Stem Cells and Cell Signalling in Skeletal Myogenesis, Volume 11 (Advances in Developmental Biology and Biochemistry, V. 11)

G E N E R A L O V E R V I E W OF THIS V O L U M E Skeletal muscle development is perhaps one of the best understood pr...

Author: D.A. Sassoon

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G E N E R A L O V E R V I E W OF THIS V O L U M E

Skeletal muscle development is perhaps one of the best understood processes at the molecular, cellular and organismal level due in large part to the fact that primary myogenic cells Imyoblasts) will grow and subsequently differentiate into myotubes in culture. This experimental advantage which was pioneered over 30 years ago, coupled with intense efforts to elucidate the causes of multiple muscle myopathies has lead to the identification of growth factors, transcription factors and cell-cell interactions that control this process. With the advent of reverse mouse genetics, many of the observations gained through the study of myogenic cells in vitro have been directly tested in vivo. What has emerged is a complex but cohesive story of how myogenic cells are initially specified in the vertebrate embryo and how muscle fibers ultimately achieve their respective identities (i.e. fast versus slow) to perform their function. While this collection of chapters is focused on these developments, it should be noted that the general paradigms that have emerged have been useful in the study of other tissue types. Skeletal muscle, like nerve cells, undergoes a permanent exit from the cell cycle upon differentiation, and it is generally thought that myofibers are long lived cells. One of the early questions in the field which came from an interest in stimulating regeneration of diseased muscle was where are the myogenic stem cells in skeletal muscle tissue? The identification of the satellite cell appeared to resolve this issue. These cells which lie under the basal lamina (cell matrix) of a myotube with highly condensed nuclei reflecting a quiescent state were observed m enter the cell cycle and reform myotubes. While these cells were identified about 30 years ago. it has been only in the last 5 years that the molecular processes underlying their regulation have become identified. Work by Rudnicki and colleagues has shown that MyoD is critical for satellite cell function and functions downstream of a patterning gene called pax7. One obstacle faced by skeletal muscle is that these stem cells are relatively rare and do not always appropriately respond to injury or disease. Indeed, as muscle undergoes normal aging processes, muscle tissue atrophies and stem cell numbers and competence appears to decline leading to much of the loss of muscle mass typical of old age in humans. Rosenthal and colleagues have identified at least one growth factor, IGF, which can restore competence to stem cells and promote myotube hypertrophy and robust appearance-mice carrying a transgene for IGF appear to resist muscle aging without collateral ill effects. The potential for application to clinical situations is now within reach. Curiously, skeletal muscle tissue is not highly prone to cancers perhaps reflecting the myogenic program which entails terminal differentiation. The field of molecular cancer cell biology has focused heavily on the role of a critical tumor suppressor gene p53 and the retinoblastoma gene, pRb. One skeletal muscle tumor which is seen primarily in children, rhabdomyosarcomas, appears to result from a novel set of molecular pathways that involve these tumor suppressors but in unexpected ways, Thayer and colleagues outline how these pathways may operate in muscle which appear to diverge greatly from the general scenario that has emerged from the study of other tumor types. vii Advances in Developmental Biology and Biochemistry Ed. Paul M. Wassarman © 2 0 0 2 Elsevier Science. Printed in the Netherlands.

vii -- viii

vnl

When muscle differentiated in vivo, the muscle tissue expresses a variety of contractile proteins and catalytic enzymes that allow it to optimally perform its function. Specific muscles in the leg for instance, contain fast fibers which allow us to run with the drawback that they fatigue easily,. In contrast, back muscles are primarily slow which do not fatigue easily and allow us to maintain body posture for prolonged periods of time. While the contribution of nerve has long been recognized to regulate this process, it is also known that the initial pattern of fiber type specification can occur in the absence of nerve and may also reflect specific myogenic cell origin or lineages. Again, this issue has been under investigation for many years, but only recently have the signaling pathways involved become better understood. Schiaffino and colleagues outline recent progress in this field and their own work showing which pathways can regulate fiber type determination. While muscle cells can proliferate and differentiate in vitro, this process is largely manipulated by the amount of serum in the culture media. The serum which contains a mixture of growth factors or cytokines acts via specific receptors and ultimately engages components of the cell cycle and myogenic regulatory program to execute the biological outcome. It is obvious that changes in serum levels is an artificial construct of culture conditions that only vaguely reflects how differentiation is regulated in vivo. Whereas one can induce the majority of myogenic cells to terminally differentiate in vitro by lowering serum concentrations, we know that differentiation in vivo occurs in a non-synchronous fashion with fully differentiated myotubes immediately adjacent to highly proliferative myoblasts. The sheer number of cytokines implicated in this process is daunting, but a cohesive picture, as outlined by Olwin and colleagues is emerging with common effector genes downstream. Despite the intense focus on understanding skeletal myogenesis over the last several decades, investigators are often guided by certain dogmas and general ideas which influence how we interpret our data. We know, for instance, that muscle regeneration relies on the recruitment of satellite stern cells...or does it? We have defined a muscle stern cell as a cell which gives rise to skeletal muscle and does not contribute to other lineages. However work outlined by Cossu and colleagues has recently called these precepts into question. It appears that satellite cells can contribute to a wide variety of lineages and that other cell lineages such as cells originating from the bone marrow can participate in muscle regeneration. Again, the potential clinical application of this for stern cell based therapies is exciting and suggests a level of plasticity never imagined for this tissue. Thus, this volume focuses on these old and new directions for the skeletal muscle field and points out directions where the field may eventually progress. This volume is dedicated to the memory of Alex Mauro who first described the satellite cell and who spent innumerable hours with colleagues and convinced many graduate students, including me, that the field of skeletal myogenesis was one of the most fascinating biological questions to pursue. I am sure he would have something unexpected and provocative to say about where we have gone these last 10 years.

CHAPTER 1

THE MYOGENIC REGULATORY FACTORS

CLAIRE M. PALMER

a n d M I C H A E L A. R U D N I C K I

Table of contents I.

II. III.

IV,

V. VI.

VII.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. The MRFs are basic helix loop helix transcription factors . . . . . . . . . . . . . . . . . . . . .

2 2

B.

MRF activity is regulated at the level of dimer formation . . . . . . . . . . . . . . . . . . . . .

3

C.

MRFs bind E boxes in the promoters and enhancers of muscle specific genes . . . . .

4

D.

Identification of residues within the bHLH domains of the MRFs required for myogenic activity: The Myogenic Code . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

4

E.

Chromatin remodeling and the MRFs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

5

F

Myocyte enhancer factor-2 (MEF2) and myogenesis. . . . . . . . . . . . . . . . . . . . . . . . .

6

G.

Evolutionary relationship of the MRFs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

6

The role o f the MRFs in embryonic myogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

7

A.

7

Embryonic expression patterns of the MRFs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Mechanisms acting upstream of the MRFs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

8

A,

Pax3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

8

B.

Synergistic regulation of muscle development by Pax3. Dach2, Eya2 and S i x l . . .

9

C.

Development of the limb musculature: Role of Mox2 and Lbx 1. . . . . . . . . . . . . . .

10

D.

Axial tissues are required for epaxial somite myogenesis . . . . . . . . . . . . . . . . . . . . .

11

E. Shh and Wuts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gene targeting experiments reveal a hierarchical relationship amongst the MRFs . . . . .

12

A.

MyoD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

12

B.

Myf5 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

12

C.

MyoD and Myf5 are myogenic determination factors . . . . . . . . . . . . . . . . . . . . . . .

13

D.

Myogenin is essential for myoblast differentiation in vivo but not in vitro . . . . . . .

13

E.

MRF4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

16

F.

Compound MRF mutant mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

17

Postnatal myogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

18

A.

18

Satellite cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

The role of the MRFs satellite cell function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

18

A.

MRF expression pattern . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

18

B.

MyoD is required for satellite cell function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

20

C.

Myf5 and regeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

20

D.

Muscle stem cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

21

E.

Pax7 and satellite cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

VIII. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 Advances in Developmental Biology and Biochemistry © 2002 Elsevier Science. Printed in the Netherlands.

Ed. David Sassoon 1 -- 32

21 21 22

C. M. Palmer and M. A. Rudnicki

I.

In~oducfion

The first hints to the existence of a dominant-acting myogenic transcription factor come from studies in which non-muscle cells were fused to myoblasts and muscle specific gene expression was activated in the non-muscle nuclei of the heterocaryons (Blau et al., 1983; Wright, 1984). This transcription factor was later identified by a subtractive cDNA approach and called MyoD ((Davis et al., 1987); for review (Weintraub et al., 1991)). MyoD is a basic helix loop helix transcription factor capable of inducing myogenic gene expression in non-muscle cells ((Davis et al., 1987); for review (Weintraub et al., 1991)). Myf5, Myogenin and MRF4 were later identified as members of the MyoD family of transcription factors, also called the myogenic regulator factors (MRFs). A similar capacity to induce muscle specific gene expression in non-muscle cells was demonstrated for these MRFs. Members of this family have subsequently been identified in a diverse array of organisms including: Drosophila (reviewed in (Abmayr et al., 1998)); Xenopus (Harvey, 1990; Hopwood et al., 1991; Jennings, 1992); Chicken (FujisawaSehara et al., 1992), mouse (Davis et al., 1987; Wright et al., 1989) and human (Pearson-White, 1991). A.

The MRFs are basic helix loop helix transcription factors

The superfamily of basic helix loop helix (bHLH) transcriptional regulators has over 400 members which are active in a diverse array of developmental processes including myogenesis and neurogenesis (for review see (Atchley et al., 1997; Atchley et al., 1999; Morgenstern et al., 1999; Atchley et al., 2000; Wollenberg et al., 2000)). Based on homology within the bHLH domain, DNA binding affinity and the presence or absence of a leucine zipper motif, these proteins are classified into five subfamilies (Atchley et al., 1997). The MRFs are members of subfamily A, other members of this subfamily include Twist, dHand and E12 (Atchley et al., 1997). Subfamily A members bind the E Box sequence 5'- GAGCTG - 3' with high affinity, lack a leucine zipper motif and have a conserved configuration of amino acids at residues 5, 8, and 13 of the bHLH domain (Atchley et al., 1997). The four vertebrate MRFs share significant amino acid sequence identity (over 80%) throughout the basic helix loop helix (bHLH) domain (Murre et al., 1989b; Dias et al., 1994). The individual MRF bHLH domains appear to be functionally equivalent however regions outside this domain give specific activities to individual MRFs (Asakura et al., 1993). Through the amphipathic helices of the bHLH domain, the MRFs can both homo and hetrodimerize (Ferre-D'Amare et al., 1993; Ma et al., 1994). The MRFs form transcriptionally active heterodimers with the widely expressed E proteins, a distinct group of bHLH proteins including E12/E47, ITF-2 and HEB (Murre et al., 1989b; Lassar et al., 1991). There is evidence to suggest that DNA binding may facilitate the dimerization of MyoD with E protein partners (Weintraub et al., 1990; Czernik et al., 1996; Maleki et al., 1997; Wendt et al., 1998).

The Myogenic Regulatory Factors

B.

