at best enigmatic. Resolving this will not only provide insight into

the involvement of NF-κB during muscle development and repair, but it may also increase our understanding of its participation in a growing list of muscle wasting

disorders including cachexia [113, 125, 203], disuse atrophy [204], muscular dystrophies

[126, 205, 206], and inflammatory myopathies [207]. To this end, we used a genetic

approach to decipher the role of NF-κB/IKK subunits during myogenic differentiation.

Our results provide an explanation for the previously reported anti- and pro-myogenic

activities of NF-κB, by revealing that myogenesis involves both classical and alternative

NF-κB pathways. While constitutive activation of the classical pathway functions in

myoblasts to inhibit differentiation, NF-κB signaling switches to the alternative pathway

late in the myogenic program to promote mitochondrial biogenesis and myotube

homeostasis.

3.2 Results

Myogenic activity is enhanced in p65-/- MEFs expressing MyoD. To extend our

understanding of NF-κB in skeletal myogenesis, we used established murine embryonic

fibroblasts (MEFs) wild type or null for individual NF-κB subunits converted to skeletal

muscle by exogenous expression of MyoD [208]. We initiated this analysis with p65

since this subunit is constitutively active in myoblast nuclei [95]. Results showed that

myogenic activity derived from a troponin-I enhancer reporter plasmid (TnI-luc) was

significantly enhanced in p65-/- MEFs compared to wild type cells (Figure 3.1A). Similar

findings were obtained with plasmids containing the acetylcholine receptor promoter

(AchR-luc) or multimerized MRF binding sites (4RTK-luc) arguing that this effect was

not reporter-specific. To verify the specificity of p65 regulation, reporter assays were

repeated in early passaged primary fibroblasts prepared from E13.5 p65+/+, p65+/-, and

p65-/- embryos. Myogenic activity was again elevated in MEFs lacking p65, which

occurred in a gene dosage dependent manner (Figure 3.1B). This confirmed that p65

regulation of myogenesis was not a consequence of cell immortalization. As a control, myogenesis was also assessed in primary MEFs wild type or null for the retinoblastoma

(Rb) protein, a cell cycle checkpoint required for skeletal muscle differentiation [209]. As

predicted, reporter activity was significantly impaired in Rb-/- MEFs (Figure 3.1B), thus

supporting the relevance of our findings with p65. To further address p65 specificity,

myogenic assays were extended to MEFs lacking other NF-κB subunits. Results showed

that activity from cRel-/- or p50-/- MEFs was considerably lower than that for p65-/- cells

(Figure 3.1C). In addition, we used MEFs deficient in IκBα that contain constitutive

levels of nuclear p65 [196]. EMSA confirmed that NF-κB binding was higher in IκBα-/-

MEFs, which correlated with lower myogenic activity (Figure 3.1D). Together, these genetic data indicated that p65 functions as a negative regulator of MyoD-induced myogenesis.

MEFs null for p65 are accelerated in their myogenic program. To examine how the

absence of p65 exerts its effects on the myogenic program, MyoD was stably expressed

in p65+/+ and p65-/- MEFs using an MSCV-MyoD-IRES-GFP retrovirus. Following

selection, cells were sorted by flow cytometry for GFP to ensure equal levels of MyoD

(Figure 3.2A). Cells were then differentiated and myogenic markers were analyzed over a

4-day period. This analysis revealed that both induction and overall expression of

markers muscle creatine kinase, troponin, myosin heavy chain (MyHC), and tropomyosin

were greater in cells lacking p65 (Figure 3.2B). Myotube formation was also strikingly

higher in p65-/- cells (Figure 3.2C), which together supported reporter data above that p65

functions as an inhibitor of myogenesis.

Transcriptional activity of p65 derives from three transactivation domains (TA)

located in its carboxyl terminus [210]. To determine whether regulation of myogenesis

was dependent on p65 transcriptional activity, reporter assays were repeated in p65-/-

MEFs reconstituted with either p65 wild type (1-551 amino acids) or mutants truncated in

TA1 (1-521) or all three (1-313) TA domains. Compared to vector, addition of wild type p65 (1-551) or TA mutant (1-521) strongly repressed myogenesis, while expression of mutant (1-313) was effective in partially rescuing this regulation (Figure 3.2D). To verify these results, MyoD was stably expressed in p65-/- MEFs along with wild type or mutant

forms of p65 (Figure 3.2E). Consistent with reporter assays, myotubes were completely

absent in p65-/- MEFs expressing wild type p65, whereas some myotubes formed in cells

reconstituted with the (1-313) mutant (Figure 3.2F). This suggested that residues within

TA (521-551) are largely dispensable for repressing myogenesis, while residues (313-

521) play a more significant role in this regulation. However, because the (1-313) mutant only partially rescued myogenesis, it further suggested that residues within the Rel

domain contributed to p65 suppressive activity. Since phosphorylation of serine 276 is

required for p65 transactivation, [211, 212], we examined the involvement of this residue

in regulating myogenesis. Reconstitution of p65-/- cells with p65 containing a 276 serine

to alanine mutation was less effective in inhibiting myogenic activity, whereas mutations outside the Rel domain in residues 529 and 536 had no effect (Figure 3.2G). This is consistent with data above showing that deletion of TA (521-551) is not required for this regulation. In addition, generation of serine to alanine 276 in the p65 (1-313) mutant caused a full rescue of myogenic activity (Figure 3.2H), demonstrating that NF-κB

regulation of myogenesis is dependent on p65 transcriptional activity mediated from both

serine 276 and other residues lying within 313-521 domain.

Myogenesis is accelerated in p65 deficient myoblasts. To determine the physiological

relevance of our findings, myogenesis was further explored in p65-/- myoblasts. Although

mice lacking p65 are embryonically lethal [196], this phenotype can be rescued with

additional deletion of TNFα [213]. Thus, TNFα-/-;p65+/- mice were crossed and primary

myoblasts were prepared from 2-4 day old neonates (Figure 3.3A). Transfections with

troponin-I or MyHCIIB reporters showed that myogenic activity was substantially

elevated in p65-/- myoblasts, and like in primary MEFs, this regulation appeared to be

gene dosage-dependent (Figure 3.3B). In comparison, myogenic activity in p50-/- myoblasts was not significantly different from wild type cells. Furthermore, under

differentiation conditions p65-/- myoblasts formed 58% more myotubes (Figure 3.3, C

and D) and expressed higher levels of myofibrillar proteins (Figure 3.3E) compared to

p65+/+ cells. It is also noteworthy that modest, but reproducible expression of troponin

was detectable in p65-/- myoblasts even under growth conditions (GM; denoted by an

asterisk). Given that myofibrillar genes are silent in myoblasts, this suggested that p65

functions as a transcriptional repressor of troponin, consistent with our recent report that

NF-κB is capable of silencing troponin enhancers through the production of YY1 and

recruitment of Ezh2 and HDAC-1 [67]. Since TNFα has recently been shown to be

required for muscle regeneration [214], admittedly it was possible that the increase in

muscle differentiation in p65-/- myoblasts occurred secondary to the loss of this cytokine.

However, siRNA knockdown of p65 in primary myoblasts and C2C12 cells again led to

increases in myogenic activity and muscle markers (Figure 3.3, F and G), supporting results that negative regulation of myogenesis is specific to p65 and unlikely related to

the absence of TNFα.

Absence of p65 enhances myogenesis in vivo. Next, we explored muscles from TNFα-/-

;p65-/- mice in an attempt to correlate our in vitro findings with an in vivo phenotype. To

our surprise, p65 null muscles displayed a large number of fibers that were noticeably

smaller in size than their wild type counterpart (Figure 3.4A). In p65 deficient tibialis

anterior muscles from 4-week old male or female mice, average fiber diameter was

reduced by 39% as compared to wild type littermates (25.6µm for wild type vs 15.5µm

for null, n=5, Figure 3.4B). This phenotype was common to multiple hind limb muscles,

and was selective to p65 since no such differences were observed in p50-/- mice (Figure

3.4A). Slow MyHC staining from gastrocnemius muscles confirmed that the absence of

p65 did not affect fiber type either (Figure 3.11A). Although muscle atrophy is an

underlying cause of fiber reduction associated with induction of E3 ubiquitin ligases,

MuRF1 and atrogene-1/MAFbx, TNFα-/-;p65-/- muscles displayed no evidence of this

regulation (Figure 3.11B). However, given our findings with primary myoblasts lacking

p65, we considered the possibility that changes in p65-/- muscles might result from an

increase in overall myotube number. Indeed, fiber counts from entire cross-sections of

tibialis muscles revealed a 76% increase in p65-/- mice compared to control littermates

(Figure 3.4C). Similar results were found when counts were extended to gastrocnemius

and quadriceps, demonstrating that this regulation is not muscle type specific (n=5,

Figure 3.4D). Nor was the p65-/- phenotype related to a compromised immune function in adult mice since increases in fiber number were also observed in P7 and P9 neonates

(Figure 3.11C and 3.11D). These findings are consistent with cell culture data suggesting

that p65 absence in vivo leads to enhanced myogenesis, a phenotype highly reminiscent

of dystrophin deficient or acutely injured muscles depleted of p65 [126].

p65 regulates myogenesis through multiple mechanisms. Next, we sought to address

the mechanism by which p65 negatively regulates myogenesis. Previous use of the

IκBα-SR inhibitor revealed that p65 can inhibit C2C12 differentiation through the

suppression of MyoD synthesis [113]. Such analysis also revealed that NF-κB is capable

of inhibiting myogenesis through cyclin D1 [95] limiting myoblasts from exiting the cell

cycle, or through YY1 to silence myofibrillar promoters in myoblasts [67]. Consistent

with these findings, MyoD was elevated in p65-/- myoblasts, while both YY1 and cyclin

D1 levels declined (Figure 3.5A). Thus it is likely that p65 negatively regulates

myogenesis through multiple mechanisms.

To determine whether these mechanisms could function independently of each other, we examined the regulation of myogenesis by p65 in MyoD-/- myoblasts. Although

myotube formation is impaired in these cells [215], myogenic activity was nevertheless

retained under differentiation conditions (Figure 3.5B). However, addition of p65 or

TNFα strongly repressed this activity. Likewise, retroviral expression of p65 in MyoD-/- myoblasts caused a noticeable reduction of myogenic markers (Figure 3.5. C and D), demonstrating that p65 can inhibit myogenesis independently of MyoD. To substantiate this finding, reporter assays were repeated in p65+/+ and p65-/- MEFs where MyoD was

substituted with myogenin. Like MyoD, myogenin is capable of converting fibroblasts to a muscle lineage albeit with lower efficiency [216]. Indeed, myogenin-induced myogenic activity was less than that for MyoD, but these levels were nonetheless greater in p65-/-

MEFs compared to wild type cells (Figure 3.5E). This regulation also appeared specific to these MRF proteins since a p53 responsive reporter was not affected by the absence of p65. Together, these results support that p65 inhibits myogenesis through multiple mechanisms.

Myogenesis is regulated by a temporal switch in IKK signaling pathways. Having gained insight on the role of p65 in muscle differentiation, we now turned our attention to

its upstream regulator, the IKK complex. Recently, our group elucidated that chronic

activation of IKKβ in mdx muscles inhibits muscle differentiation [126]. Interestingly,

Mourkioti and colleagues have also reported that skeletal muscle deletion of IKKβ increased intermediate fiber numbers in 4-month old mice [125], a phenotype that closely matched that of younger p65-/- mice (Figure 3.4). Such results suggested that p65 and

IKKβ share overlapping functions in skeletal muscle differentiation. Indeed, analogously to p65, myogenic activity was increased in primary fibroblasts and myoblasts deleted for

IKKβ floxed (f/f) alleles using Cre recombinase (Figure 3.12A and 3.12B, Figure 3.6A).

Likewise, 4-week old mice lacking skeletal muscle IKKβ exhibited an increase in total

fiber number (Figure 3.6, B and C), reaffirming that IKKβ like p65 functions as a

negative regulator of myogenesis. Since IKKβ and p65 are components of classical NF-

κB signaling, myogenesis was also tested in IKKγ-/- MEFs to address whether this

pathway acts as a general inhibitor of differentiation. Consistent with this thinking,

myogenic activity was increased in IKKγ-/- MEFs, but decreased in MEFs lacking IKKα,

the latter of which is not considered part of the classical pathway [29] (Figure 3.6D).

To further explore the myogenic functions of IKK we measured its activity in

differentiating myoblasts. Results showed that IKK activation was relatively low in

undifferentiated cells, but became induced at 48 hr into the myogenic program (Figure

3.6E), a time when cell fusion and contractile myofibrillar expression is well underway.

This activity was specific to IKK since phosphorylation was undetectable when assays

were repeated with a mutated substrate, nor was the increase in activity a consequence of

altered protein expression since IKK subunits remained unchanged during myogenesis

(Figure 3.6E).

Next, we analyzed endogenous IKK substrates to ascertain which IKK complex became activated during late stage myogenesis. IKKβ activation, as part of the classical

pathway, results in phosphorylation of IκBα and p65 [192]. IKKα on the other hand

predominantly phosphorylates p100 leading to its proteolysis and conversion to p52.

Results revealed that levels of phosphorylated IκBα and p65 decreased during C2C12

differentiation, while total protein levels remained unchanged (Figure 3.6F). Consistent

with this decrease, nuclear p65 levels declined, and by ChIP, p65 binding activity on the

IκBα promoter was also lost (Figure 3.6F). In comparison, processing of p100 to p52 was

induced during myogenesis with similar kinetics to in C2C12 as well as primary

myoblast differentiation (Figure 3.6G and data not shown). Since IKKα activation results

in formation of RelB/p52, we also investigated the contribution of these NF-κB subunits

by repeating myogenic assays in RelB-/- and p52-/- MEFs. Consistent with findings in

IKKα-/- MEFs, myogenic activities were reduced in both p52-/- and RelB-/- fibroblasts

(Figure 3.6H). Together, these data suggest that skeletal myogenesis is characterized by a

temporal switch in NF-κB signaling pathways, whereby reduction of classical NF-κB is

followed by activation of the alternative pathway relatively late in the myogenic program.

IKKα functions as a regulator of myotube maintenance under metabolic stress. The

above data suggested that components of the alternative pathway might function to

promote myogenesis. However, stable expression of an HA tagged version of IKKα in 55

C2C12 myoblasts did not affect the induction of early or late myogenic markers,

myogenin and troponin, respectively (Figure 3.7A), nor was myogenic activity affected

when IKKα or a kinase dead version (K/M) of this kinase was over expressed in MyoD

expressing 10T1/2 fibroblasts (Figure 3.13A). In comparison, expression of classical

signaling components, IKKβ, IKKγ, or p65 led to clear reductions in myogenic activity in

these same cells. Furthermore, no differences in skeletal muscle gene expression were

detected by Affymetrix microarray between control and HA-IKKα expressing C2C12

myotubes (data not shown), and siRNA-mediated depletion of IKKα from differentiating

myoblasts also revealed little change in myogenic markers (Figure 3.7B). Therefore,

although results from IKKα-/-, RelB-/-, and p52-/- MEF suggested that the alternative pathway is pro-myogenic, overall, the data do not support that activation of this NF-κB

signaling pathway is necessary for myotube formation (see Discussion section).

However, under long-term differentiation conditions (6 days) without medium

replenishment we observed that myotubes expressing HA-IKKα were better maintained than control cells (Figure 3.7C). Specifically, IKKα expressing myotubes were 48% less

atrophic (26.0 ± 5.7µm in fiber diameter compared to 13.5 ± 3.5µm in control cells) and

overall exhibited a “healthier” morphological appearance. In addition, IKKα expressing

myotubes were also more resistant to low glucose, but not to heat shock, oxidative stress,

or DNA damage (Figure 3.7C, and data not shown), suggesting a selective resistance to

metabolic stress. This effect was dependent on the kinase activity of IKKα since myotube maintenance was lost upon expression of a kinase dead mutant (Figure 3.7D). Moreover,

siRNA deletion of alpha, but not the beta subunit, negated this protective effect upon glucose deprivation confirming the specificity of IKKα in this regulation (Figure 3.7E).

Furthermore, 6-day starved IKKα expressing myotubes displayed higher levels of myogenic markers, myogenin, troponin, MyHC and activated p38, whereas these markers were reduced upon IKKα knockdown (Figure 3.7F). Together, these data suggest that activation of IKKα and the alternative pathway during myogenesis functions to maintain myotubes in response to metabolic stress.

IKKα regulates mitochondrial biogenesis. Finally, we attempted to address the process by which IKKα controls myotube maintenance. Because IKKα regulation appeared selectively linked to starvation stress, we speculated that this kinase was involved in regulating the energy capacity of differentiating muscle. Energy production during myogenesis occurs through a switch from glycosidic to oxidative phosphorylation

resulting from an increase in mitochondrial content [142, 143]. Using semi-quantitative

PCR and the mitochondrial marker gene, cytochrome oxidase 1 (MTCO1), we readily detected an increase in mitochondrial DNA during C2C12 myogenesis (Figure 3.8A).

Examination of HA-IKKα differentiating myoblasts also revealed significantly higher

levels of MTCO1 DNA compared to vector (Vect) cells, whereas depletion of IKKα led

to reduction of MTCO1 (Figure 3.8B). This suggested the possible novel finding that

IKKα is a regulator of mitochondrial biogenesis. Consistent with this notion, myotubes

overexpressing IKKα contained higher levels of the mitochondrial dye, MitoTracker

(Figure 3.8C). In addition, mitochondrial fractions from these cells were also enriched in 57

cytochrome c, which were in turn reduced upon expression of IKKα siRNA (Figure

3.8D). Furthermore, total ATP was elevated by 72% in IKKα expressing myotubes

(p=0.001), while IKKα knockdown caused an 18% reduction in ATP levels (p=0.02,

Figure 3.8E). To address whether the increase in mitochondrial content by IKKα

reflected mitochondrial function, biochemical assays were performed for citrate synthase

and dehydrogenase enzymes. Results showed that enzyme activities were significantly

increased in IKKα expressing myotubes (Figure 3.8F). This function appeared selective

to the NF-κB alternative pathway since p52/RelB, but not p50/p65, increased MTCO1 and ATP (Figure 3.8G). Furthermore, ATP levels were also increased in MEF and HeLa

cells over expressing IKKα (data not shown), suggesting that IKKα regulation of

mitochondria is not specific to skeletal muscle.

To further investigate this regulation, ultrastructural analysis was performed in

IKKα over expression and knockdown conditions. Remarkably, HA-IKKα expressing

myotubes displayed elongated networks of mitochondria, a hallmark of extensive

proliferation (Figure 3.9A). Conversely, IKKα knockdown resulted in degenerating

organelles, evidenced by swelling and absence of cisternae in the mitochondrial matrix

(Figure 3.9B). In addition, genome-wide L2L analysis ([217] identified 126 selectively

enriched biological processes in upregulated genes (>1.5 fold) from IKKα myotubes.

From these, 48% were involved in mitochondrial and metabolic regulation (Figure 3.9C),

while significantly fewer IKKα regulated genes associated with transcription/translation

(12%) or even skeletal myogenic processes (6%). To examine whether regulation of

mitochondria is linked to myotube maintenance, we treated C2C12 myotubes with

mitochondrial inhibitors chloramphenicol and oligomycin under glucose deprivation.

This led to visibly less numbers of preserved myotubes (Figure 3.13B), a phenotype strikingly similar to IKKα depleted cells. Taken together, our results strongly support

that activation of NF-κB alternative signaling during myogenesis does not function to

promote myotube formation, but rather is important for regulating mitochondrial

biogenesis and myotube homeostasis.

3.3 Discussion

Recent studies have shown that chronic activation of NF-κB is detrimental to

muscle function. In skeletal muscles, NF-κB has been linked with disease states such as

cachexia and various forms of muscular dystrophies and inflammatory myopathies [126,

204-207]. Although such studies implicate NF-κB as a therapeutic target,

mechanistically, relatively little is known how this transcription factor mediates its

pathological effects. Elucidation of these mechanisms might be better achieved by

studying NF-κB function in basic models of skeletal myogenesis. However, even in

tissue culture systems reports have conflicted as to whether NF-κB acts as a repressor or

promoter of myogenesis. In this current study we describe what we believe to be a new

understanding for the role of NF-κB in skeletal muscle differentiation. Our findings

reveal that NF-κB is capable of functioning as both a repressor of differentiation and a

promoter of myotube maintenance depending on specific activities of IKK and NF-κB

subunits. 59

p65 and the classical NF-κB signaling pathway function as negative regulators of

myogenesis. Utilization of knockout MEFs demonstrated that myogenic activity was

enhanced in cells lacking p65 and comparisons with all five NF-κB subunits showed that

this activity was highest in p65-/- cells. Therefore, although myoblast nuclei have been

shown to contain constitutive activity for p50 and p65 [95], our current data argue that

suppression of myogenesis by NF-κB is mediated specifically through p65. This notion is

consistent with results in primary myoblasts where myogenic activity was also elevated in p65, but not p50 null cells. Together, these genetic data reaffirm that p65 activity in proliferating myoblasts functions as a negative regulator of myogenesis. This function of p65 is evident in muscle injury where lack of p65 enhances myogenesis in mdx and toxin treated mice [126]. Given that p65 deficiency also correlated with increases in overall fiber numbers in young and adult mice, it suggests that p65 is also relevant during postnatal muscle growth, as evidenced by the high levels of NF-κB activity in muscles from neonates [126]. Why p65 would function in this capacity at this stage of development is not yet known, and whether it functions in a similar manner during embryonic or fetal myogenesis remains to be investigated.

Our current results also demonstrate that regulation of myogenesis is dependent on p65 transcriptional activity. This notion is in line with our previous findings that NF-

κB inhibits myogenesis through the transcriptional activation of cyclin D1 [95].

Repression of myogenesis by p65 has also been seen in response to TNFα leading to the loss of MyoD [113], and more recently to the gain of YY1 resulting in silencing of

60 myofibrillar genes [67]. Thus, p65 requires its transactivation function to suppress muscle differentiation, and results from MyoD-/- myoblasts support that this can occur via multiple mechanisms.