MRF activity is regulated at the level of dimer formation

Several proteins, for example Id, Mistl and MyoR. have been identified which either sequester E proteins from MyoD or prevent MyoD/E protein heterodimerization b y binding the bHLH domain of MyoD (Benezra et al.. 1990; Lemercier et al., 1998; Lu et al., 1999). ld (inhibitors of DNA binding) proteins are helix loop helix proteins which lack a DNA binding domain (Benezra et al., 1990; Christy et al., 1991: Biggs et al.. 1992: Ellmeier et al., 1992: Deed et al.. 1993; Riechmann et al., 1994). Idshave been suggested to avidly bind E proteins sequestering them from bHLH transcription factors (Benezra et al.. 1990; Langlands et al., 1997). In addition. Idl and Id2 bind to MyoD and Myf5 strongly preventing dimer formation (Langlands et at.. 1997). Id expression is down regulated during myoblast differentiation presumably allowing MRF activity to regulate differentiation (Benezra et al., 1990). Furthermore. ectopic expression of Id in differentiating cultures of C2C12 myoblasts inhibits terminal differentiation (Jen et al.. 1992). Ids may function to prevent MRF dependent entry into the differentiation program during expansion o f myogenic cells (Langlands et al., 1997). However, mice null for Idl, Id2. Id3. or Idl and Id3 do not have any overt muscle defects (Lyden et al.. 1999; Yokota et al., 1999; Rivera et al.. 2000; Yokota et al., 2000). The lack of a muscle phenotype may in part be due to functional redundancy between Id proteins: analysis of additional Id compound mutant mice may address this question. MyoR is a muscle restricted basic helix-loop-helix protein which antagonises MyoD (Lu et al., 1999). In the developing mouse, MyoR is expressed in a subset of muscle precursor cells predominantly during primary myogenesis between 10.5 and 16.5 days postcoitum (dpc). In cell culture, the expression o f MyoR is down regulated during differentiation. In addition to sequestering E proteins. MyoR/E12 heterodimers bind E boxes within the promoters of muscle specific genes competing with MRF/E protein heterodimer for access (Lu et al., 1999). MyoR may function to modulate the timing af muscle-specific gene expression and delay muscle fiber maturation (Lu et al.. 1999). Activators of myogenesis include proteins that stimulate the heterodimerization of MyoD and E proteins. In solution the formation of MvoD/E12 heterodimers is not favoured over the formation of MyoD homodimers (Maleki et al., 1997). However, M0s-dependent phosphorylation of MyoD promotes MyoD-E12 heterodimerization while inhibiting the ?.VfyoDhomodimer DNA binding activity (Lenom~and et al.. 1997 ). c-Mos is a serine/threonine ldnase expressed in aduk skeletal muscle (Leibovitch et al.. 1991). c-Mos directly phospliorylates serine 237 of MyoD (Pelpel K et al., 2000). Mutation of this serine resMu,e tO alanine abolishes the c-Mos positive regulation of MyoD (Pelpel K et al., 2000). Although in vitro data supports a role for c-Mos in :myogenic differentiation. homozygous c-Mos mutant mice do not have obvious skeletal muscle defects (Colledge et .al., 1394; tlashimoto et al., 1994; Lenormand et al., 1:997). Since the phoshorylation status of serine 237 was not determined in the c-Mos -/ mice, phosphorylation of this residue by yet4to-be identified kinases may compensate for the loss of c-Mos activity m muscle. In addition, the skeletal muscle defects in MyoD deficient mice are r~ot overtly apparent expect during muscle regeneration (Rudnicki et al., 1992; Megeney et al., t996). If c-Mos is essential for MyoD activity in vivo c-mos -/- mice e ~ b i t a muscle regeneration defect similar to mat of MyoD mutant animals.

4

C.


MRFs bind E boxes in the promoters and enhancers of muscle specific genes

Dimerization of bHLH proteins juxtaposes their basic domains forming a functional DNA binding domain (Ferre-D'Amare et al., 1993; Ma et al., 1994). In general, bHLH protein dimers bind to the consensus sequence 5'-CANNTG-3' (each basic region binds 1/2 of the consensus site), called an E box (Murre et al., 1989b). MyoD/E protein heterodimers preferentially bind the E box sequence 5'CA(G/C) (G/C)TG-3' (Blackwell et al., 1990). Highly conserved residues within the basic domain are responsible for consensus binding, In particular, a glutarnic acid residue conserved in the majority of bHLH proteins is in direct contact with the 'CA' nucleotides of the E box (reviewed ha (Robinson et al., 2000)). Residues within the basic domain in part dictate the consensus sequence specificity of different bHLH protein dimers. For example, Arg- 111 of MyoD makes contact with the 3'G of the E box. A similar contact does not occur when non-myogenic bHLH proteins bind DNA (reviewed in (Robinson et al., 2000)). The unique conformation of the MRF bHLH is likely due to the small size of Ala-114 (a residue required for myogenic activity). In non-myogenic bHLH proteins this position is occupied by a bulkier amino acid, it is this amino acid which is in contact with the G of CANNTG (Ferre-D'Amare et al., 1993; Ellenberger et al., 1994; Ma et al., 1994; Huang et al., 1998). D.

Identification of residues within the bHLH domains of the MRFs required for myogenic activity: The myogenic code

Mutagenesis studies have identified conserved residues within the basic region of the MRFs essential for their myogenic activity; residues Ala-ll4 and Thr-ll5 of MyoD and residues Ala-86 and Thr-87 of Myogenin (Davis et al., 1990; Brennan et al., 1991). These residues are referred to as the myogenic code. Replacement of Ala-114 of MyoD with asparagine or histidine, the residue normally found at that position in E proteins and Max respectively, alters the conformation of the basic domain such that Arg-111 does not make direct contact with the E box DNA (Huang et al., 1998). Furthermore this local conformational change prevents the unmasking of the N-terminal activation domain of MyoD upon DNA binding (Huang et al., 1998). Interestingly, myogenic specificity can be acquired by other bHLH proteins by the introduction of the myogenic code into the appropriate position (Weintraub et al., 1991; Davis et al., 1992). Similarly, the neurogenic bHLH protein MASH-1 acquires myogenic specificity by replacement of leucine-130 with lysine and the addition of one alpha helical turn without significantly altering cell free DNA binding properties (Dezan et al., 1999). Although the bHLH domains of the four MRFs are highly conserved, there is little sequence conservation outside this region, a notable exception is a cystein/histidine rich region (Tapscott et al., 1988; Weintraub et al., 1991). The transactivation domain is not conserved between individual MRFs. The transactivation domain of Myf5 is located in the C terminus while that of MyoD and MRF4 is in the N terminus (Braun et al., 1990; Weintraub et al., 1991; Mak et al., 1992). Strong transcriptional activation domains have been identified in both the N- and C-termini of myogenin (Olson, 1992). Additional functional studies are required to define more precisely the MRF transactivation domains.


E.

Chromatin remodeling and the MRFs

Activation of the myogenic differentiation program requires the alleviation of condensed chromatin structure. Both MyoD and Myf5 remodel chromatin in the regulatory regions of skeletal muscle specific genes (Gerber et al.. 1997). MyoD-dependem chromatin remodelling is mediated by two domains, a carboxy terminal domain and a cytseine/histidine rich domain located between the activation domain and the bHLH (Gerber et al., 1997). Interestingly, a number of previously identified MyoD binding partners (including COUP-TFII, N-CoR. p300 and PCAF) can modulate chromatin structure (Puri et al., 1997: Sartorelli et al., 1997; Bailey et al., 1998; Bailey et al.. 1999~ Sartorelli et al., 1999). The interaction of MyoD with binding partners that can modulate chromatin structure could provide a mechanism for targeting chromatin remodelling to muscle specific targets. The MyoD repressor COUP-TFII is an orphan nuclear receptor which recruits histone deacetylases via its interaction with N-CoR and other co-repressors (Bailey et al., 1998). It directly binds the N-terminal activation domain of MyoD. Interestingly MyoD also directly interacts with N-CoR through its bHLH domain (Bailey et al., 1999). In addition to recruiting histone deacetylases. COUP-TFII represses MyoD activity by competitively inhibiting the binding of p300 as a result this repression can be alleviated by overexpression of p300 fBailey et al., 1998). It and other orphan nuclear receptors including REV-erbc~ and RVR expression are down regulated during myogenic differentiation and inhibit MyoD-mediated transcription (see (Bailey et al.. 1998; Bailey et al., 1999)). These orphan nuclear receptors negatively regulate muscle differentiation and appear to have a role in maintaining the proliferative state of myoblasts (Dowries et aL, 1995; Muscat et al., 1995; Burke et al.. 1996: Bailey et al., 1998). Interestingly, MyoD recruits the histone acetyltransferase p300. and PCAF (Puri et al., 1997; Sartorelli et al., 1997; Lau et al., 1999). CBP/p300 and PCAF interact directly with MyoD and are required for MyoD-dependent transactivation (Yuan et al., 1996: Puri et al., 1997; Purl et ai.. 1997; Sartorelli et al.. 1997). A dominant negative p300 that is unable to bind MyoD inhibits both MyoD dependent myogenic conversion of 10T1/2 fibroblasts and transactivation by MyoD (Sartorelti et al., 1997), This is consistent with an essential role for endogenous p300 as a coactivator of MyoD. Both CBP/p300 and PCAF acetylate MyoD on two evolutionarily conserved lysine residues at the boundary of the 'bHLH domain (Sartorelli et ai.. 1999; Polesskaya et al., 2000). Acetylation of MyoD by either CBP/p300 or PCAF increases both in vitro DNA binding affinity and activity of muscle specific promoters (Sartorelli et al.. 1999: Polesskaya et al., 2000). Conservational substitutions of these lysine residues with nonacetylatable arginines inhibit MyoD activity (Sartorelli et al., 1999; Polesskaya et al., 2000). Recently an inhibitor of p300 histone acetylase activity, EID-1, was identified as a repressor of MyoD function (MacLellan et al., 2000). EID-1 is a 187 amino acid E1A like protein preferentially expressed in adult cardiac and skeletal muscle tissue (MacLellan et al., 2000). Skeletal muscle specific transcription is inhibited when EID-1 is overexpressed and appears to be due to its ability to bind and inhibit p300's histone acetylase activity (MacLellan et at., 2000).


F.