Similar to p65, we also discovered that myogenic activity was enhanced in MEFs lacking classical components IKKβ and IKKγ. Like p65, IKKβ deletion in muscle led to increases in fiber number and to enhanced myogenesis in mdx mice [126]. Collectively, these data argue strongly that classical NF-κB signaling functions as a negative regulator of muscle differentiation in both physiological and disease processes.

IKKα signaling promotes myotube maintenance through mitochondrial biogenesis.

With respect to alternative NF-κB signaling, our results showed that activation of IKK during myogenesis is selective to IKKα as this activity tightly correlated with p100 processing. Such activation was preceded by a decline in classical pathway activity, depicted by decreases in IκBα and p65 phosphorylation, as well as p65 nuclear and

DNA-bound levels. In contrast to recent findings that nuclear localization of IKKα is required in skin differentiation [200, 201], or NF-κB dependent gene expression [20, 22,

218], we were unable to detect nuclear IKKα in myoblasts or myotubes (Bakkar and

Guttridge, unpublished observations). Although our current results do not rule out the possibility that IKKα might still phosphorylate an unknown target to modulate myogenic gene expression, we favor instead that IKKα function in skeletal muscle differentiation is

represented by the alternative pathway requiring the cytoplasmic form of IKKα to

activate p52/RelB complexes.

Evidence from IKKα-/-, p52-/- and RelB-/- MEFs indicated that alternative

activation of NF-κB is required for myogenic activity. These results appear consistent

with previous findings implicating IKKα as a positive regulator of myogenesis [102].

However, in contrast to these findings, we were unable to demonstrate by either forced

expression or RNAi depletion that IKKα is essential for induction of myogenic genes or

myotube formation. Although genetic evidence from p65 and IKKβ knockout MEFs

were consistent with how these classical signaling components were found to function in

muscle cells, we do not yet understand why this same consistency was not present

between IKKα-/- MEFs and C2C12 cells depleted of IKKα with siRNA. Possibly, the

fraction of IKKα that remains in cells after siRNA depletion is sufficient to mask a

phenotype that otherwise requires its complete absence, or perhaps the increase in

myogenic activity derived from established IKKα-/- MEFs might be an indirect

consequence of immortalization and continued subculturing. We suspect that additional myogenic reporter assays in primary IKKα-/- MEFs and myoblasts will be needed to

clarify this issue.

Nevertheless, our observations led to the novel discovery that IKKα acts as a

regulator of mitochondrial biogenesis. Although the mechanism remains unknown, we

predict that IKKα activation functions through p52/RelB to promote mitochondrial

biogenesis and meet the metabolic needs of a newly formed contractile myotube. Given

62 that the inhibitor compounds of mitochondria were also seen to decrease myotube maintenance suggests that IKKα regulation of mitochondria is necessary for myotube homeostasis in response to changing metabolic conditions.

A model for IKK/ NF-κB signaling in skeletal muscle differentiation. Collectively, our data support a model whereby IKK/NF-κB signaling both inhibits and promotes the differentiation state of muscle cells (Figure 3.10). This model helps unify the literature on the contradictory functions of NF-κB in myogenesis and predicts that during differentiation, a temporal switch occurs between NF-κB classical and alternative signaling pathways. In myoblasts, classical signaling is constitutively active and functions to maintain cells in an undifferentiated state. This function is regulated through the control of MyoD as well as other MyoD-independent mechanisms, involving cyclin

D1 and YY1. Once differentiation cues are initiated, classical signaling is turned down while the alternative pathway is induced late in the myogenic program. The activation of

IKKα leading to p52/RelB association in turn regulates myogenesis by mediating the production of mitochondria necessary to satisfy the metabolic needs of contractile muscle cells. Although cooperative functions of NF-κB signaling pathways are important for mammary and osteoclast tissue development [42, 219], skeletal muscle is to the best of our knowledge the first example of a differentiation system regulated through a functional switch of classical and alternative NF-κB signaling pathways.

3.4 Materials and Methods

Materials. Antibodies to p100/p52, IκBα (C21), IKKβ, IKKγ (FL419), myogenin (M-

225), p38, MyoD (M-318), p65 (N-terminal) were obtained from SantaCruz

Biotechnology (Santa Cruz, CA), MyHC IIB (MY-32), MyHC slow (NOQ7.5.4D), troponin T (JLT-12), sarcomeric tropomyosin (CH1), and α-sarcomeric actin (5C5) from

Sigma-Aldrich (St. Louis, MO). p65 antibody was obtained from Rockland

Immunochemicals, Inc (Gilbertsville, PA), hemagglutin (HA) from Covance (Princeton,

NJ), IKKα from Imgenex (San Diego, CA), phospho IκBα, p38 and p65 from Cell

Signaling (Beverly,MA), and cytochrome c from BD Pharmingen (San Jose CA). Bovine insulin, collagen type I, and gelatin came from Sigma (St-Louis, MO), while TNFα was purchased from Roche (Mannheim, Germany). Both collagenase P and dispase (grade II) were obtained from Boehringer Mannheim (Mannheim, Germany), basic human FGF from Promega (Madison, MI), and oligomycin from Alexis Biochemicals (San Diego,

CA). Mitotracker Green and secondary antibodies for immunofluorescence were obtained from Molecular Probes (Eugene, OR), while other materials for immunohistochemical analysis came from Vector Laboratories (Burlingame, CA).

Plasmids. Reporter and p65 expression plasmids were previously described [95, 220,

221] with the exception of the p65(1-313;S276A) mutant, generated by mutating serine

276 to alanine in the p65(1-313) plasmid. MSCV-MyoD was generated by subcloning the MyoD cDNA from a pBabepuroMyoD retroviral construct [113]. IKK plasmids were

64 designed by subcloning IKKα, IKKβ, and IKKγ into the pBSx-HSAvpA plasmid, whereby transgene expression is driven from the human skeletal actin promoter.

Transfections, luciferase assays and retrovirus infections. Sub-confluent C2C12 cells were transfected in low serum Opti-MEM using Lipofectamine (Invitrogen, Carlsbad,

CA) according to the manufacturer. For luciferase assays, cells were transiently transfected using Superfect (Qiagen, Valencia, CA for MEFs), or Lipofectamine

(Invitrogen, Carlsbad, CA) for primary myoblasts. All transfections were normalized to

CMV-βGAL expression. Cells were lysed in MPER solution (Pierce), and assays were performed as previously reported [95]. IKKα, IKKβ and p65 siRNAs were obtained from Dharmacon, Inc. (Lafayette, CO) and transfections were performed using

Lipofectamine 2000 (Invitrogen). Retrovirus production and infection were performed as previously described [95].

Mice and genotyping. Animals were housed in the animal facility at the Ohio State

University Heart and Lung Research Institute under sterile conditions maintaining constant temperature and humidity, and fed a standard diet. Treatment of mice was in accordance to the institutional guidelines for Animal Care and Use Committee. Mice null for p65 were generated as previously described [213], p50 mice were obtained from

Jackson Laboratories (Bar Harbor, Maine), and IKKβ flox mice [222] were crossed to

MCK-Cre mice to delete IKKβ in skeletal muscle. Mice genotypes were confirmed by

PCR analysis from prepared tail DNA.

Cell culture. C2C12 murine myoblasts and fibroblasts were cultured as previously described [113]. Primary myoblasts were prepared from 2-day old neonates adopted from the described procedures [223]. Briefly, limbs from pups were skinned and incubated with collagenase/dispase mixture at 370C for 1h. Then the cell suspension was

further homogenized by pipetting and pre-plated on uncoated cell culture plates in F10

media (Gibco/Invitrogen, Carlsbad, CA) to selectively enrich for myoblasts. Following 2

rounds of pre-plating, the cell suspension was plated on gelatin pre-coated plates, in the

presence of 20% FBS and 6ng/ml bFGF. Primary myoblasts were used at passage 3-5

post-isolation.

Immunoblotting, Northerns, ChIP, and kinase assays. Westerns, Northern, and kinase

analyses were performed as described [221]. For ChIP, assays were performed as

recommended by the manufacturer (Upstate Biotechnology, Inc).

Histology, electron microscopy, and immunofluorescence. For muscle analysis, tissues

were sectioned at 10 microns on a cryostat (Leica) and stained with hematoxylin and

eosin or processed for immunohistochemistry. The internal diameters (shortest diameter) from 1200 fibers in random fields throughout the muscle were recorded using Olympus

BX50 microscope and MetaVue 6.2r6 software (Universal Imaging Corporation, PA).

Fiber number was recorded in 25 randomly selected fields throughout the muscle and

averaged for comparisons. Muscles from 3-5 different animals per group were used.

Immunostaining procedures on cell lines and muscle sections were performed as

described [220, 224], and all images were captured with a Zeiss Axioskop 40 fluorescent

microscope using an AxioCam HRc camera and the AxioVision 3.1 software.

Ultrastructural analysis was performed on fixed cells, and sectioned using a Leica EM

UC6 microtome at 70nm. Sections were then stained and visualized using FEI Spirit

Tecnai Transmission electron microscope at 80kV and images were captured with an

AMT camera.

Mitochondrial Assays. Both CellTiter-Glo Luminescent Assay for ATP determination and MTS cell viability assays were obtained from Promega (Madison, MI) and performed

as per manufacturer’s recommendations. Citrate Synthase activity was determined by

using Ellman’s reagent with acetyl-CoA and oxaloacetate [225]. Procedures for primer

design and PCR of mitochondrial cytochrome oxidase subunit 1 (MTCO1), as well as

mitochondrial extraction for identification of cytochrome C were followed as described

[226, 227].

Statistical Analysis. All quantitative data are represented as mean ± SEM. Analysis was

performed between different groups using a two-tailed Student’s t test. Statistical

significance was set at a p value of <0.05.

Figure 3.1: Loss of p65 enhances myogenic activity in MEFs.

A. p65+/+ and p65-/- MEFs were co-transfected with CMV-MyoD and either of the following reporter constructs: TnI- luc, AchR-luc, or 4RTK-luc. Next day cells were switched to differentiation media (DM), and after 48 hr lysates were prepared and assayed for luciferase activity. B. p65+/+, p65+/-, p65-/- and pRb+/+, and pRb-/- primary MEFs were transfected with MyoD and TnI-luc. Cells were differentiated as in A., and luciferase assays were performed. C. p65-/-, cRel-/- and p50-/- MEFs were transfected with MyoD, and TnI-luc, differentiated and monitored for luciferase activity. D. Myogenic assays similar to those described in A-C were performed in IκBα+/- and IκBα-/- cells. Insert: EMSA analysis of IκBα+/- and IκBα-/- MEFs.

Figure 3.2: Loss of p65 accelerates the myogenic program in MEFs.