Myocyte enhancer factor-2 (MEF2) and myogenesis

The MEF2 family of transcription factors play an important role in skeletal, cardiac and smooth muscle development (for review see (Black et al., 1998)). Four vertebrate MEF2 genes have been identified (A to D) and appear to have arose from a common ancestor (for review see (Black et al., 1998)). MEF2 factors are members of the MADS-box family of transcriptional regulators. Members of this family share a 57 amino acid motif (MADS box)which provides minimal DNA-binding activity (Black et al., 1998). In addition to the MADS box, MEF2 factors contain a 29 amino acid MEF2 domain required for high affinity DNA binding and dimerization (Molkentin et al., 1996; Black et al., 1998). MEF2 proteins bind the consensus sequence 5' YTA (A/T)4 TAR 3' with varying degrees of affinity (Pollock et al., 1991; Yu et al., 1992; Molkentin et al., 1996). The transactivation domain of MEF2 factors is in the C-terminus and there is limited sequence homology in this region (for review see (Black et al., 1998)). In addition to it role as a MyoD co-factor, p300 also potentiates the transactivation by MEF2 family members (Sartorelli et al., 1997). In Drosophila, MEF2 expression is both necessary and sufficient to activate myogenesis (Bout et al., 1995; Lilly et al., 1995; Ranganayakulu et al., 1995; Lin et al., 1997). Both cardiac and skeletal muscle in Drosophila MEF2 mutant embryos fail to differentiate (Bout et al., 1995; Lilly et al., 1995; Ranganayakulu et al., 1995). However the myoblasts are appropriately specified and positioned suggesting MEF2 acts late in myogenesis to regulate terminal differentiation and fusion (Black et al., 1998). Interestingly MEF2 does not appear to be sufficient to activate the myogenic differentiation program in mammalian cells and is insufficient to induce myoblast differentiation in myogenin null mice (Yu et al., 1992; Hasty et al., 1993; Nabeshima et al., 1993; Molkentin et al., 1995; Ornatsky et al., 1997). MEF2 factors act synergistically with the MRFs to initiate myogenesis (Kaushal et al., 1994; Molkentin et al., 1995; Black et al., 1998). This synergy has been suggested to be mediated by direct MEF2/MRF interactions (Molkentin et al., 1995; Black et al., 1998). Because of functional redundancy between mouse MEF2 factors and cardiac insufficiency resulting in lethality, MEF2 knockout mice have been uninformative in regards to the in vivo role of MEF2 factors in skeletal myogenesis (Lin et al., 1997; Bi et al., 1999). However, an essential role for MEF2 factors is suggested by studies in which a dominant-negative form of MEF2A inhibits MyoD dependent conversion of 10T1/2 fibroblasts (Omatsky et al., 1997; Kolodziejczyk et al., 1999). G.

Evolutionary relationship of the MRFs

Unlike vertebrates, invertebrates such as C. elegans, sea urchins and Drosophila have a single myogenic regulatory factor (Krause et al., 1990; Michelson et al., 1990; Venuti et al., 1991). Despite little sequence homology outside the bHLH domain, the functional conservation between vertebrate and invertebrate myogenic factors is remarkable (Zhang et al., 1999). The Drosophilia MRF (nautilus) and chicken MyoD rescues the myogenic defect observed in C. elegans with a loss of function mutations in CeMyoD (hlh-1) (Zhang et al., 1999).

7


Phylogenic analysis suggests the four vertebrate MRFs arose from a single ancestral myogenic transcription factor (see Figure 1) (Atchley et al., 1994; Megeney et al., ~995). This factor initially gave rise to two subgroups by gene duplication. MyoD and Myf5 evolved from one subgroup and myogenin and MRF4 the other (Atchley et al.. 1994)o Interestingly, in some chordates a single gene encodes two differentially expressed MRFs suggesting there was a selective pressure for multiple MRFs prior to gene duplication (Meedel et al., 1997). Multiple MRFs likely act as specification and differentiation factors in multiple lineages allowing for coordinated muscle specific gene expression during embryogenesis (Meedel et al.. 1997).

M y o g e n i n (chr 1) MRF4 (chr 10) M y o D (chr 7) Myf5 (chr 10)

Figure 1. The phylogenetic relationship of the four vertebrate MRF genes. The vertebrates MRFs arose from a common ancestor by gene duplication prior to the radiation of the vertebrates.

II.

The role of the MRFs in embryonic myogenesis

A.

Embryonic expression patterns of the MRFs

In vertebrates, skeletal muscles with the exception of the anterior head. and extraocular muscles orginate from mesodermal precursor cells in the somites (for review see (Borycki et al.. 2000)). The paraxial head mesoderm gives rise to the anterior head and extraocular muscles (Wachtler et al.. 1992; Christ et al., 1995). Somites are epithelial spheres of paraxial mesoderm which flank the neural tube and form in a rostrocaudal progression (for review see (Christ et al., 1995)). As somites mature, they become subdivided into the ventral scelerotome, which contributes to the vertebra and fibs. and the dorsal dermomyotome (skeletal muscle, distal ribs and dorsal dermis) (for review see (Tajbakhsh el al., 2000)). The dorsomedial region of the dermamyotome gives rise


to epaxial muscles (axial muscles of the back). The limb or hypaxial muscles arise from cells which migrate into the limb bud from the ventrolateral dermamyotome (for review see (Molkentin et al., 1996)). Signals from the neural tube, notochord and paraxial mesoderm are involved in regulating somitic myogenesis (Cossu et al., 1996). The four vertebrate MRFs have distinct spatiotemporal expression patterns. MRF expression occurs initially in rostral somites and progresses caudal with somite maturation. Myf5 is the first MRF expressed in the developing embryo. Using in situ hybridization, Myf5 is first detected in the dorsomedial quadrant of the most rostal somite at day 8 postcoitum (pc.) (Ott et al., 1991; Tajbakhsh et al., 1996). As development progresses, Myf5 expression spreads to the whole myotome and is subsequently downregulated as the myotome matures. Myf5 expression in the developing limb bud is detected between day 10 pc. and 12 pc. MyoD mRNA first apprears in the hypaxial somitic domain at 9.75 dpc (Sassoon et al., 1989; Faerman et al., 1995). MyoD expression is detected throughout the myotome by day 11.0 and is maintained throughout development. MyoD expression in the limb bud occurs together with Myf5 expression. Myogenin mRNA is detected approximately 12hrs after Myf5 mRNA in the rostral somites. Myogenin expression spreads caudally as development progresses and is maintained throughout fetal life (Sassoon et al., 1989; Ott et al., 1991). Its expression is downregulated postnatally. Myogenin mRNA is detected in the developing limb bud after 10.5 dpc. MRF-4 is expressed transiently between 9.0 and 11.5 dpc in the myotome (rostral to caudal progression). It is subsequently re-expressed in the muscles of the embryo at 16 dpc becoming the most abundant MRF mRNA postnatally (Bober et al., 1991; Hinterberger et al., 1991; Hannon et al., 1992). Successive waves of myogenic determination and differentiation likely contribute to the complex developmental expression pattern of the MRFs.

III.

Mechanisms acting upstream of the MRFs

A.

Pax3

Members of the Pax (paired box) family of proteins are developmentally important transcription factors characterized by the presence of a paired domain (Reviewed in (Borycki et al., 1997; Rawts et al., 1997)). The paired domain is a 128 amino acid DNA binding motif, which is essential for Pax protein function (Treisman et al., 1991; Xu et al., 1995). In addition to the paired domain, several Pax proteins including Pax3 and Pax7 contain a second highly conserved motif, the homeodomain. Pax3 and Pax7 are highly related proteins with similar embryonic expression patterns and as a result may be in part functional redundant (Jostes et al., 1990; Goulding et al., 1991). Identification of both Pax3/FKHR and Pax7/FKHR translocations in rhabdomyosarcomas provided the initial evidence for a role of Pax3 and Pax7 in myogenesis (Galili et al., 1993; Shapiro et al., 1993; Davis et al., 1994). Pax3 expression is initially observed in the neural tube at 8 dpc (Goulding et al., 1991). By 8.5 dpc, Pax3 expression is observed in the neuroepithelium and the dermomyotome. As development progresses, Pax3 expression in the dermomyotome is further restricted. Pax3 is also expressed in myoblasts migrating to the limb buds.


Mice deficient in Pax3 (Splotch mice) die in utero before day 15 pc (reviewed in (Borycki et al., 1997)). The phenotypic presentation of Pax3 null mutations is suggestive of not only a role for Pax3 in neural development but also in myogenesis. The muscles derived from migratory myoblasts (including limb muscles, back muscle (latissimus dorsi), shoulder muscles and diaphragm) are not present in Splotch embryos (10 dpc) (TajbakJash el al.. 1997). The limb muscle deficiency observed in Splotch mice is partially: the result of a failure of somite cells in the lateral dermomyotome to migrate to the limb buds (Daston et al, 1996). A similar muscle phenotype is observed in c-met receptor tyrosine kinase mutant mice, consistent with the observation that c-met is a downstream target of Pax3 (Bladt et at., 1995). Although the Pax3 expressing migrating myogenic precursor cells are committed to the myogenic lineage, they do not differentiate until reaching the appropriate location within the limb. Recent work by Bendall and colleagues have provided some clues on how these lineage committed migrating cells postpone terminal differentiation (Bendall et al., 1999). Msxl. an homeodomain containing protein, sequesters Pax3 m migrating cells preventing it from activating transcription (Bendall et at.. 1999). In addition, Msxl represses MyoD transcription by directly binding a MyoD enhancer (Woloshin et al., 1995). Although Msxl has been disrupted in mice, muscle defects have not been reported (Satokata et al., 1994; Houzelstein et al., 1997). Tajbakhsh et al. (1997) generated 1nice lacking Myf5 and Pax3. Unlike homozygous Pax3 or Myf5 mutant embryos, double homozygous mutant embryos do not express MyoD and as a result body skeletal muscle is not present in these embryos. :In addition. ectopic expression of Pax3 can activate MyoD expression in a subset of cell types (Epstein et al., 1995; Maroto et at., 1997: Tajbakhsh et al., 1997). For example ectopic Pax3 can induce myogenesis in neural tube cells, however cannot induce myogenesis m 10T 1/2 fibroblasts and inhibits myogenic differentiation in C2C 12 myoblasts and 10T1/2 fibroblasts transfected with a MyoD expression plasmid (Epstein et al., 1995; Maroto e1 al.. 1997). Although Pax3 is normal expressed in the neural tube. its myogemc activity is suppressed by presumably a myogenic specific Pax3 inhibitor (RaMs et al., 1997; Maroto et al., 1997). The inability of Pax3 to induce myogenesis in other cell types may reflect either a lack of positive cofactors or the presence of Pax3 inhibitors these cells (Rawt.s et at.. 1997; Maroto et al., 1997), These findings are consistent with Pax3 acting as an upstream regulator of MyoD expression, with its ability to promote or inhibit myogenesis dependent on cell type, Similar to in vitro studies, the myogenic promoting activity of Pax3 is restricted in the embryo, suggesting that an inhibitor of Pax3 myogenic promoting activity is present in these tissues or cells (reviewed in (RaMs et al., 1997)). Alternatively an essential cofactor maybe absent from non-permissive tissues or cells (see Section 3.2 below). In addition, re~ent studies suggest Pax3 promotes expansion of the uncommitted somitic mesoderm by preYenting apoptosis of these cells (Borycki et al., 1999l, B.

Synergistic regulati0n of muscle development by Pax3, Dach2. Eya2 and Six1

The genetic hierarchy regulating myogenesls is strikingly similar to that which regulates both insect and mammalian eye development (for review (Relaix et al., 1999)).

10


During eye development, eyeless (Drosophilia) or the vertebrate homolog Pax6 participates in a positive regulatory feedback loop with dachshund (Dach) and eyes absent (Eya). Both dachshund and eyes absent synergize with the homeodomain containing protein sine oculis (So) to regulate gene expression (for review (Relaix et al.. 1999; Kawakami et al., 2000)). Ectopic expression of eyeless, eyes absent or dachshund results in ectopic eye development (for review (Relaix et al., 1999; Kawakami et al., 2000)). Heanue et al. (1999) recently demonstrated that an homologous genetic network regulates vertebrate myogenesis (Heanue et al.. 1999: Relaix et al., 1999). During myogenesis, Pax3 and Dach2 appear to participate in a positive regulatory feedback loop, similar to Pax6 and Dach during eye development (Heanue et al., 1999). Dach2 can synergizes with Eya2 to induce myogenesis. In addition, Eya2 can also synergize with Sixl (a vertebrate homolog of sine oculis) to induce myogenesis (Heanue et al., 1999). C.

Development of the limb musculature: Role of Mox2 and Lbx 1

Mox2 and the highly related Moxl are homeobox genes expressed in the somitic mesoderm. Moxl expression is detected prior to somite formation in the paraxial mesoderm (Candia et al., 1992; Candia et al., 1996). The development of limb musculature is perturbed in homozygous Mox2 mutant mice (Mankoo et al., 1999). These mice have a general reduction in limb musculature and are missing specific limb muscles (Mankoo et al., 1999). Unlike the Pax3 or c-met null mice, migration of myogenic precursors is normal in Mox2 deficient animals (Mankoo et al., 1999). However Mox2 is essential for the appropriate expression of both Pax3 and Myf5 in limb bud, it is not however required for appropriate MyoD expression (Mankoo et al., 1999). The mammalian homolog of the Drosophila homeobox protein ladybird, Lbxl, is essential for the lateral migration of muscle progenitor cells into the limb (Jagla et al., 1995; Brohmann et al., 2000; Gross et al., 2000). A complete absence or severe reduction of the lateral limb muscles is observed in Lbxl null mice (Schafer et al., 1999; Brohmann et al., 2000; Gross et al., 2000). Other muscles which require contributions from migratory progenitor cells including the ventral limb muscles, tongue muscle and diaphragm muscles are normal consistent with the notion that only a subset of muscle progenitors require Lbxl for appropriate migration (Schafer et al., 1999; Brohmann et al., 2000; Gross et al., 2000). Lbxl expression is detected by in situ hybridization in the lateral somite and dermomyotome at 8 dpc. By 10 dpc, Lbxl expressing migratory cells originating from the ventrolateral lip of the dermamyotome are detected, these cells are Pax3 and c-met positive (Uchiyama et al., 2000). Analysis of Lbxl expression in Splotch mice suggest Pax3 may act as an upstream regulator of Lbxl in the ventrolateral somite (Mennerich et al., 1998). Although Pax3 is required for Lbxl expression, it is not sufficient to induce Lbxl expression, signals derived from the limb bud are also necessary (Mennerich et al., 1998).


D.

11

Axial tissues are required for epaxial somite myogenesis

Cells within the newly formed somite are positionally determined as a result of extrinisic signals acting during later stages of somite development (Aoyama et al., 1988: Ordaht et al., 1992). Analysis of the phenotypes of notochord mutant mice and zebrafish suggest an essential role of the notochord in somite myogenesis (see Figure 3) (for review see (Borycki et al.. 2000)). These studies along with grafting and explant studies suggest a combination of signals from the neural tube, notochord and surface ectoderm are sufficient to induce expression of the MRFs and somitic myogenesis (for review see IArnold et al.. 1998: Borycki et al.. 2000)). In presomitic mesoderm explants from mouse 9:5 dpc embryos, axial structures preferentially activate myogenesis through a Myf5-dependent pathway, whereas axial structure isolated from a 10.5 dpc embryo preferentially induce MyoD expression independent of Myf5 (Tajbakhsh et al., 1998). This suggests the temporal separation of Myf5 dependent and independent pathways is the result of axial tissue maturation (Tajbakhsh et al., 1998). The positive acting factors include Sonic hedgehog, and Wnt family members (see below, for review see (Borycki et al.. 2000)). E.

Shh and Wnts

The notochord and floor plate secrete factors including somc hedgehog which positively regulate myotome formation (for review see (Borycki et al., 2000)). Sonic hedgehog (Shh) with it two related mammalian proteins Indian hedgehog and Desert hedgehog are small secreted signaling proteins. It is synthesized in the notochord, floorplate and zone of polarizing activity in the developing embryo (Marti et al.. 1995). Shh is required for early activation of Myf5 and MyoD in somite cells that give rise to the epaxial lineage (see Figure 3) (Borycki et al., 1999). It is however not essential for the activation of Myf5 and MyoD in the hypaxial dermomyotome or the myogenic progenitors that form limb muscle (Borycki et al.. 1999]. The downstream effectors of Shh signaling include Ptc I the Shh receptor) and the Gli transcription factors (for review see (Borycki et al., 200(3)). Although analysis of the expression patterns of these proteins suggest a possible role in control of myogenesls, the involvement of molecules downstream of Shh have yet to be determined. Similarly Wnt-1. Wnt-3 and Wnt-4, which are expressed in dorsal regions o f the neural t u ~ , can activate myogeneis m vitro (see Figure 3) (Munsterberg et al.. 19957. Analysis of Wntl +, Wnt3a 4- null mice suggests Wnt signals regulate early myogenesis. in particular early Myf5 expression (Ikeya et al.. 1998). Wnt7a is expressed in the dorsal ectoderm and preferentially activates MvoD (Tajbakhsh et al.. 1998). The myogenic promoting activity of Wnt7a is inhibited by BMP4. These results highlight the role of Shh and Wnt signalling in embryonic myogenesis. In addition° these results support the hypothesis that the epaxial lineage is Myf5-dependent and distinct from the MyoD-dependent hypaxial lineage.

12

IV.


Gene targeting experiments reveal a hierarchical relationship amongst the MRFs

The four MRFs have similar capacities to convert 10T1/2 fibroblasts into myoblasts. In addition, the DNA binding and dimerization properties are similar amongst the MRFs (Braun et al., 1991). Cross-activation of the indigenous MRF genes by ectopically expressed MRFs complicates analysis of individual MRF function (Braun et al., 1989; Thayer et al., 1989). Targeted disruption of the MRFs was essential to define the individual roles of:these proteins in myogenesis. A.

MyoD

Mice deficient in MyoD are viable, fertile and do not display any overt skeletal muscle abnormalities (Rudnicki et al., 1992). MyoD deficiency does not affect sacromere ultrastructure, fast/slow fiber ratio or muscle specific gene expression. Myogenin and MRF4 expression in MyoD mutant mice is unaffected. Both heterozygous and homozygous MyoD mutant mice however show significant postnatal overexpression of Myf5; 1.8 and 3.5 fold respectively (Rudnicki et al., 1992). While the epaxial muscles in a MyoD 4- embryo develop normally, development of the hypaxial musculature or migrating muscle lineage is delayed by about 3.5 days (Kablar et al., 1997; Kablar et al., 1998; Kablar et al., 1999; Kaul et al., 2000). In addition, adult MyoD deficient muscle have a severe regenerative defect (Megeney et al., 1996). The effect of MyoD deficiency on regeneration will be discussed later. B.

My f5

Similar to MyoD mutant mice, newborn Myf5-/- mice have no overt morphological defects in their skeletal muscle. Expression of the three remaining MRFs is not upregulated in Myf5 mutant mice (Braun et al., 1992). Using a Myf5 targeted nls-lacZ reporter gene, Tajbakhsh and colleagues tracked the migratory routes of Myf5 expressing muscle progenitor cells ([3-gal positive cells) i n both heterozygous and homozygous mutant embryos. Interestingly in Myf5 mutant embyos, muscle progenitor cells that have activated Myf5 expression ([3 -gal positive) migrate aberrantly and fail to respond to positional cues. These ceils ultimately adopt a non-myogenic fate (Tajbakhsh et al., 1996). Epaxial myogenesis in Myf5 mutant embryos is markedly delayed and is rescued by the induction of MyoD expression at 13.5 dpc (Braun et al., 1992; Kablar et al., 1997; Kablar et al., 1998; Kablar et al., 1999; Kaul et al., 2000). Hypaxial myogenesis is normal in Myf5 -/- embryos. Taken together the delay in epaxial and hypaxial myogenesis observed in Myf5 mutant and MyoD mutant mice respectively supports a multiple lineage hypothesis in which Myf5 regulates epaxial muscle formation and MyoD regulates hypaxial muscle formation (Kablar et al., 1997; Kablar et al., 1998; Kablar et al., 1999). Nevertheless, the lack of obvious skeletal muscle defect in Myf5 and MyoD mutant mice suggests that the distinct myogenic lineages can to some extent compensate for each other. This hypothesis is further supported by the finding that


13

Myf5 and MyoD axe not co-expressed in muscle precursor cells but rather expressed in a mutually exclusive manner (Braun et al., 1996). Several Myf5 mutant mice have been generated since the initial targeting) It was initiNly ~ocked-out by Braun et al. (1992) (Braun: et al., t992). ~ e s e Myf5 mutant mice die perinatally due to a severe rib defect. A similar rib defect was o b s e r v ~ when a nls-lacZ,reporter gene, together with the PGK-neo selection cassette, Was inserted into exon 1 of the Myf5 locus (Tajbakhsh et al., 1996). However subsequent targeted disruption of the Myf5 locus in which the PGK-neo-TK selection cassette was excised by Cre-recombinase do not have any skeletal defects and are both viable arid fertile (Kaul et al., 2000). C.

MyoD and Myf5 are myogenic determination factors

To investigate whether Myf5 and MyoD could functional compensate for one another in embryonic myogenesis, compound Myf5/MyoD mutant mice were generated (Rudnicki et al., 1993), (Kaul et al., 2000). Although born alive, Myf5/MyoD compound mutant mice are immobile and die shortly after birth. These mice are devoid of both myoblasts and skeletal myofibers. Adipose and amorphous loose connective tissue occupies the spaces normally occupied by muscle (Rudnicki et al., 1993). A sequential ablation of motor neurons from the spinal cord to the brain occurs during development in Myf5+: MyoD + mutant embryos, highlighting the intimate connection between nervous development and skeletal myogenesis (Kablar et al., 1999), The muscle phenotype of the compound mutant mice is consistent with a model in which MyoD and Myf5 acl as primary MRFs and are required for myogenic lineage determination (see Figure 2 for summary) (Rudnicki et al., 1993). Interestingly, Myf5+/-: MyoD + mice unlike Myf5+: MyoD +/- mice are not viable suggesting postnatal survival of MyoD mutants requires two functional Myf5 alleles (Rudnicki et al, 1993). These mice have a substantial reduction in muscle fiber number. Although these observations confirm that Myf5 and MyoD can functional substitute for each other during development, they suggest Myf5 and MyoD are not functional equivalent. Similarly myogenin targeted into the Myf5 locus can partially substitute for Myf5. Myf5 myg'k~lrnyg-ki.MyoD -/ mutant mice die perinatally because of reduced skeletal muscle mass (Wang et al.. 1997). The skeletal muscle formed in Myf5 myg'ki/myg-ki, MyoD + mutant mice is normal suggesting the observed reduction in skeletal muscle mass is the result of inefficient recruitment of precursors into the myogenic lineages (Wang et al., 1997). D.