A. p65+/+ and p65-/- MEFs were infected with MSCV-MyoD, and following puromycin selection sorted for GFP to ensure equal MyoD levels. Cells were then probed for p65 and MyoD. α-tubulin was used as a loading control. B. p65+/+ and p65-/- MEFs stably expressing MyoD were differentiated and lysates were then probed for indicated myogenic differentiation markers. C. Cells were differentiated as in B., and MyHC immunofluorescence was performed. D. p65-/- MEFs were transfected with TnI-luc and either vector plasmid, wild type p65 (1-551), or p65 TA mutants (1-521, 1-313), along with MyoD. RLU: Relative Light Units. E. p65-/- MEFs were reconstituted with either vector, full-length or truncated p65, along with MSCV-MyoD. Following selection, whole cell lysates were prepared and probed for p65, MyoD and α-tubulin. F. Cells were infected as in E., differentiated for 72 hr, fixed, and stained for MyHC. G. p65-/- MEFs were transfected with MyoD, TnI-luc, and either vector control, wild type p65 (WT), or p65 constructs containing S/A mutation at positions 276, 529, and 536. MEFs were differentiated and harvested after 48 hr for luciferase assays. H. Relative luciferase activities from p65-/- MEFs transfected with MyoD, TnI-luc and either vector control, WT p65, or p65 (1-313) containing the S276A mutation.

Figure 3.3: Loss of p65 enhances differentiation of primary myoblasts.

A. Primary myoblasts were prepared from 2-4 day old TNFα-/-;p65+/+, TNFα-/-;p65+/- , TNFα-/-;p65-/- neonates and genotypes were verified by westerns for p65. B. p65 and p50 primary myoblasts were transfected with TnI-luc or MyHC-luc plasmids, differentiated for 48 hr and subsequently harvested for luciferase assays. C. TNFα-/-;p65+/+ and TNFα-/-;p65-/- myoblasts were differentiated for 0 hr (GM), or 48 hr (DM), and subsequently stained for MyHC. D. Quantification of myogenesis was performed by scoring MyHC positive cells from a minimum of 25 fields, normalized to total cell number as determined by Hoechst staining. E. Myoblasts were differentiated for 0 (GM) and 48 hr (DM), and lysates were probed for MyHC and troponin. The asterisk indicates troponin expression under GM conditions. F. Primary or C2C12 myoblasts were transfected with siControl (siCont) or siRNA against p65 (sip65) along with Tn-luc reporter. Cells were switched to DM and luciferase assays performed. G. C2C12 myoblasts were transfected with siCont or sip65 and switched to DM for 48h, after which lysates were prepared and westerns performed.

Figure 3.4: Myogenesis is enhanced in p65 deficient mice.

A. H&E-stained cryosections of tibialis anterior (TA), gastrocnemius (Gastroc), and quadriceps (Quad) muscles from TNFα-/-;p65+/+ and TNFα-/-;p65-/- mice or p50+/+ and p50-/- gastrocnemius. B. Fiber diameters were measured from gastrocnemius muscles sections from a total of 1500 fibers (n=5 mice per group). C. Fiber numbers were determined in whole cross sections from TA muscles from TNFα-/-;p65+/+ and TNFα-/-;p65-/- mice (n=3). D. Fiber numbers were recorded from pre-measured randomly selected areas (minimum of 25 per animal) throughout the TA, Gastroc and Quad muscles (n=5 mice per genotype).

Figure 3.5: p65 regulation of myogenesis occurs through multiple mechanisms.

A. C2C12 myoblasts were transfected with control and p65 siRNA and lysates were harvested for western analysis. B. MyoD-/- myoblasts were transfected with TnI-luc along with either an empty vector (CMV-Vect) or a p65 expression plasmid (CMV-p65), or transfected with vector and subsequently treated with 5ng/ml TNFα. Cells were differentiated for 2 days and luciferase assays were performed. C. MyoD-/- myoblasts were infected with pBabe-Puro or pBabe-p65 retroviruses. Following selection and differentiation for 3 and 5 days, protein lysates were prepared for western analysis. D. MyoD-/- myoblasts stably expressing pBabe-Puro and pBabe-p65 were differentiated, fixed and stained for MyHC (red), and nuclei (Hoechst, blue). E. p65+/+ and p65-/- MEFs were transfected with TnI-luc with either MyoD or myogenin plasmids. As a control, transfections were also performed with a p53 expression plasmid and responsive reporter (pGL13-luc). Cells were subsequently differentiated and luciferase assays performed.

Figure 3.6: IKK signaling is temporally regulated and functionally distinct in myogenesis.

A. IKKβ (f/f) MEFs or myoblasts prepared from E13.5 embryos or 3-day old pups, respectively, were infected with pBabe-Puro or pBabe-Cre retrovirus. B. H&E stained cryosections from TA muscles of 4-6 week old IKKβ f/f and IKKβ f/f;MCK-Cre mice. C. Fiber numbers were determined from pre-measured randomly selected areas throughout the TA muscle (n=3 mice per genotype). D. IKK wild type and null MEFs were transiently transfected with MyoD and TnI-luc, and after 2 days in DM lysates were prepared for luciferase assays. E.C2C12 myoblasts were differentiated and lysates prepared for IKK kinase assays using wild type or serine to alanine mutant IκBα proteins as substrates (KA: kinase assay; WB: western blot). F. C2C12 cells were differentiated and at indicated time points, extracts were prepared to probe for phosphorylated IκBα, total IκBα, phosphorylated p65, and total p65. Parallel differentiated C2C12 cells were immunoprecipitated with a p65 antibody, and processed for ChIP. Fragments from the IκBα promoter were amplified by PCR before (input) or after immunoprecipitation. G. Lysates from differentiating C2C12 cells were prepared and used to probe for p100/p52, and α-tubulin. H. MEFs wild type or null for p52 and RelB were transfected with MyoD and TnI-luc and prepared for luciferase assays.

Figure 3.7: IKKα regulates myotube maintenance.

A. C2C12 were transfected with Vector or HA-IKKα expression plasmids. Following selection, cells were differentiated and harvested for western analysis probing for HA and myogenic markers. B. Myoblasts were transfected with siCont or siIKKα oligonucleotides, differentiated and westerns were performed as in A. C. 3-day differentiated myotubes stably expressing Vector or IKKα were subjected to varying stress conditions including no media replenishment for 6 days (6 days in DM) or low glucose (1gram/L glucose in DM for 48 hr). Cells were then fixed and photographed by phase-contrast at 20x magnification. D. Differentiated myotubes stably expressing wild type or kinase dead (KD) version of IKKα were switched to low glucose for 24 hr and photographed by phase contrast. E. Myotubes expressing siCont, siIKKα or siIKKβ were differentiated for 3 days and then switched to low glucose for 20 hr before fixation. Parallel samples were harvested for westerns to confirm knockdown efficiency. F. C2C12 cells expressing vector or IKKα were differentiated for 6 days. Lysates were subsequently prepared for westerns probing for IKKα and myogenic markers.

Figure 3.8: IKKα regulates mitochondrial biogenesis.

A. DNA was prepared from GM or 3-day DM C2C12 cells, diluted, and then used to amplify a 648-bp fragment from MTCO1. Separate PCR for GAPDH was used to normalize for loading. B. C2C12 cells stably expressing Vector (Vect) or IKKα were differentiated and DNA samples were prepared for determination of mitchondrial number as in A. C. Vect and IKKα overexpressing cells were differentiated for 3 days and then stained for mitochondria with Mitotracker Green. Staining was viewed by fluorescence at 20x magnification. D. Mitochondrial and cytoplasmic extracts were prepared from HA-IKKα or IKKα-depleted myotubes and lysates were probed for cytochrome c. E. IKKα expressing or depleted cells were differentiated for 3 days, lysed, and ATP production was measured by luminescence. F. Vect and IKKα cells were differentiated, lysed, and prepared for a citrate synthase assay. All experiments were initiated with equal protein and performed during the linear phase of the reaction to ensure adequate substrate amounts (asterisk represents p= 0.01, left). A parallel set of myotubes expressing Vect or IKKα were cultured, switched to DM, and dehydrogenase activity was measured (asterisk denotes p=0.001). G. C2C12 cells were transfected with Vect, p52/RelB, or p50/p65, and differentiated for 3 days. DNA was prepared as in A., or processed for determination of ATP production as in E. (asterisk denotes p=values of 0.001 and 0.03, respectively).

Figure 3.9: IKKα controls mitochondrial structure.

A. Ultrathin sections from Vect or IKKα expressing myotubes were analyzed by EM at 18500X direct magnification (scale bar =500nm). B. Myotubes expressing control or IKKα siRNA were sectioned and visualized by EM as in A. C. Microarray analysis was performed on Vect and IKKα expressing myotubes using the murine MG 430.20 Affymetrix chip. Genes upregulated in IKKα myotubes as compared to Vect were analyzed by L2L analysis (http://depts.washington.edu/l2l/) for statistically significant enriched biological processes (p<0.05).

Figure 3.10: A model for IKK/NF-κB signaling and function in skeletal myogenesis.

The model depicts different phases of myogenesis, from proliferating myoblasts to differentiated myotubes. In proliferating myoblasts, classical NF-κB signaling mediated by IKKβ and IKKγ leads to the activation of p65 that binds DNA and regulates gene expression to inhibit myogenesis. During differentiation, classical NF-κB is downregulated while the alternative signaling becomes activated. Activation of alternative signaling occurs late in the myogenic program to regulate mitochondrial biogenesis and myotube maintenance.

Figure 3.11: Lack of p65 leads to increased fiber numbers independent of age, fiber type or muscle atrophy.

A. Gastroc muscle cryosections were stained with slow MyHC to differentiate slow (type I, dark), and fast (type II, light) fibers. Black arrows indicate a representative slow fiber, while the blue arrows are representative of a fast fiber. B. Northern blot for the E3 ligases MuRF1 and MAFBx in gastroc muscles from TNFα-/-;p65+/+ and TNFα-/-;p65-/- mice (n=2, each). Expression of MuRF1 was normalized to GAPDH. C. H&E analysis of the lower limb from 9 day- old TNFα-/-;p65+/+ and TNFα-/-;p65-/- mice. D. Fiber numbers were measured from serial sections of the lower limb (lateral to the fibula) of 7 and 9 day-old TNFα-/-;p65+/+ and TNFα-/-;p65-/- neonates (n=1 for each genotype and time point).

Figure 3.12: Conditional deletion of IKKβ in muscles.

A. IKKβ excision post Cre recombinase infection was verified by PCR (upper panel), and westerns (WB, bottom panel). Arrowhead indicates specific IKKβ band. B. Gastroc muscle lysates were prepared from IKKβ f/f and IKKβ f/f; MCK-Cre and probed for IKKβ to verify Cre efficiency.

Figure 3.13: Mitochondrial inhibitors lead to myotube cell death.

A. 10T1/2 fibroblasts were transfected with MyoD, TnI-luc, and either vector (Vect), p65, or IKKα, β, γ wild type or mutant versions (WT, wild type; KD, kinase dead; CA, constitutively active). Fibroblasts were then differentiated for 48 hr and lysates were prepared for luciferase assays. B. 3-day differentiated C2C12 cells were treated for 20 hr with vehicle (control), 250µg/ml chloramphenicol; or 10µg/ml oligomycin after which myotubes were subsequently fixed and photographed at 20x magnification.

REFERENCES

1. Ghosh, S. and M. Karin, Missing pieces in the NF-kappaB puzzle. Cell, 2002. 109 Suppl: p. S81-96.

2. Sen R, B.D., Inducibility of kappa immunoglobulin enhancer-binding protein NF- kappaB by a posttranslational mechanism. Cell, 1986. 47: p. 921-928.

3. Hayden, M.S. and S. Ghosh, Shared principles in NF-kappaB signaling. Cell, 2008. 132(3): p. 344-62.

4. Pahl, H.L., Activators and target genes of Rel/NF-kappaB transcription factors. Oncogene, 1999. 18(49): p. 6853-66.

5. Tong, X., et al., The p50-p50 NF-kappaB complex as a stimulus-specific repressor of gene activation. Mol Cell Biochem, 2004. 265(1-2): p. 171-83.

6. Ghosh, S., M.J. May, and E.B. Kopp, NF-kappa B and Rel proteins: evolutionarily conserved mediators of immune responses. Annu Rev Immunol., 1998. 16: p. 225-260.

7. Perkins, N.D., Post-translational modifications regulating the activity and function of the nuclear factor kappa B pathway. Oncogene, 2006. 25(51): p. 6717- 30.

8. Muta, T., IkappaB-zeta: an inducible regulator of nuclear factor-kappaB. Vitam Horm, 2006. 74: p. 301-16.

9. Carmody, R.J., et al., Negative regulation of toll-like receptor signaling by NF- kappaB p50 ubiquitination blockade. Science, 2007. 317(5838): p. 675-678.

10. Kashatus, D., P. Cogswell, and A.S. Baldwin, Expression of the Bcl-3 proto- oncogene suppresses p53 activation. Genes Dev, 2006. 20(2): p. 225-35.

11. Westerheide, S.D., et al., The putative oncoprotein Bcl-3 induces cyclin D1 to stimulate G(1) transition. Mol Cell Biol, 2001. 21(24): p. 8428-36.

12. Zandi, E., et al., The IkappaB kinase complex (IKK) contains two kinase subunits, IKKalpha and IKKbeta, necessary for IkappaB phosphorylation and NF-kappaB activation. Cell, 1997. 91(2): p. 243-52.

13. Mercurio, F., et al., IKK-1 and IKK-2: cytokine-activated IkappaB kinases essential for NF-kappaB activation. Science, 1997. 278(5339): p. 860-6.

14. May, M.J., R.B. Marienfeld, and S. Ghosh, Characterization of the Ikappa B- kinase NEMO binding domain. J Biol Chem, 2002. 277(48): p. 45992-6000.

15. Makris, C., J.L. Roberts, and M. Karin, The carboxyl-terminal region of IkappaB kinase gamma (IKKgamma) is required for full IKK activation. Mol Cell Biol, 2002. 22(18): p. 6573-81.

16. Sebban, H., S. Yamaoka, and G. Courtois, Posttranslational modifications of NEMO and its partners in NF-kappaB signaling. Trends Cell Biol, 2006. 16(11): p. 569-77.

17. Hacker, H. and M. Karin, Regulation and function of IKK and IKK-related kinases. Sci STKE, 2006. 2006(357): p. re13.

18. Schwabe, R.F. and H. Sakurai, IKKbeta phosphorylates p65 at S468 in transactivaton domain 2. Faseb J, 2005. 19(12): p. 1758-60.

19. Xiao, G., A. Fong, and S.C. Sun, Induction of p100 processing by NF-kappaB- inducing kinase involves docking IkappaB kinase alpha (IKKalpha) to p100 and IKKalpha-mediated phosphorylation. J Biol Chem, 2004. 279(29): p. 30099- 30105.

20. Hoberg, J.E., F. Yeung, and M.W. Mayo, SMRT derepression by the IkappaB kinase alpha: a prerequisite to NF-kappaB transcription and survival. Mol Cell, 2004. 16(2): p. 245-255.

21. Huang, W.C., et al., Phosphorylation of CBP by IKKalpha promotes cell growth by switching the binding preference of CBP from p53 to NF-kappaB. Mol Cell, 2007. 26(1): p. 75-87.

22. Anest, V., et al., A nucleosomal function for IkappaB kinase-alpha in NF-kappaB- dependent gene expression. Nature, 2003. 423(6940): p. 659-663.

23. Prajapati, S., et al., IKKalpha regulates the mitotic phase of the cell cycle by modulating Aurora A phosphorylation. Cell Cycle, 2006. 5(20): p. 2371-80.

24. Lee, N.K. and S.Y. Lee, Modulation of life and death by the tumor necrosis factor receptor-associated factors (TRAFs). J Biochem Mol Biol, 2002. 35(1): p. 61-6.

25. Meylan, E. and J. Tschopp, The RIP kinases: crucial integrators of cellular stress. Trends Biochem Sci, 2005. 30(3): p. 151-9.

26. Hsu, H., et al., TNF-dependent recruitment of the protein kinase RIP to the TNF receptor-1 signaling complex. Immunity, 1996. 4(4): p. 387-96.

27. Gerondakis, S., et al., Genetic approaches in mice to understand Rel/NF-kappaB and IkappaB function: transgenics and knockouts. Oncogene, 1999. 18(49): p. 6888-6895.

28. Pomerantz, J.L. and D. Baltimore, Two pathways to NF-kappaB. Mol Cell, 2002. 10(4): p. 693-695.

29. Senftleben, U., et al., Activation by IKKalpha of a second, evolutionary conserved, NF-kappa B signaling pathway. Science, 2001. 293(5534): p. 1495- 1499.

30. Hauer, J., et al., TNF receptor (TNFR)-associated factor (TRAF) 3 serves as an inhibitor of TRAF2/5-mediated activation of the noncanonical NF-kappaB pathway by TRAF-binding TNFRs. Proc Natl Acad Sci U S A, 2005. 102(8): p. 2874-9.

31. Dejardin, E., The alternative NF-kappaB pathway from biochemistry to biology: pitfalls and promises for future drug development. Biochem Pharmacol, 2006. 72(9): p. 1161-79.

32. Liang, C., M. Zhang, and S.C. Sun, beta-TrCP binding and processing of NF- kappaB2/p100 involve its phosphorylation at serines 866 and 870. Cell Signal, 2006. 18(8): p. 1309-17.

33. Bonizzi, G., et al., Activation of IKKalpha target genes depends on recognition of specific kappaB binding sites by RelB:p52 dimers. Embo J, 2004. 23(21): p. 4202- 4210.

34. Dejardin, E., et al., The lymphotoxin-beta receptor induces different patterns of gene expression via two NF-kappaB pathways. Immunity, 2002. 17(4): p. 525- 535.

35. Asagiri, M. and H. Takayanagi, The molecular understanding of osteoclast differentiation. Bone, 2007. 40(2): p. 251-64.

36. Franzoso, G., et al., Requirement for NF-kappaB in osteoclast and B-cell development. Genes Dev, 1997. 11(24): p. 3482-96.

37. Iotsova, V., et al., Osteopetrosis in mice lacking NF-kappaB1 and NF-kappaB2. Nat Med, 1997. 3(11): p. 1285-1289.

38. Xing, L., et al., NF-kappaB p50 and p52 expression is not required for RANK- expressing osteoclast progenitor formation but is essential for RANK- and cytokine-mediated osteoclastogenesis. J Bone Miner Res, 2002. 17(7): p. 1200-10.

39. Xing, L., et al., Expression of either NF-kappaB p50 or p52 in osteoclast precursors is required for IL-1-induced bone resorption. J Bone Miner Res, 2003. 18(2): p. 260-9.

40. Yamashita, T., et al., NF-kappaB p50 and p52 regulate receptor activator of NF- kappaB ligand (RANKL) and tumor necrosis factor-induced osteoclast precursor differentiation by activating c-Fos and NFATc1. J Biol Chem, 2007. 282(25): p. 18245-53.

41. Novack, D.V., et al., The IkappaB function of NF-kappaB2 p100 controls stimulated osteoclastogenesis. J Exp Med, 2003. 198(5): p. 771-81.

42. Ruocco, M.G., et al., I{kappa}B kinase (IKK){beta}, but not IKK{alpha}, is a critical mediator of osteoclast survival and is required for inflammation-induced bone loss. J Exp Med, 2005. 201(10): p. 1677-1687.

43. Chaisson, M.L., et al., Osteoclast differentiation is impaired in the absence of inhibitor of kappa B kinase alpha. J Biol Chem, 2004. 279(52): p. 54841-8.

44. Vaira, S., et al., RelB is the NF-kappaB subunit downstream of NIK responsible for osteoclast differentiation. Proc Natl Acad Sci U S A, 2008. 105(10): p. 3897- 902.

45. Vaira, S., et al., RelA/p65 promotes osteoclast differentiation by blocking a RANKL-induced apoptotic JNK pathway in mice. J Clin Invest, 2008. 118(6): p. 2088-97.

46. Bottero, V., S. Withoff, and I.M. Verma, NF-kappaB and the regulation of hematopoiesis. Cell Death Differ, 2006. 13(5): p. 785-97.

47. Bugarski, D., et al., Signaling pathways implicated in hematopoietic progenitor cell proliferation and differentiation. Exp Biol Med (Maywood), 2007. 232(1): p. 156-63.

48. Denk, A., T. Wirth, and B. Baumann, NF-kappaB transcription factors: critical regulators of hematopoiesis and neuronal survival. Cytokine Growth Factor Rev, 2000. 11(4): p. 303-320.

49. Grumont, R.J., et al., B lymphocytes differentially use the Rel and nuclear factor kappaB1 (NF-kappaB1) transcription factors to regulate cell cycle progression and apoptosis in quiescent and mitogen-activated cells. J Exp Med, 1998. 187(5): p. 663-74.

50. Pohl, T., et al., The combined absence of NF-kappa B1 and c-Rel reveals that overlapping roles for these transcription factors in the B cell lineage are restricted to the activation and function of mature cells. Proc Natl Acad Sci U S A, 2002. 99(7): p. 4514-9.

51. Grossmann, M., et al., The combined absence of the transcription factors Rel and RelA leads to multiple hemopoietic cell defects. Proc Natl Acad Sci U S A, 1999. 96(21): p. 11848-53.

52. Horwitz, B.H., et al., Failure of lymphopoiesis after adoptive transfer of NF- kappaB-deficient fetal liver cells. Immunity, 1997. 6(6): p. 765-72.

53. Grossmann, M., et al., The anti-apoptotic activities of Rel and RelA required during B-cell maturation involve the regulation of Bcl-2 expression. Embo J, 2000. 19(23): p. 6351-60.

54. Liou, H.C., Regulation of the immune system by NF-kappaB and IkappaB. J Biochem Mol Biol, 2002. 35(6): p. 537-546.

55. Buckingham, M., et al., The formation of skeletal muscle: from somite to limb. J Anat, 2003. 202(1): p. 59-68.

56. Le Grand, F. and M.A. Rudnicki, Skeletal muscle satellite cells and adult myogenesis. Curr Opin Cell Biol, 2007. 19(6): p. 628-33.

57. Pownall, M.E., M.K. Gustafsson, and C.P. Emerson, Jr., Myogenic regulatory factors and the specification of muscle progenitors in vertebrate embryos. Annu Rev Cell Dev Biol, 2002. 18: p. 747-783.

58. Tapscott, S.J., The circuitry of a master switch: Myod and the regulation of skeletal muscle gene transcription. Development, 2005. 132(12): p. 2685-95.