Myogenin is essential for myoblast differentiation m vivo but not in vitro

Two independent groups generated myogenin mutant alleles by homologous recombination-mediated gene targeting (Hasty et al.. 1993; Nabeshima et al., 1993). Homozygous myogenin mutant mice die perinatally presumabl5 because of a severe defect in the diaphragm. A significant reduction in diaphragm thickness and skeletal muscle content is Observed in myogenin null mice (Hasty et al.. 1993: Nabeshima et al., 1993). Unlike the MyoD -- and Myf5 -- mice. myogenin mutant mice have major skeletal

14


Ceils in Somite

Primary MRFs

Myf5 ~ myogenic

Secondary

MRFs

~

Myogenin

myoblasts

precursors

Muscle

Dots.! S~bdoma~*l

~ myogenic precursors Ventral Subdomain

MyoD ~

~ myoblasts

Myogenin MRF4 ~" Muscle

Figure 2. Extracellular signals from the surrounding tissue regulate myotome formation. Positive signals including Sonic hedgehog mad Wnts from the axial structures induce expression of the primary MRFs in the myotome. MyoD expression in the ventral myotome is induced by Wnt7a secreted from the ectoderm and lateral plate. The anti-myogenic activity of BMP4 is inhibited by Noggin secreted from the dorsal medial lip

muscle abnormalities (Hasty et al., 1993; Nabeshima et al., 1993). A significant reduction in skeletal muscle mass is observed throughout the body. In addition, the skeletons of mygenin -/- mice show abnormal curvature of the spine and rib cage deformities. Subsequent analysis of an hypomorphic myogenin allele in which myogenin transcription is reduced to 25% wildtype, suggests distinct myogenin threshold levels are required for myogenesis and thoracic skeletal development (Vivian et al., 1999). Muscle hypoplasia is observed in both homozygous myogenin hypomophs and myogenin (hypo/null) mice. The severity of the hypoplasia is dependent on the level of myogenin expression. The thoracic skeletal defect observed in myogenin mutant mice appears to be due to delayed rib cartilage migration (Vivian et al., 1999). Histological analysis of skeletal muscle from myogenin null neonates and embryos revealed extensive disorganization of the tissue. There is a substantial reduction in myofibre density in homozygous mutant embryos. However the number of myoblasts appear normal suggesting myogenin is not required for lineage commitment but rather terminal differentiation. Consistent with this myogenin expression is upregulated during myoblast differentiation both in culture and in vivo (Sassoon et al, 1989; Wright et al., 1989; Smith et al., 1994). The generation of myogenin / Myf5 and myogenin/MyoD mutant mice demonstrates the function of myogenin does not overlap with either MyoD or Myf5 (Rawls et al., 1995). These results together suggest myogenin is a secondary MRF acting downstream of the primary MRFs, MyoD and Myf5.

The Myogenic Regulatory Factors'

15

Figure 3. Schematicrepresentation of the hierarchial genetic relationships of the MRFs in the epaxial and hypaxial myotOmes.Cells from the dorsal and ventral subdomains differentiate into myoblastsby the expression of Myf5and MyoDrespectively. Subsequentlybothprimary MRFs are co-expressedin these cells. Gene targeting experiments demonstrate that Myf5 or MyoDare required for myogeniclineage determination while myogeninand MRF4 are necessaryfor terminal differentiation.

The muscle defect in myogenin mutant mice is most severe in muscle arising from the ventral subdomain of the dermomyotome (hypaxial) (Nabeshima et al.. t993). The hypaxial dermomyotome also gives rise to a migratory population of myogenic precursors. Very few. if any myofibres are present in these muscles. MyoD and myogenin are the predominate MRFs expressed in this region of the dermamyotome before and after myoblast differentiation respectively (Smith et al.. 1994). This is consistent with the hypothesis that myogenin is the predominate differentiation factor of a MyoD-dependent lineage. Myofibres are present in axial, intercostal and back muscles however the majority are disorganized and lack Z lines (Nabeshima et al.. 1993). Studies by Venuti and colleagues (1995) demonstrate that these myofibres arise during primary myogenesis (Venuti et al.. 1995). Suggesting primary myogenesis is unaffected by the absence of myogenin. The levels of MRF4 transcript are markedly reduced in myogenin mutant embryos while the MyoD levels are unaffected (Hasty et al., 1993: Nabeshima et al., 1993). This reduction in MRF4 could reflect a lack of differentiated myofibres as opposed to direct regulation of MRF4 by myogenin. MRF4 can partially substitute for myogenin during embryonic myogenesis (Zhu et al.. 1997). Myogenin does not appear have an essential role in myoblast differentiation in vitro, Continuous cell lines of myoblasts isolated from myogenin mutant embryos differentiate normally in culture. I n addition, myogenin -- fibroblasts convert to myotubes by ectopic

16


expression of MyoD (Nabeshima et al., 1993). However. analysis of primary MyoD 4myogenic cells suggest continuous culture of primary myoblasts effects differentiation potential. The differentiation defect observed in low-passage primary MyoD + myoblasts is not observed in later-passage MyoD + cultures (Sabourin et al., 1999). In fight of these results, it is important to characterize the differentiation potential of low-passage primary myogenin + myoblasts. E.

MRF4

The gene encoding mouse MRF4 is linked to the Myf5 gene in a head to tail orientation. Approximately 8.5 kb of sequence separates the two loci (Miner et at., 1990). Three laboratories have generated MRF4 mutant mice (Braun et at., 1995; Patapoutian et aL, 1995; Zhang et al., 1995). Although the identical PGK-neo selection cassettes was used in the targeting constructs, orientation of the cassette and deleted MRF4 sequence is unique t o each construct [for review see (Olson et at., 1996)]. Analysis of the role of MRF4 in embryonic myogenesis has been complicated by the close proximity of the two MRF genes. Targeted disruption of MRF4 affects Myf5 expression via a cis-acting mechanism tFloss et at., 1996; Yoon et al., 1997). The negative cis effect of MRF4 mutations on Myf5 transcription was confirmed by the generation of compound heterozygous MRF4; Myf5 rmce in which the disrupted alleles are on different chromosomes. These mice have a significant reduction in Myf5 transcript compared to control animals suggesting the wildtype Myf5 allele is partially inactivated (Floss et at., 1996; Olson et at., 1996: Yoon et at., 1997). Myf5 enhancer elements have been identified in a 14 kb region spanning the MRF4/Myf5 locus (Summerbell et al., 2000). These enhancers regulate Myf5 expression in epaxial muscle precursor cells, the branchial arches and the central nervous system. MRF4 null mice generated in Barbara Wold's laboratory die perinatally and show defects in axial myogenesis and rib structure (Patapoutian et at., 1995). In light of recent insights into the neo cassette-dependent rib defects observed in some Myf5 mutant mice, the rib anomalies observed in the MRF4 null mice may be the result of cis effects on a yet to be identified gene (Kaul et al., 2000). A deficit in myotome development is observed in the MRF4 -/- mice between 9 and 11 dpc. This corresponds to the first wave of MRF4 expression during development. A significant reduction in Myf5, MyoD and myogenin expression is observed in mutant mice at day 10 pc. A decrease in Myf5, MyoD and myogenin levels is also observed in heterozygous mice at this time (Patapoutian et al., 1995). By 14 pc, myogenesis in these MRF4 mutant animals appears normal. Minor defects are observed in the intercostal muscle of neonates (Patapoutian et al., 1995). Although studies using transgenic mice have shown the MRF4 sequences deleted in the Wold mice are not required for appropriate Myf5 expression, interference of Myf5 expression by PGK-neo can not be ruled out (Summerbell et al., 2000). The Olson MRF4 mutant mice are viable however have malformations of the ribs including bifurcations, fusions and supernumerary processes (Zhang et al., 1995). Neonatal muscle is grossly normal although the level of embryonic myosin heavy chain is slightly decreased. Similar to the Wold MRF4 -/- mice, defects in myotome differentiation are observed in 10.5 dpc MRF4 -/- embryos (Vivian et at., 2000). Myogenin and


17

Myf5 expression in the developing myotome are severely reduced indicative of retarded myotome differentiation. There is a significant increase in postnatal myogenin levels in MRF4 mutant mice. This is consistent with the notion that MRF¢ is required to downregulate postnatal myogenin expression (Zhang et al.. 1995). Alternatively, increased myogenin expression may compensate for the loss of MRF4 (Zhang et al.. I995). The sequence deleted in the Olson MRF4 allele is more extensive than either the Arnold or Wold alleles (for review see (Olson et al.. 1996)). In addition, the PGK~neo cassette ~s inserted in the opposite orientation (opposite to Myf5 transcription), The enhancer element that regulates Myf5 epaxial somite expression is deleted in the Olson al!ele (Summerbell et al.. 2000). The reduction in Myf5 expression :in the epaxial somite may contribute to the delay in myotome differentiation observed in these MRF4 nnll mice. MRF4 mutant mice generated in the laboratory of Hans Arnold die perinatally of respiratory distress due to severe malformation of the ribs (Vivian et al., 2000). Myf5 expression is disrupted in this mutant possibly because of interference by the PGK-neo cassette (Braun et al., 1995: Summerbell et al., 2000). As a result, these mice are phenotypic representatives of Mvf5+: MRF4 + double knock-out mice. These mice are essentially a phenocopy of the Myf5 mutant mice with a delay in early myotome formation (Braun et al., 1995). The deep axial muscles are reduced in the Arnold MRF4 hornozygous knock-out mouse. Consistent with the hypothesis that Myf5 and MRF4 are important in regulating the development of muscles arising from the myotome (Braun et al., 1995). F.

Compound MRF mutant mice

Compound MyoD-/-:MRF44-tOlson allele) and MyoD+:Myogenin -/- mice have been generated (Rawls e~ at.. 1998). The muscle phenotype of a MRF4/ myogenin double mutant mouse is no more severe than the myogenin single mutant (Rawls et al., 1998). The number of residual myofibres is comparable in mice lacking myogenin or myogenin/MRF4 suggesting neither myogenin nor MRF4 is required for primary myofibre differentiation. Interestingly continuous lines of myoblasts isolated from MRF4/myogenin double mutant mice differentiate normally in vitro, highlighting the differences between in vivo and in vitro myogenesis. The MRF4-/-:MyoD-/- mouse is a phenocopy of the myogenin mutant mouse with only residual myofibres and undifferientated myoblasts (Rawls et al.. 1998). This is consistent with the notion that terminal differentiation of myobtasts originating from the MyoD~dependent hypaxia~ lineage requires myogenm while terminal differentiation of myoblasts originating fi'om the Myf5-dependent epaxial lineage requires MRF4. Myoblasts isolated from MyoD/MRF4 double mutant neonates differentiate normally in vitro suggesting the in vivo cellular environment may restrict the compensatory capacity of the remaining MRFs {Valdez et al.0 2000). Mice lacking all of the MRFs except Myf5 have been generated (Valdez et al.. 2000). Triple mutant myoblast fail to fuse both in vitro and in vivo. Unlike the myogenin or MRF4/MyoD mutant mice, these mice do not have residual myofibres. The observed differentiation defect in the triple mutant mice was suggested to reflect a failure to meet sufficient threshold levels of myogenic bHLH factors (Valdez et al.. 2000), (Rawls et al.. 1998). However. these results are consistent

18


with the notion that individual myogenic bHLH proteins have evolved specialized functions and distinct MRFs may be required m switch on the expression of different muscle specific genes (Valdez et al., 2000). For example MyoD and Myf5 unlike myogenm, have domains which mediate chromatin remodelling, allowing for efficient activation of genes in regions of transcriptionally silent chromatin (Gerber et al.. 1997). Although a threshold level of MRF may be required for both in vitro and in vivo myogenesis, MRF embryonic expression patterns and the phenotypes of both single and compound mutant mice are consistent with a distinct myogenic lineage hypothesis. Targeting experiments demonstrate the hierarchial relationship between the MRFs. MyoD and Myf5 are primary MRFs required for the conversion of pre-myogenic cells into skeletal myoblasts. Myogenin and MRF4 are secondarv MRFs required for myoblast differentiation and myotube fusion. In addition, these experiments support the notion that MyoD/myogenin and Myf5/MRF4 regulate the hypaxial and epaxial myogenic lineages respectively and that these lineages can to some extent compensate for each other (see Figure 2).

V.

Postnatal myogenesis

A.

Satellite cells

In postnatal skeletal muscle a distinct lineage of myogenic progenitor cells; satellite cells, is responsible for growth, maintenance and repair of the muscle tissue (for review see (Seale et al., 2000)). Satellite cells are intimately juxtaposed to the plasmalemma (sarcolemma) of mature myofibres such that a single basal lamina surrounds both the myofibre and the satellite cell. The nuclei of satellite cells are oval and are heterochromatic in comparison to myonuclei (reviewed in (Bischoff, 1994)). Satellite cells contain little cytoplasm however have abundant plasmalemmal vesicles (reviewed in (Bischoff, 1994)). In neonatal rodents, approximately 30% of muscle nuclei represent satellite cells (Bischoff, 1994). The number of satellite cells decreases with ageing to less than 5% in adult muscle (Gibson et al., 1983; Bischoff, 1994; Grounds, 1998). Normally mitotically quiescent, satellite cells are induced to proliferate in response to muscle damage and various other stresses [for review see (Seale et al., 2000). After multiple rounds of division, t h e progeny of activated satellite cells (muscle precursor cells; mpcs) fuse to the damaged myofibres or form new fibers. Satellite cells and their daughter mpcs are distinct cell populations as defined by biological and biochemical criteria (Bischoff, 1994).

VI.

The role of the MRFs satellite cell function

A.

MRF expression pattern

The expression of the MRFs in quiescent satellite ceils and during satellite cell activation, proliferation and differentiation has been assayed by both RT-PCR and

19


transgene analysis (Smith et al., 1994; Yablonka-Reuveni et al.. 1994: Cornelison et al.. 1997: Creuzet et al., 1998; Cooper et al.. 1999). MRF expression is not detected in quiescent satellite cells by RT-PCR (Smith et al.. 1994; Yabtonka-Reuveni et al.. 1994: Cornelison et al., 1997). However Myf5(nlacZ) has been reported to be expressed in CD34 positive quiescent satellite cells (Beauchamp et al., 2000). Upon activation satellite cells quickly upregulate either MyoD or Myf5 expression prior to proliferation (Smith et al., 1994: Cornelison et al., 1997; Cooper et al.. 1999). Subsequently. mosl proliferating mpcs express both MyoD and Myf5 (Cornelison et al.. 1997: Cooper et al., 1999). Myogenin and MRF4 are the last MRFs expressed coincidental with MPC differentiation and fusion (Smith et al., 1994; Yablonka-Reuveni et al.. 1994: Cooper et al.. 1999). The MRF expression pattern dm'ing satellite cell activation. proliferation and differentiation is analogous to the pattern observed during embryonic myogenesis (see Figure 4). It is intereseting to speculate that MyoD and Myf5 may regulate distinct Satellite cell populations similar to the epaxial and hypaxial myogenic progenitors during embryogenesls.

r Myf5

MPC expansion MyoD ] Myi5

terminal differentiation Myogenin]MRF~

fu

repair damage fiber

Figure 4.

new fiber

A schematic representation of satellite cell dependent muscle repair. Upon activation of satellite

cells by muscle damage either MyoD or Myf5 expression is upregulated prior to proliferating. Subsequent!y, most proliferating satellite cells (myogenic precursor ceils, MPCs ~express both primary MRFs. Expression of the secondary MRFs is coincidental with MPC differentiation and fusion.

20


B.

MyoD is required for satellite cell function

Insights into the role of MyoD in satellite cell function came from studies in which MyoD mutant mice were interbred with mdx mice (Megeney et al., 1999). mdx mice have a loss of function point mutation in the dystrophin gene and are a model for human Duchenne and Becker muscular dystrophies (Bulfield et al., 1984; Sicinski et al., 1989). Although extensive muscle fiber necrosis is observed in young mdx mice, skeletal muscle integrity is maintained because of a high regenerative capacity of the tissue. The muscle of m d x mice is significantly hypertrophic with a predominant number of fibers containing centrally located nuclei (Anderson et al., 1987; Carnwath et al., 1987; Coulton et al., 1988). Centrally located nuclei are typical of repaired fibers (for review see (Grounds et al., 1993)). The severity of myopathic changes in the skeletal muscle of mdx: M y o D 4- mice are profound in comparison to those observed in mdx mice and result in premature death. Compound mutant mice develop a severe dorsal-ventral curvature of the spine and have an abnormal waddling g a i t (Megeney et al., 1996). The muscle of mdx: M y o D -i- mice, unlike that of mdx mice is not hypertrophic suggesting the loss of MyoD reduces the regenerative capacity of the tissue. This was tested directly by injury-induced regeneration experiments in MyoD null mice. MyoD-deficient muscle does not regenerate efficiently despite an increase in the number of satellite cells suggesting MyoD is required for progression through the differentiation program and in the absence of MyoD satellite cells undergo selfrenewal (Megeney et al., 1996). A similar differentiation defect is observed in both primary MyoD 4- myoblasts and single-fiber culture (Sabourin et al., 1999; Yablonka-Reuveni et al., 1999; Cornelison et al., 2000). Consistent with chronic muscle regeneration in vivo, fibers isolated from MyoD 4- muscle are abnormally branched (Cornelison et al., 2000). Although the majority of MyoD -/- satellite cells in fiber culture entered the cell cycle and upregulated Myf5 expression, major defects in both myogenic gene expression and differentiation are observed (Cornelison et al., 2000). In particular MyoD -/- satellite cells in single fiber culture fail to upregulate MRF4 expression and have reduced levels of m-caderin (Cornelison et al., 2000). These results suggest primary MyoD -/- mpcs represent a normally transient intermediate between quiescent satellite cells and activated mpcs. These cells provide an unique opportunity to study the early stages of satellite cell activation (Sabourin et al., 1999). C.

Myf5 and regeneration

The role of Myf5 in muscle regeneration has yet to be established. However the propensity of MyoD negative Myf5 expressing MPCs for self-renewal suggests Myf5 may be involved in maintaining the satellite cell pool in adult muscle (Sabourin et al., 1999; Yablonka-Reuveni et al., 1999). Although inefficient, muscle regeneration does occur in the absence of MyoD (Megeney et al., 1996). In addition, 20% of satellite cells express Myf5 and not MyoD 3 hours post injury suggesting Myf5 expression is sufficient to activate the satellite cell developmental program (Cooper et al., 1999). Myf5 dependent regeneration may account for the regeneration observed in mdx; M y o D -/- mice.


2I

With the recent generation of viable Myf5 null mice. the role of Myf5 in muscle repair will likely be elucidated in the near future. D,

Muscle stem cells

Stem cells isolated from various adult tissues have recently been shown t o have myogenic potential (for review see (Seale et al., 2000)). The initial studies demonstrated that bone marrow derived stem cells could acquire myogenic specificity and participate in the repair of chemically induced muscle damage in immunodeficient mice (Ferrari et al., 1998). A similar capacity of bone marrow derived stem cells to contribute to both cardiac and skeletal muscle was later demonstrated using the mdx mouse model of Duchenne and Becker muscular dystrophy (Bittner et al.. 1999). Hematopoietic stem cells can by purified from bone marrow by fluorescence activated cell sorting (FACS) using Hoechst 33342 dye (Goodell et al., 1996; Goodelt et al., 1997). Similar to bone marrow, the purified stem cells can participate in muscle repair (Gussoni et al., 1999). Using the Hoechst/FACS method, Gussoni et al. later demonstrated that a similar adult stem cell potential could be isolated directly from muscle tissue and that these cells can both contribute to regenerating myofibers and reconstitute the bone marrow compartment of lethally irradiated mice (Gussoni et al.. 1999). Unlike hematopoietic stem cells, muscle derived stem cells appear to give rise to satellite cells. E.

Pax7 and satellite cells

Pax7 is a paired homeobox transcription factor essential for satel!ite cell development (Seale et al.i 2000)~ Pax7 mutant mice are devoid of satellite cells (Seale et al., 2000). Interestingly differential expression of alternately spliced Pax7 transcripts correlates with muscle regenerative efficiency indifferent strains of mice (Kay et al., i995; Kay et al:, t998). The specific requirement of Pax7 in satellite cell ontogeny is analagous to the essential function of a highly related family member, Pax3, in somitic myogenesis. Analysis of Pax7 + mice demonstrates that satellite cells and muscle derived stem cells are distinct celt populations (Seale et al., 2000). Interestingly, stem cells derived from the muscle of: Pax7 -/ mice have an increased capacity produce hematopoietic colonies in Culture suggesting muscle derived stem cells are upstream of satellite cells and via a Pax7 dependent pathway give rise to satellite cells (Scale et al., 2000).

VH.

Acknowledgements

We thank Kay Palmer, Patrick Seale and Mark Gillespie f o r helpful discussions and for careful reading of the manuscript. The work from the laboratory o f M.A.R. was supported by grants from the National Institutes of Health, the Canadian Institutes of Heatth Research, the Muscular Dystrophy Association, the Human Frontier's ScienCe Program, and the Canada Research Chmr Program, C.M.P. is a recipient of an OGSST studentship. M.A,R. holds the Canada Research Chair in Molecular Genetics and is a member of the Canadian Genetic Disease Network.

22

VIII.