59. Wang, D.Z., et al., The Mef2c gene is a direct transcriptional target of myogenic bHLH and MEF2 proteins during skeletal muscle development. Development, 2001. 128(22): p. 4623-33.

60. Lassar, A.B., et al., Functional activity of myogenic HLH proteins requires hetero-oligomerization with E12/E47-like proteins in vivo. Cell, 1991. 66(2): p. 305-15.

61. Puri, P.L., et al., Differential roles of p300 and PCAF acetyltransferases in muscle differentiation. Mol Cell, 1997. 1(1): p. 35-45.

62. Simone, C., et al., p38 pathway targets SWI-SNF chromatin-remodeling complex to muscle-specific loci. Nat Genet, 2004. 36(7): p. 738-43.

63. Melnikova, I.N., et al., Differential biological activities of mammalian Id proteins in muscle cells. Exp Cell Res, 1999. 247(1): p. 94-104.

64. Palacios, D. and P.L. Puri, The epigenetic network regulating muscle development and regeneration. J Cell Physiol, 2006. 207(1): p. 1-11.

65. Fulco, M., et al., Sir2 regulates skeletal muscle differentiation as a potential sensor of the redox state. Mol Cell, 2003. 12(1): p. 51-62.

66. Caretti, G., et al., The Polycomb Ezh2 methyltransferase regulates muscle gene expression and skeletal muscle differentiation. Genes Dev, 2004. 18(21): p. 2627- 38.

67. Wang, H., et al., NF-kappaB regulation of YY1 inhibits skeletal myogenesis through transcriptional silencing of myofibrillar genes. Mol Cell Biol, 2007. 27(12): p. 4374-4387.

68. Lluis, F., et al., Regulation of skeletal muscle gene expression by p38 MAP kinases. Trends Cell Biol, 2006. 16(1): p. 36-44.

69. Keren, A., Y. Tamir, and E. Bengal, The p38 MAPK signaling pathway: a major regulator of skeletal muscle development. Mol Cell Endocrinol, 2006. 252(1-2): p. 224-30.

70. Perdiguero, E., et al., Genetic analysis of p38 MAP kinases in myogenesis: fundamental role of p38alpha in abrogating myoblast proliferation. Embo J, 2007. 26(5): p. 1245-56.

71. Suelves, M., et al., Phosphorylation of MRF4 transactivation domain by p38 mediates repression of specific myogenic genes. Embo J, 2004. 23(2): p. 365-375.

72. Guttridge, D.C., Signaling pathways weigh in on decisions to make or break skeletal muscle. Curr Opin Clin Nutr Metab Care., 2004. 7(4): p. 443-450.

73. Rommel, C., et al., Mediation of IGF-1-induced skeletal myotube hypertrophy by PI(3)K/Akt/mTOR and PI(3)K/Akt/GSK3 pathways. Nature Cell Biol., 2001. 3(11): p. 1009-1013.

74. Olwin, B.B., K. Hannon, and A.J. Kudla, Are fibroblast growth factors regulators of myogenesis in vivo? Prog Growth Factor Res, 1994. 5(2): p. 145-58. 86

75. Armand, A.S., I. Laziz, and C. Chanoine, FGF6 in myogenesis. Biochim Biophys Acta, 2006. 1763(8): p. 773-8.

76. Vaidya, T.B., et al., Fibroblast growth factor and transforming growth factor beta repress transcription of the myogenic regulatory gene MyoD1. Mol Cell Biol, 1989. 9(8): p. 3576-9.

77. Taylor, D.A., et al., E1A-mediated inhibition of myogenesis correlates with a direct physical interaction of E1A12S and basic helix-loop-helix proteins. Mol Cell Biol, 1993. 13(8): p. 4714-27.

78. Lassar, A.B., et al., Transformation by activated ras or fos prevents myogenesis by inhibiting expression of MyoD1. Cell, 1989. 58(4): p. 659-67.

79. Weyman, C.M., et al., Distinct signaling pathways regulate transformation and inhibition of skeletal muscle differentiation by oncogenic Ras. Oncogene, 1997. 14(6): p. 697-704.

80. Lee, S.-J., Regulation of Muscle Mass by Myostatin. Ann. Rev. Cell Dev. Biol., 2004. 20: p. 61-86.

81. Kollias, H.D. and J.C. McDermott, Transforming growth factor-beta and myostatin signaling in skeletal muscle. J Appl Physiol, 2008. 104(3): p. 579-87.

82. Walsh, F.S. and A.J. Celeste, Myostatin: a modulator of skeletal-muscle stem cells. Biochem Soc Trans, 2005. 33(Pt 6): p. 1513-7.

83. McPherron, A.C., A.M. Lawler, and S.J. Lee, Regulation of skeletal muscle mass in mice by a new TGF-beta superfamily member. Nature, 1997. 387(6628): p. 83- 90.

84. Artaza, J.N., et al., Endogenous expression and localization of myostatin and its relation to myosin heavy chain distribution in C2C12 skeletal muscle cells. J. Cell Physiol., 2002. 190(2): p. 170-179.

85. Lee, S.J. and A.C. McPherron, Regulation of myostatin activity and muscle growth. Proc. Natl Acad. Sci. USA, 2001. 98(16): p. 9306-9311.

86. Welle, S., et al., Muscle growth after postdevelopmental myostatin gene knockout. Am J Physiol Endocrinol Metab, 2007. 292(4): p. E985-91.

87. Siriett, V., et al., Antagonism of myostatin enhances muscle regeneration during sarcopenia. Mol Ther, 2007. 15(8): p. 1463-70.

88. Ji, M., et al., Myostatin induces p300 degradation to silence cyclin D1 expression through the PI3K/PTEN/Akt pathway. Cell Signal, 2008. 20(8): p. 1452-8. 87

89. Thomas, M., et al., Myostatin, a negative regulator of muscle growth, functions by inhibiting myoblast proliferation. J Biol. Chem., 2000. 275(51): p. 40235-40243.

90. Taylor, W.E., et al., Myostatin inhibits cell proliferation and protein synthesis in C2C12 muscle cells. Am. J Physiol Endocrinol. Metab., 2001. 280(2): p. E221- E228.

91. Rios, R., et al., Myostatin is an inhibitor of myogenic differentiation. Am. J. Physiol. Cell Physiol., 2002. 282(5): p. C993-C999.

92. Joulia, D., et al., Mechanisms involved in the inhibition of myoblast proliferation and differentiation by myostatin. Exp. Cell Res., 2003. 286(2): p. 263-275.

93. Langley, B., et al., Myostatin inhibits myoblast differentiation by down-regulating MyoD expression. J Biol. Chem., 2002. 277(51): p. 49831-49840.

94. McFarlane, C., et al., Myostatin signals through Pax7 to regulate satellite cell self-renewal. Exp Cell Res, 2008. 314(2): p. 317-29.

95. Guttridge, D.C., et al., NF-kappaB controls cell growth and differentiation through transcriptional regulation of cyclin D1. Mol. Cell. Biol., 1999. 19(8): p. 5785-5799.

96. Judge, A.R., et al., Role for IkappaBalpha, but not c-Rel, in skeletal muscle atrophy. Am J Physiol Cell Physiol, 2007. 292(1): p. C372-82.

97. Hunter, R.B., et al., Activation of an alternative NF-kappaB pathway in skeletal muscle during disuse atrophy. FASEB J., 2002. 16(6): p. 529-538.

98. Bakkar, N., et al., IKK/NF-kappaB regulates skeletal myogenesis via a signaling switch to inhibit differentiation and promote mitochondrial biogenesis. J Cell Biol, 2008. 180(4): p. 787-802.

99. De Alvaro, C., et al., Nuclear exclusion of forkhead box O and Elk1 and activation of nuclear factor-kappaB are required for C2C12-RasV12C40 myoblast differentiation. Endocrinology, 2008. 149(2): p. 793-801.

100. Baeza-Raja, B. and P. Munoz-Canoves, p38 MAPK-induced nuclear factor- kappaB activity is required for skeletal muscle differentiation: role of interleukin- 6. Mol Biol Cell, 2004. 15(4): p. 2013-2026.

101. Caamano, J.H., et al., Nuclear factor (NF)-kappa B2 (p100/p52) is required for normal splenic microarchitecture and B cell-mediated immune responses. J Exp Med, 1998. 187(2): p. 185-196.

102. Canicio, J., et al., Nuclear factor kappa B-inducing kinase and Ikappa B kinase- alpha signal skeletal muscle cell differentiation. J. Biol. Chem., 2001. 276(23): p. 20228-20233.

103. Conejo, R., et al., Insulin restores differentiation of Ras-transformed C2C12 myoblasts by inducing NF-kappaB through an AKT/P70S6K/p38-MAPK pathway. Oncogene, 2002. 21(23): p. 3739-3753.

104. Conejo, R., et al., Insulin produces myogenesis in C2C12 myoblasts by induction of NF-kappaB and downregulation of AP-1 activities. J. Cell. Physiol., 2001. 186(1): p. 82-94.

105. Kaliman, P., et al., Insulin-like growth factor-II, phosphatidylinositol 3-kinase, nuclear factor-kappaB and inducible nitric-oxide synthase define a common myogenic signaling pathway. J. Biol. Chem., 1999. 274(25): p. 17437-17444.

106. Lee, K.H., et al., NF-kappaB-dependent expression of nitric oxide synthase is required for membrane fusion of chick embryonic myoblasts. Biochem J, 1997. 324 ( Pt 1): p. 237-42.

107. Kim, S.S., et al., Inhibitors of the proteasome block the myogenic differentiation of rat L6 myoblasts. FEBS Lett, 1998. 433(1-2): p. 47-50.

108. Hirasaka, K., et al., Clinorotation prevents differentiation of rat myoblastic L6 cells in association with reduced NF-kappa B signaling. Biochim Biophys Acta, 2005. 1743(1-2): p. 130-40.

109. Lehtinen, S.K., et al., Down-regulation of transcription factors AP-1, Sp-1, and NF-kappa B precedes myocyte differentiation. Biochem Biophys Res Commun, 1996. 229(1): p. 36-43.

110. Catani, M.V., et al., Nuclear factor kappaB and activating protein 1 are involved in differentiation-related resistance to oxidative stress in skeletal muscle cells. Free Radic Biol Med, 2004. 37(7): p. 1024-1036.

111. Dee, K., A. DeChant, and C.M. Weyman, Differential signaling through NFkappaB does not ameliorate skeletal myoblast apoptosis during differentiation. FEBS Lett., 2003. 545(2-3): p. 246-252.

112. Bakkar, N., H. Wackerhage, and D.C. Guttridge, Myostatin and NF-kB Regulate Skeletal Myogenesis Through Distinct Signaling Pathways. Signal Transduction, 2005. 4(202-210).

113. Guttridge, D.C., et al., NF-kB-Induced Loss of MyoD Messenger RNA: Possible Role in Muscle Decay and Cachexia. Science, 2000. 289(5488): p. 2363-2366.

114. Girgenrath, M., et al., TWEAK, via its receptor Fn14, is a novel regulator of mesenchymal progenitor cells and skeletal muscle regeneration. Embo J, 2006. 25(24): p. 5826-39.