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Weintraub, H., Davis, R., Tapscott, S., Thayer, M., Krause, M., Benezra, R., Blackwell, T. K., Turner, D., Rupp, R., Hollenberg, S., and et al. (1991). The myoD gene family: nodal point during specification of the muscle cell lineage. Science 251, 761-6. Weintraub, H., Dwarki, V. J., Verma, I., Davis, R., Hollenberg, S., Snider, L., Lassar, A., and Tapscott, S. J. (1991). Muscle-specific transcriptional activation by MyoD. Genes Dev 5, 1377-86. Wendt, H., Thomas, R. M., and Ellenberger, T. (1998). DNA-mediated folding and assembly of MyoD-FA7 heterodimers. J Biol Chem 273, 5735-43. WoUenberg, K. R., and Atchley, W. R. (2000). Separation of phylogenetic and functional associations in biological sequences by using the parametric bootstrap. Proc Natl Acad Sci USA 97, 3288-91. Woloshin, P., Song, K., Degnin, C., Killary, A. M., Goldhamer, D. J., Sassoon, D. A., and Thayer, M. J. (1995). MSX1 inhibits myoD expression in fibroblast x 10T1/2 cell hybrids. Cell 82, 611-20. Wright, W. E. (1984). Expression of differentiated functions in heterokaryons between skeletal myocytes, adrenal cells, fibroblasts and glial ceils. Exp Cell Res 151, 55-69. Wright, W. E., Sassoon, D. A., and Lin, V. K. (1989). Myogenin, a factor regulating myogenesis, has a domain homologous to MyoD. Cell 56, 607-17. Xu, W., Rould, M. A., Jun, S., Desplan, C., and Pabo, C. O. (1995). Crystal structure of a paired domalnDNA complex at 2. 5 A resolution reveals structural basis for Pax developmental mutations. Cell 80, 639-50. Yablonka-Reuveni, Z., and Rivera, A. J. (1994). Temporal expression of regulatory and structural muscle proteins during myogenesis of satellite ceils on isolated adult rat fibers. Dev Biol 164, 588-603. Yablonka-Reuveni, Z., Rudnicki, M. A., Rivera, A. J., Primig, M., Anderson, J. E., and Natanson, P. (1999). The transition from proliferation to differentiation is delayed in satellite ceils from mice lacking MyoD. Dev Biol 210, 440-55. Yokota, Y., Mansouri, A., Moil, S., Sugawara, S., Adachi, S., Nishikawa, S., and Gruss, P. (1999). Development of peripheral lymphoid organs and natural killer cells depends on the helix-loop-helix inhibitor Id2. Nature 397,702-6. Yokota, Y., Mori, S., Nishikawa, S. I., Mansouri, A., Gruss, P., Kusnnoki, T., Katakai, T., and Sbimizu, A. (2000). The helix-loop-helix inhibitor Id2 and cell differentiation control [In Process Citation]. Curr Top Microbiol Immunol 251, 35-41. Yoon, J. K., Olson, E. N., Arnold, H. H., and Wold, B. J. (1997). Different MRF4 knockout alleles differentially disrupt Myf-5 expression: cis-regulatory interactions at the MRF4/Myf-5 locus. Dev Biol 188,349-62. Yu, Y. T., Breithart, R. E., Smoot, L. B., Lee, Y., Mahdavi, V., and Nadal-Ginard, B. (1992). Human myocytespecific enhancer factor 2 comprises a group of tissue-restricted MADS box transcription factors. Genes Dev 6, 1783-98. Yuan, W., Condorelli, G., Caruso, M., Felsani, A., and Giordano, A. (1996). Human p300 protein is a coactivator for the transcription factor MyoD. J Biol Chem 271, 9009-13. Zhang, J. M., Chert, L., Kranse, M., Fire, A., and Paterson, B. M. (1999). Evolutionary conservation of MyoD function and differential utilization of E proteins. Dev Biol 208,465-72. Zhang, W., Behringer, R. R., and Olson, E. N. (1995). Inactivation of the myogenic bHLH gene MRF4 results in up-regulation of myogenin and rib anomalies. Genes Dev 9, 1388-99. Zhu, Z., and Miller, J. B. (1997). MRF4 can substitute for myogenin during early stages of myogenesis. Dev Dyn 209, 233-41.

CHAPTER 2

MYOFIBER SPECIFICATION AND SURVIVAL

A N T O N I O M U S A R O and N A D I A R O S E N T H A L

Table of contents I. II. III.

IV.

V. VI.

I.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Myofiber specification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The signal transduction pathways of muscle survival and plasticity . . . . . . . . . . . . . . . . A. Muscle survival factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. The calcium pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C, The PI3-kinase pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. MAP-Kinase pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The proteolytic systems in skeletal muscle tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. The calpain pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. The ubiquitiu-proteasome pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Therapeutic strategies to attenuate muscle wasting and promote myofiber survival . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

33 33 36 37 40 42 44 45 45 46 47 48

Introduction

Skeletal m u s c l e development is characterized by an orderly progression of molecular signals leading to generation of heterogeneous muscle fibers (Figure 1). Adult skeletal muscle retains a considerable degree o f plasticity, which makes the muscle capable of adopting specific phenotyplc properties in response to a wide range o f factors including physical activity, injury, stimulation by motor neurons, oxygen and nutrient supply, and changes in hormone levels. In this context, fiber type is an essential determinant of muscle function and alteration in fiber composition represents a major component in the muscle degeneration associated with muscle diseases.

II.

Myofiber specification

The emergence o f adult fiber types is a relatively 1ate process in m a m m a l i a n myogenesis. extending into the first weeks o f postnatal life. Adult skeletal muscle fibers are classified according to their relative velocities o f contraction (Type I or II, respectively). Slow (Type I) and fast (Type II) fiber types are named according to the unique myosin 33 Advanees in Developmental Biology and Biochemistry Ed. David Sassoon 33 © 2002 Elsevier Science. Printed in the Netherlands.

52

34

A. Musarb and N. Rosenthal

heavy chain (MyHC) isoform they express. Mammalian Type II myofibers are further classified into three major subdivisions, Type IIA, IIX and IIB, in order of increasing rates of contraction. In addition to myosin heavy chain, each fiber type expresses distinct proteins comprising the contractile apparatus, calcium-sequestering components, and metabolic enzymes, so that relatively skeletal muscle-specific proteins, such as actin, are expressed in all myofibers. In mammals, primary myofibers form during the early fetal period and rapidly develop a slow phenotype. A second wave of myogenesis occurs later in fetal development, with the primary fibers acting as a scaffolding for secondary myofiber formation (Figure 1). The secondary fibers contribute the vast bulk of the growing musculature that matures to the various fast phenotypes during the early postnatal period.

Figure 1. The origin of myoblasts during embryogenesis (Modified from Miller et al., 1993).

Adult mammalian muscles contain mixtures of the four different types of fast and slow myofibers, each muscle having a characteristic distribution and spatial organization. Developing myofibers initially express a special set of contractile protein isoforms. As individual cells mature, their patterns of muscle-specific gene expression become coordinated (Wade et al., 1990; Lyons et al., 1990; Ontell et al., 1993; Suthedand et al., 1993). Thus the proteins used in the initial assembly of the contractile apparatus are replaced to confer specialized properties to each individual fiber type in the adult (Schiaffino and Reggiani, 1996; Pette, 1998). The signals responsible for initiation and maintenance of fiber type-specific gene expression and the organization of the fibers into a muscle are only partially understood.

Myofiber Specification and Survival

35

The restricted expression of fiber-specific isoforms that gives rise to the phenotypic differences between muscle cells is generally regulated at the level of transcription. Unlike the successful determination of the precise cis-regulatory elements and transacting factors that direct muscle-specific gene expression (Buonanno and Rosenthal, 1996: Buonanno and Fields. 1999), the molecular dissection of fiber specification has been less tractable, partly because differentiated muscle cell cultures do not form mature fiber types. In addition, muscle-specific transgenes are rarely recapitulate the fiberspecific expression pattern of the endogenous gene presumably because the complete set of the fiber-specific modular regulatory elements have not been included. Potential mediators of fiber-specific transcription include the myogenic determination factors (MDFs), muscle-restricted members of the large superfamily of basic-helixloop-helix (bHLH) transcription factors. Although MyoD transcript preferentially accumulates in muscles largely composed of fast fibers (Hughes et al., 1993). little fiber-specificity has been demonstrated for the remaining three MDFs. Moreover, null mutations in the MyoD gene caused only relatively minor perturbations in fiber type gene expression (Hughes et al., 1997; Rudnicki and Jaenisch 1995), and mice carrying null mutations in the other three MDFs (myogenin myf5 artd MRF4) either failed to form differentiated muscle or showed no obvious fiber phenotype (Arnold and Braun. 1996). MDFs form heterodimers with E proteins, a related bHLH family of transcription factors (Lassar et al.. 1991). Although important in B cell development (Barndt and Zhuang, 1999), the E proteins may potentially provide a target for signaling pathways

36

IlL


The signal transduction pathways of muscle survival and plasticity

While much has been learned about skeletal muscle formation in the embryo, less is known about the molecular pathways controlling skeletal myocyte survival and myofiber plasticity in the adult. Tissue remodeling is an important physiological process which allows skeletal muscle to respond to environmental demands. This specific characteristic allows muscle fibers to undergo adaptive changes, such as changes in cytoarchitecmre and protein composition, in response to a variety of stimuli. Extracellular agonists, receptors, protein kinases, intermediate molecules and transcription factors are all components of one or more signal transduction pathways promoting specific cellular responses in adult myocytes (Figure 2). Chronic protein degradation is one of the most devastating consequences of defects in muscle survival mechanisms. For example, one of the most severe characteristics of muscular dystrophy is the progressive loss of muscle tissue due to chronic degeneration of muscle and to the exhaustion of satellite cells that replace damaged fibers. Thus, the persistent protein degradation observed in neuromuscular diseases reflects a pathological muscle catabolism (Figure 3).

Figure 2. Cartoon of the skeletal muscle contractile apparatus and cytoskeletalinteractions (from Dalakas et al., 2000).


37

Figure 3. Pathwaysin muscle survivaland Wasting.

A.