115. Langen, R.C., et al., Inflammatory cytokines inhibit myogenic differentiation through activation of nuclear factor-kappaB. FASEB J, 2001. 15(7): p. 1169- 1180.

116. Dogra, C., et al., Tumor necrosis factor-like weak inducer of apoptosis inhibits skeletal myogenesis through sustained activation of nuclear factor-kappaB and degradation of MyoD protein. J Biol Chem, 2006. 281(15): p. 10327-10336.

117. Munz, B., et al., RIP2, a checkpoint in myogenic differentiation. Mol Cell Biol, 2002. 22(16): p. 5879-5886.

118. Ardite, E., et al., Glutathione depletion impairs myogenic differentiation of murine skeletal muscle C2C12 cells through sustained NF-kappaB activation. Am J Pathol, 2004. 165(3): p. 719-728.

119. Kumar, A., et al., Cyclic mechanical strain inhibits skeletal myogenesis through activation of focal adhesion kinase, Rac-1 GTPase, and NF-kappaB transcription factor. Faseb J, 2004. 18(13): p. 1524-35.

120. Sitcheran, R., P.C. Cogswell, and A.S. Baldwin, Jr., NF-kappaB mediates inhibition of mesenchymal cell differentiation through a posttranscriptional gene silencing mechanism. Genes Dev., 2003. 17(19): p. 2368-2373.

121. Dhawan, J. and T.A. Rando, Stem cells in postnatal myogenesis: molecular mechanisms of satellite cell quiescence, activation and replenishment. Trends Cell Biol, 2005. 15(12): p. 666-73.

122. Charge, S.B. and M.A. Rudnicki, Cellular and molecular regulation of muscle regeneration. Physiol Rev, 2004. 84(1): p. 209-38.

123. Wagers, A.J. and I.M. Conboy, Cellular and molecular signatures of muscle regeneration: current concepts and controversies in adult myogenesis. Cell, 2005. 122(5): p. 659-67.

124. Grefte, S., et al., Skeletal muscle development and regeneration. Stem Cells Dev, 2007. 16(5): p. 857-68.

125. Mourkioti, F., et al., Targeted ablation of IKK2 improves skeletal muscle strength, maintains mass, and promotes regeneration. J Clin Invest, 2006. 116(11): p. 2945-2954.

126. Acharyya, S., et al., Interplay of IKK/NF-kappaB signaling in macrophages and myofibers promotes muscle degeneration in Duchenne muscular dystrophy. J Clin Invest, 2007. 117(4): p. 889-901.

127. Collins, R.A. and M.D. Grounds, The role of tumor necrosis factor-alpha (TNF- alpha) in skeletal muscle regeneration. Studies in TNF-alpha(-/-) and TNF- alpha(-/-)/LT-alpha(-/-) mice. J Histochem Cytochem, 2001. 49(8): p. 989-1001.

128. Chen, S.E., et al., Role of TNF-{alpha} signaling in regeneration of cardiotoxin- injured muscle. Am J Physiol Cell Physiol, 2005. 289(5): p. C1179-1187.

129. Pelosi, L., et al., Local expression of IGF-1 accelerates muscle regeneration by rapidly modulating inflammatory cytokines and chemokines. Faseb J, 2007. 21(7): p. 1393-402.

130. Messina, S., et al., Nuclear factor kappa-B blockade reduces skeletal muscle degeneration and enhances muscle function in Mdx mice. Exp Neurol, 2006. 198(1): p. 234-41.

131. Thaloor, D., et al., Systemic administration of the NF-kappaB inhibitor curcumin stimulates muscle regeneration after traumatic injury. Amer. J. Physiol., 1999. 277(2 Pt 1): p. C320-329.

132. Ryan, M.T. and N.J. Hoogenraad, Mitochondrial-Nuclear Communications. Annu Rev Biochem, 2007.

133. Chan, D.C., Mitochondria: dynamic organelles in disease, aging, and development. Cell, 2006. 125(7): p. 1241-52.

134. Berdanier, C.D., Mitochondrial gene expression: influence of nutrients and hormones. Exp Biol Med (Maywood), 2006. 231(10): p. 1593-601.

135. Frazier, A.E., et al., Mitochondrial morphology and distribution in mammalian cells. Biol Chem, 2006. 387(12): p. 1551-8.

136. Zeviani, M. and S. Di Donato, Mitochondrial disorders. Brain, 2004. 127(Pt 10): p. 2153-72.

137. Schapira, A.H., Mitochondrial disease. Lancet, 2006. 368(9529): p. 70-82.

138. Kelly, D.P. and R.C. Scarpulla, Transcriptional regulatory circuits controlling mitochondrial biogenesis and function. Genes Dev, 2004. 18(4): p. 357-68.

139. Puigserver, P., et al., A cold-inducible coactivator of nuclear receptors linked to adaptive thermogenesis. Cell, 1998. 92(6): p. 829-39.

140. Lin, J., C. Handschin, and B.M. Spiegelman, Metabolic control through the PGC- 1 family of transcription coactivators. Cell Metab, 2005. 1(6): p. 361-370.

141. Lin, J., et al., Transcriptional co-activator PGC-1 alpha drives the formation of slow-twitch muscle fibres. Nature, 2002. 418(6899): p. 797-801.

142. Lyons, C.N., S.C. Leary, and C.D. Moyes, Bioenergetic remodeling during cellular differentiation: changes in cytochrome c oxidase regulation do not affect the metabolic phenotype. Biochem Cell Biol, 2004. 82(3): p. 391-399.

143. Moyes, C.D., et al., Mitochondrial biogenesis during cellular differentiation. Am J Physiol, 1997. 272(4 Pt 1): p. C1345-1351.

144. Kraft, C.S., et al., Control of mitochondrial biogenesis during myogenesis. Am J Physiol Cell Physiol, 2006. 290(4): p. C1119-1127.

145. Hood, D.A., et al., Coordination of metabolic plasticity in skeletal muscle. J Exp Biol, 2006. 209(Pt 12): p. 2265-75.

146. Jager, S., et al., AMP-activated protein kinase (AMPK) action in skeletal muscle via direct phosphorylation of PGC-1alpha. Proc Natl Acad Sci U S A, 2007. 104(29): p. 12017-22.

147. Arany, Z., et al., The transcriptional coactivator PGC-1beta drives the formation of oxidative type IIX fibers in skeletal muscle. Cell Metab, 2007. 5(1): p. 35-46.

148. Mortensen, O.H., et al., PGC-1alpha and PGC-1beta have both similar and distinct effects on myofiber switching toward an oxidative phenotype. Am J Physiol Endocrinol Metab, 2006. 291(4): p. E807-16.

149. St-Pierre, J., et al., Bioenergetic analysis of peroxisome proliferator-activated receptor gamma coactivators 1alpha and 1beta (PGC-1alpha and PGC-1beta) in muscle cells. J Biol Chem, 2003. 278(29): p. 26597-603.

150. Lelliott, C.J., et al., Ablation of PGC-1beta results in defective mitochondrial activity, thermogenesis, hepatic function, and cardiac performance. PLoS Biol, 2006. 4(11): p. e369.

151. Handschin, C., et al., Skeletal muscle fiber-type switching, exercise intolerance, and myopathy in PGC-1alpha muscle-specific knock-out animals. J Biol Chem, 2007. 282(41): p. 30014-21.

152. Baldwin, A.S., Control of oncogenesis and cancer therapy resistance by the transcription factor NF-kappaB. J Clin Invest, 2001. 107(3): p. 241-246.

153. Karin, M. and Y. Ben-Neriah, Phosphorylation meets ubiquitination: the control of NF-[kappa]B activity. Annu Rev Immunol., 2000. 18: p. 621-663.

154. Senftleben, U. and M. Karin, The IKK/NF-kappa B pathway. Crit Care Med, 2002. 30(1 Suppl): p. S18-26.

155. Sabourin, L.A. and M.A. Rudnicki, The molecular regulation of myogenesis. Clin. Genet., 2000. 57(1): p. 16-25.

156. Naya, F.S. and E. Olson, MEF2: a transcriptional target for signaling pathways controlling skeletal muscle growth and differentiation. Curr Opin Cell Biol, 1999. 11(6): p. 683-688.

157. Myer, A., E.N. Olson, and W.H. Klein, MyoD cannot compensate for the absence of myogenin during skeletal muscle differentiation in murine embryonic stem cells. Dev Biol, 2001. 229(2): p. 340-350.

158. Li, Y.P., et al., Skeletal muscle myocytes undergo protein loss and reactive oxygen-mediated NF-kappaB activation in response to tumor necrosis factor alpha. FASEB J., 1998. 12(10): p. 871-880.

159. Li, Y.P. and M.B. Reid, NF-kappaB mediates the protein loss induced by TNF- alpha in differentiated skeletal muscle myotubes. Am J Physiol Regul Integr Comp Physiol, 2000. 279(4): p. R11650-11670.

160. Ladner, K.J., M.A. Caligiuri, and D.C. Guttridge, Tumor necrosis factor- regulated biphasic activation of NF-kappa B is required for cytokine-induced loss of skeletal muscle gene products. J. Biol. Chem., 2003. 278(4): p. 2294-2303.

161. Gonzalez-Cadavid, N.F. and S. Bhasin, Role of myostatin in metabolism. Curr Opin Clin Nutr Metab Care., 2004. 7(4): p. 451-457.

162. Lee, S.J. and A.C. McPherron, Myostatin and the control of skeletal muscle mass. Curr Opin Genet Dev, 1999. 9(5): p. 604-607.

163. Sharma, M., et al., Myostatin in muscle growth and repair. Exerc. Sport Sci. Rev., 2001. 29(4): p. 155-158.

164. McPherron, A.C. and S.J. Lee, Double muscling in cattle due to mutations in the myostatin gene. Proc. Natl Acad. Sci. USA, 1997. 94(23): p. 12457-61.

165. Grobet, L., et al., A deletion in the bovine myostatin gene causes the double- muscled phenotype in cattle. Nature Genet., 1997. 17(1): p. 71-74.

166. Reisz-Porszasz, S., et al., Lower skeletal muscle mass in male transgenic mice with muscle-specific overexpression of myostatin. Am J Physiol. Endocrinol. Metab., 2003. 285(4): p. E876-E888.

167. Zhu, X., et al., Dominant negative myostatin produces hypertrophy without hyperplasia in muscle. FEBS Lett., 2000. 474(1): p. 71-75.

168. Schuelke, M., et al., Myostatin mutation associated with gross muscle hypertrophy in a child. N Engl J Med., 2004. 350(26): p. 2682-2688.

169. Roth, S.M. and S. Walsh, Myostatin: a therapeutic target for skeletal muscle wasting. Curr Opin Clin Nutr Metab Care., 2004. 7(3): p. 259-263.

170. Reardon, K.A., et al., Myostatin, insulin-like growth factor-1, and leukemia inhibitory factor mRNAs are upregulated in chronic human disuse muscle atrophy. Muscle Nerve, 2001. 24(7): p. 893-899.

171. Zachwieja, J.J., et al., Plasma myostatin-immunoreactive protein is increased after prolonged bed rest with low-dose T3 administration. J. Grav. Physiol., 1999. 6(2): p. 11-15.

172. Gonzalez-Cadavid, N.F., et al., Organization of the human myostatin gene and expression in healthy men and HIV-infected men with muscle wasting. Proc. Natl. Acad Sci. USA, 1998. 95(25): p. 14938-14943.

173. Whittemore, L.A., et al., Inhibition of myostatin in adult mice increases skeletal muscle mass and strength. Biochem. Biophys. Res. Commun., 2003. 300(4): p. 965-971.

174. Wagner, K.R., et al., Loss of myostatin attenuates severity of muscular dystrophy in mdx mice. Ann Neurol, 2002. 52(6): p. 832-836.

175. Bogdanovich, S., et al., Functional improvement of dystrophic muscle by myostatin blockade. Nature, 2002. 420(6914): p. 418-421.

176. Zimmers, T.A., et al., Induction of cachexia in mice by systemically administered myostatin. Science, 2002. 296(5572): p. 1486-1488.

177. McCroskery, S., et al., Myostatin negatively regulates satellite cell activation and self-renewal. J. Cell Biol., 2003. 162(6): p. 1135-1147.

178. Zhu, X., et al., Myostatin signaling through Smad2, Smad3 and Smad4 is regulated by the inhibitory Smad7 by a negative feedback mechanism. Cytokine, 2004. 26(6): p. 262-272.

179. Han, S.H., et al., Transforming growth factor-beta 1 (TGF-beta1) promotes IL-2 mRNA expression through the up-regulation of NF-kappaB, AP-1 and NF-AT in EL4 cells. J Pharmacol Exp Ther., 1998. 287(3): p. 1105-1112.

180. Azuma, M., et al., TGF-beta1 inhibits NF-kappaB activity through induction of IkappaB-alpha expression in human salivary gland cells: a possible mechanism of growth suppression by TGF-beta1. Exp. Cell Res., 1999. 250(1): p. 213-222.

181. Arsura, M., et al., Transient activation of NF-kappaB through a TAK1/IKK kinase pathway by TGF-beta1 inhibits AP-1/SMAD signaling and apoptosis: implications in liver tumor formation. Oncogene, 2003. 22(3): p. 412-425.

182. Bitzer, M., et al., A mechanism of suppression of TGF-beta/SMAD signaling by NF-kappa B/RelA. Genes Dev., 2000. 14(2): p. 187-197.

183. Nagarajan, R.P., et al., Repression of transforming-growth-factor-beta-mediated transcription by nuclear factor kappaB. Biochem J, 2000. 348 Pt 3: p. 591-596.

184. Ma, K., et al., Characterization of 5'-regulatory region of human myostatin gene: regulation by dexamethasone in vitro. Am J Physiol. Endocrinol. Metab., 2001. 281(6): p. E1128-E1136.

185. Finco, T.S., et al., Oncogenic Ha-Ras-induced signaling activates NF-kappaB transcriptional activity, which is required for cellular transformation. J Biol Chem, 1997. 272(39): p. 24113-24116.

186. Madrid, L.V., et al., Akt suppresses apoptosis by stimulating the transactivation potential of the RelA/p65 subunit of NF-kappaB. Mol Cell Biol, 2000. 20(5): p. 1626-1638.

187. Rios, R., et al., Differential response to exogenous and endogenous myostatin in myoblasts suggests that myostatin acts as an autocrine factor in vivo. Endocrinology, 2004. 145(6): p. 2795-2803.

188. Bengal, E., et al., Functional antagonism between c-Jun and MyoD proteins: a direct physical association. Cell, 1992. 68(3): p. 507-519.

189. Hirsinger, E., et al., Notch signalling acts in postmitotic avian myogenic cells to control MyoD activation. Development, 2001. 128(1): p. 107-116.

190. Spiller, M.P., et al., The myostatin gene is a downstream target gene of basic helix-loop-helix transcription factor MyoD. Mol. Cell. Biol., 2002. 22(20): p. 7066-7082.

191. Kocamis, H., et al., Follistatin alters myostatin gene expression in C2C12 muscle cells. Acta Vet Hung, 2004. 52(2): p. 135-141.

192. Hayden, M.S. and S. Ghosh, Signaling to NF-kB. Genes Dev., 2004. 18: p. 2195- 2224.

193. Huxford, T., et al., The crystal structure of the IkappaBalpha/NF-kappaB complex reveals mechanisms of NF-kappaB inactivation. Cell, 1998. 95(6): p. 759-770.

194. Karin, M., How NF-kappaB is activated: the role of the IkappaB kinase (IKK) complex. Oncogene, 1999. 18(49): p. 6867-6874.

195. Rudolph, D., et al., Severe liver degeneration and lack of NF-kappaB activation in NEMO/IKKgamma-deficient mice. Genes Dev, 2000. 14(7): p. 854-862.

196. Beg, A.A., et al., Embryonic lethality and liver degeneration in mice lacking the RelA component of NF-kappa B. Nature, 1995. 376(6536): p. 167-170.

197. Tanaka, M., et al., Embryonic lethality, liver degeneration, and impaired NF- kappa B activation in IKK-beta-deficient mice. Immunity, 1999. 10(4): p. 421- 429.

198. Xiao, G., E.W. Harhaj, and S.C. Sun, NF-kappaB-inducing kinase regulates the processing of NF-kappaB2 p100. Mol Cell, 2001. 7(2): p. 401-409.

199. Weih, F., et al., Both multiorgan inflammation and myeloid hyperplasia in RelB- deficient mice are T cell dependent. J Immunol, 1996. 157(9): p. 3974-3979.

200. Sil, A.K., et al., IkappaB kinase-alpha acts in the epidermis to control skeletal and craniofacial morphogenesis. Nature, 2004. 428(6983): p. 660-664.

201. Hu, Y., et al., IKKalpha controls formation of the epidermis independently of NF- kappaB. Nature, 2001. 410(6829): p. 710-714.

202. Takeda, K., et al., Limb and skin abnormalities in mice lacking IKKalpha. Science, 1999. 284(5412): p. 313-316.

203. Cai, D., et al., IKKbeta/NF-kB activation causes severe muscle wasting in mice. Cell, 2004. 119: p. 285-298.

204. Hunter, R.B. and S.C. Kandarian, Disruption of either the Nfkb1 or the Bcl3 gene inhibits skeletal muscle atrophy. J Clin Invest, 2004. 114(10): p. 1504-1511.

205. Baghdiguian, S., et al., Calpain 3 deficiency is associated with myonuclear apoptosis and profound perturbation of the IkappaB alpha/NF-kappaB pathway in limb-girdle muscular dystrophy type 2A. Nat Med, 1999. 5(5): p. 503-511.

206. Kumar, A. and A.M. Boriek, Mechanical stress activates the nuclear factor- kappaB pathway in skeletal muscle fibers: a possible role in Duchenne muscular dystrophy. FASEB J., 2003. 17(3): p. 386-396.

207. Monici, M.C., et al., Activation of nuclear factor-kappaB in inflammatory myopathies and Duchenne muscular dystrophy. Neurology, 2003. 60(6): p. 993- 997.

208. Davis, R.L., H. Weintraub, and A.B. Lassar, Expression of a single transfected cDNA converts fibroblasts to myoblasts. Cell, 1987. 51(6): p. 987-1000.

209. Novitch, B.G., et al., pRb is required for MEF2-dependent gene expression as well as cell-cycle arrest during skeletal muscle differentiation. Curr Biol, 1999. 9(9): p. 449-459.

210. Schmitz, M.L., et al., Structural and functional analysis of the NF-kappa B p65 C terminus. An acidic and modular transactivation domain with the potential to adopt an alpha-helical conformation. J Biol Chem, 1994. 269(41): p. 25613- 25620.

211. Zhong, H., R.E. Voll, and S. Ghosh, Phosphorylation of NF-kappa B p65 by PKA stimulates transcriptional activity by promoting a novel bivalent interaction with the coactivator CBP/p300. Mol. Cell, 1998. 1(5): p. 661-671.

212. Schmitz, M.L., S. Bacher, and M. Kracht, I kappa B-independent control of NF- kappa B activity by modulatory phosphorylations. Trends Biochem Sci, 2001. 26(3): p. 186-190.

213. Doi, T.S., et al., Absence of tumor necrosis factor rescues RelA-deficient mice from embryonic lethality. Proc Natl Acad Sci U S A, 1999. 96(6): p. 2994-2999.

214. Chen, S.E., B. Jin, and Y.P. Li, TNF-alpha regulates myogenesis and muscle regeneration by activating p38 MAPK. Am J Physiol Cell Physiol, 2007. 292(5): p. C1660-1671.

215. Sabourin, L.A., et al., Reduced differentiation potential of primary MyoD-/- myogenic cells derived from adult skeletal muscle. J Cell Biol, 1999. 144(4): p. 631-643.

216. Gerber, A.N., et al., Two domains of MyoD mediate transcriptional activation of genes in repressive chromatin: a mechanism for lineage determination in myogenesis. Genes Dev, 1997. 11(4): p. 436-450.

217. Newman, J.C. and A.M. Weiner, L2L: a simple tool for discovering the hidden significance in microarray expression data. Genome Biol, 2005. 6(9): p. R81.

218. Yamamoto, Y., et al., Histone H3 phosphorylation by IKK-alpha is critical for cytokine-induced gene expression. Nature, 2003. 423(6940): p. 655-659.

219. Demicco, E.G., et al., RelB/p52 NF-kappaB complexes rescue an early delay in mammary gland development in transgenic mice with targeted superrepressor IkappaB-alpha expression and promote carcinogenesis of the mammary gland. Mol Cell Biol, 2005. 25(22): p. 10136-10147.

220. Acharyya, S., et al., Cancer cachexia is regulated by selective targeting of skeletal muscle gene products. J Clin Invest., 2004. 114(3): p. 370-378.

221. Hertlein, E., et al., RelA/p65 regulation of IkappaBbeta. Mol Cell Biol, 2005. 25(12): p. 4956-4968.

222. Li, Z.W., et al., IKK beta is required for peripheral B cell survival and proliferation. J Immunol, 2003. 170(9): p. 4630-4637.

223. Rando, T.A. and H.M. Blau, Primary mouse myoblast purification, characterization, and transplantation for cell-mediated gene therapy. J Cell Biol, 1994. 125(6): p. 1275-1287.

224. Acharyya, S., et al., Dystrophin glycoprotein complex dysfunction: a regulatory link between muscular dystrophy and cancer cachexia. Cancer Cell, 2005. 8(5): p. 421-432.

225. Leek, B.T., et al., Effect of acute exercise on citrate synthase activity in untrained and trained human skeletal muscle. Am J Physiol Regul Integr Comp Physiol, 2001. 280(2): p. R441-447.

226. Huo, L. and R.C. Scarpulla, Mitochondrial DNA instability and peri-implantation lethality associated with targeted disruption of nuclear respiratory factor 1 in mice. Mol Cell Biol, 2001. 21(2): p. 644-654.

227. Liu, T., B. Brouha, and D. Grossman, Rapid induction of mitochondrial events and caspase-independent apoptosis in Survivin-targeted melanoma cells. Oncogene, 2004. 23(1): p. 39-48.