Muscle survival factors

It is generally accepted that the primary cause of functional impairment in muscle is a cumulative failure to repair damage resulting from sustained muscular activity, related to an overall decrease in anabolic processes. Among growth factors, the insulin-like growth factors (IGFs) have been implicated in the control of skeletal muscle growth and differentiation in embryonic development (Liu et al.. 1993). In the adult, IGF-J has been implicated in many anabolic pathways in skeletal muscle, where it plays a central role during muscle regeneration (Liu et al.. 1993; Powell-Braxton et at.. 1993: Florini et al.. 1996). Since IGF-1 levels decline with a g e in both rodents and hmnan muscle, this growth factor has been considered a promising therapeutic agent in staving off advancing muscle weakness during ageing and promoting regeneration. Circulating IGF-1 is synthesized in the liver; however, specific isoforms of the IGF-1 gene product. produced by alternate promoter usage and splicing, are locally synthesized in skeletal muscle fibers themselves in response to stretch exercise as well as to injury (McCoy et al.. 1999). Recent studies of the effects of IGFs on the myogenic program in muscle cell culture have focused specifically on IGF-1. since it is the predominant form in mature muscle tissue. In cultured myoblasts, as in other cell types, IGF-1 induces cell proliferation (Quinn et al.. 1994: Engert et ah. 1996). However unlike other growth factors, IGF-1 also stimulates myogenic differentiation and generates pronounced myocyte hypertrophy (Florini et al., 1986; Quinn et al., 1994; Engert et al., 1996), suggesting that this growth factor can regulate both proliferative and differentiative responses in muscle cells. Studies in muscle cell culture (Engert et al., 1996) have established that administration of exogenous IGF-1 elicits a biphasic response, first stimulating cell proliferation and

38

A. MusarO and IV. Rosenthal

subsequently enhancing myogenic differentiation, a sequence of events ideally timed for the tissue regeneration. To determine whether IGF-1 can stimulate myogenesis in the absence of proliferation, we established an experimental cell culture system where the action of IGF-1 on proliferation and on differentiation could be uncoupled. The neonatal rat L6E9 myogenic line has been used extensively to study the role of IGF-1 in myogenesis (Yaffe, 1968; Nadal-Ginard, 1978), as it does not express this growth factor itself, and can respond to IGF-1 stimulation through presentation of IGF-1 receptors. Treatment of L6E9 myoblasts with exogenous IGF-1 in defined serumfree medium induces a transient increase in cell cycle markers and cell proliferation (Rosenthal and Cheng, 1995; Engert et al., 1996). This response is rapidly followed by withdrawal from the cell cycle and activation of myogenic factors, resulting in a net increase in structural gene expression and larger myotubes (Engert et al., 1996). To bypass the proliferative effects of IGF-1. we stably transfected L6E9 cultures with a cDNA encoding a muscle-specific, locally actin~ IGF-1 isoform, (mIGF-1) driven by the fast muscle-restricted regulatory elements from the myosin light chain (MLC)I/3 locus, which activate gene expression only after myoblasts have withdrawn from the cell cycle and have committed to differentiation. In this way, the influence of IGF-1 on myogenic differentiation could be dissected and manipulated independently of its role in myoblast proliferation. Studies on the MLC/IGF-1 transfectants suggest that post-mitotic expression of IGF-1 potentiates and accelerates the myogenic program. induces dramatic myotube hypertrophy, activates a new battery of genes and initiates a shift towards a more mature fiber type. These studies in cell culture support a role for IGF-1 in the establishment and maintenance of the mature muscle phenotype in normal and cultured muscle tissue. Although cell culture models generate valuable models of IGF-1 action, elucidating the molecular basis of IGF-1 function in vtvo is critical for exploring the subcellular events leading to muscle atrophy, as well as for designing and testing therapeutic strategies for enhancing the robustness of native muscle. The fact that IGF-1 can act either as a hormone or as a local growth factor has complicated the analysis of animal models in which transgenic IGF synthesized in extrahepatic tissues was released into the circulation (Mathews et al., 1988; Coleman et al.. 1995; Reiss et al., 1996; Delaughter et al.. 1999), therefore determining pathological side effects. Transgenic mine generated with an alternate isoform of IGF-1 driven by a skeletal actin promoter demonstrated significant increases in muscle mass (Coleman et al., 1995). Likewise, transgenic mice carrying an MLC/mIGF-1 gene exhibited sustained muscle hypertrophy, producing pronounced increases in muscle mass and strength with no undesirable side effects (Musar5 et al., 2001). In these studies, restricting the action of supplemental IGF-1 to the tissue of origin by choice of the appropriate isoforms allowed the assessment its autocrine/paracrine role in skeletal muscle throughout the lifespan of the animal, exclusive of possible endocrine effects on other tissues. Expression of tile mlGF-1 transgene safely enhanced and preserved muscle fiber integrity even at advanced ages (Musar5 et al., 2001), suggesting that the MLC/mIGF-1 transgene acts as a survival factor by prolonging the regenerative potential of younger muscle (Figure 4). This is supported by preliminary evidence demonstrating reactivation of myoblast proliferation following terminal differentiation of MLC/mIGF-1 primary


39

)

Tg C D X - 6 d .

Figure 4.

-

T g C D X -2d-:

-

Muscle regenerative capacity after injury is retained in senescent MLC/mIGF-1 transgenic mice.

TA muscles of 22 month old Des/nls-lacZ transgenic mice (for simplicity indicated as Wtj or MLC/mIGF-1 x Des/LacZ double transgemc mice (Tg) were injured by a single injection of cardioxin (CDX: see Materials and MethodsJ and monitored for Des/nls-LacZ transgene activation by X-gal staining. Injured Des/nls-LacZ muscle did not activate the transgene at any time (Wt CDX -2d- panel, and data not shown ~.The MLC/mIGF- 1 x Des/nls-LacZ muscle at day 2 post-injury displayed X-gal transgene activation both in undamaged and damaged portions of the muscle (Tg CDX -2d- panels). Activation of satellite cells from quiescent status to proliferative phase was confirmed by PCNA (Tg CDX -2d- insem and MyoD (data not showm expression. Satellite cell activation in the MLC/mIGF-1 x Des/nls-LacZ muscle fibers was also confirmed by endogenous desmin gene activation (data not shown). A1 day 6. X-gal staining nuclei were present in MLC/mIGF-1 x Des/nls-LacZ myofibers, concomitant with the activation of the neonatal isoform of MyHC expression (Tg CDX -6d- panels). X-gal staining in MLC/mIGF-1 x Des/nls-LacZ myofibers at days 12 after injury documented the peripherization of positive nuclei, suggesting the end point of the regeneration program (from Musar6 et al.. 2001 J.

m y o c y t e c u l t u r e s ( A . M . , u n p u b l i s h e d o b s e r v a t i o n s ) so that m I G F - 1 p r o d u c e d b y postm i t o t i c m y o c y t e s a p p e a r s to p r o m o t e r e g e n e r a t i o n b y e x t e n d i n g s t e m cell p r o l i f e r a t i v e capacity as well. T h e d r a m a t i c i n c r e a s e in r o b u s t n e s s a n d s p o n t a n e o u s c o n t r a c t i l i ~ o f d i f f e r e n t i a t e d p r i m a r y cultures f r o m the M L C / m l G F - 1 t r a n s g e n i c m u s c l e , e v e n after p r o l o n g e d p e r i o d s (up to 60 d a y s ) in s e r u m - f r e e m e d i a a n d in the a b s e n c e o f

40


innervation, further suggests that expression of mIGF-1 could increase the survival of postmitotic myocytes ex vivo. The increase and retention of regenerative capacity due to sustained supplemental IGF-1 production is indicative of a fundamental change m muscle metabolism resulting in the persistent activation and replenishment of stem cells, and selective sparing or replacement of the most powerful fibers. The capacity of the mIGF-1 transgene to attenuate the structural and functional consequences of muscle aging was independent of its action during embryogenesis or early postnatal life. since local delivery of mIGF-/ in individual mouse muscles by AAV virus mediated gene transfer also permanently blocked age-related loss of muscle size and strength, presumably by improving regenerative capacity (Barton-Davis et al., 1998) through increases in satellite cell activity (Barton-Davis et al., 1999). Young adult muscles receiving virally delivered IGF-I exhibited muscle hypertrophy, while the functional properties of old muscles injected with the IGF-I virus approached those of their younger uninjected counterparts. Changes in muscle mass and cross sectional area of mIGF-I injected or transgenic muscles also translated into increased force production. In addition, age-related reduction in force production and loss of fast fibers, all of which are typical of ageing skeletal muscle, were prevented both by virally delivered and by transgenic IGF-I gene expression (Barton-Davis et al., 1998; Musaro et al., 2001). These studies provide a model wherein skeletal muscle performance is enhanced and maintained by local IGF-I synthesis throughout the lifespan of the muscle. Local persistent synthesis of supplemental IGF-I could also prevent the loss of muscle function that begins at mid adulthood in healthy individuals, and is vastly accelerated in patients with neuromuscular pathologies. B.

The calcium pathway

The plasticity of fiber identity is an essential feature of muscle function in the adult animal. Changing innervation patterns cause a corresponding reprogramming of fiberspecific gene expression, but the intracellular mechanisms responsible for transducing the neuronal input are less clear. Calcium is an important intracellular messenger in muscle, controlling numerous cellular process including proliferation, cell growth, differentiation, and apoptosis (Berridge et al., 2000). All muscle fibers use Ca z+ as their main regulatory and signal molecule (Berchtold et al., 2000). Slow twitch oxidative fibers maintain intracellular Ca 2+ concentration at high levels (100-300 nM), whereas fast twitch glycolitic fibers are characterized by brief, high ampfitude Ca 2+ transients and lower ambient Ca 2+ levels (50 nM). Defects in signaling pathways can compromise the survival of the myofibers in muscle wasting. Dystrophin, a membrane associated protein that contributes to muscle structural integrity, also regulates intracellular Ca 2+ homeostasis (Gillis, 1996). In the absence of dystrophin, alterations in intracellular Ca 2+ may contribute to an imbalance between muscle protein synthesis and protein degradation, culminating in necrosis, fibrosis, and shift in fiber content (Engel et al., 1994).


41

The intermediate signaling molecule modulating many of these processes is the phosphatase calcium-dependent calcineurin (Olson and Williams, 2000). Calcium signaling is not only important for muscle function, but is also involved in specifying the slow muscle phenotype (Chin et al.. 1998~. Intracellular calcium levels may therefore modulate fiber-specific signal transduction pathways, adding to the plasticity of the differentiated state. The pronounced myocyte hypertrophy induced by post-mitotic IGF-1 gene expression in stably transfected muscle cultures (Musar6 and Rosenthal, 1999) is the result of a calcineurin-mediated response to increased intracellular calcium, which underlies cardiomyocyte hypertrophy (Musaro et al., 1999). Forced expression of either mlGF-1 or activated calCineurin in muscle cultures promote muscle hypertrophy inducing expression of GATA-2. a transcription factor which accumulates exclusively in hypertrophic myotube nuclei complexed with calcineurin and NFATcl, a target of calcineurin phosphatase activity. Exploiting a variety of genetic and pharmacological manipulations has established a conclusive link between IGF-1 action and the calcineurin signaling pathway in myocyte hypertrophy. Analysis of the molecular parameters of muscle hypertrophy in MLC/mIGF-1 animals confirmed that the signal transduction pathways responsible for activating hypertrophic gene expression programs in cell culture models also operate to induce skeletal muscle hypertrophy in vivo. Since expression of the mIGF-1 transgene in otd MLC/mIGF-1 transgenic mice is protective against normal loss of muscle mass during senescence (Musar6 et al., 2001), the calcineurin signal transduction pathway activated by IGF- 1 appears to play a pivotal role in the survival of muscle as well. This notion is supported by the reports that transplant patients maintained on cyclosporine (CsA)treatments develop muscle disorders (Breil and Chariot, 1999). It has been reported that CsA. which is one of the specific inhibitors of calcineurin activity, promotes myopathy manifesting by myalgia, muscle weakness, and plasma creatin kinase elevation (Fernandez-Sola et al., 1990: Goy et al.. 1989: Grezard et al., 1989: Noppen et al., 1987), all symptoms associated to muscle wasting. In fact histological analysis has revealed muscle atrophy, necrotic fibers, excessive number of central nuclei, and mitochondria abnormalities, including ragged-red fibers and lipid vacuoles (Femandez-Sola et al., 1990; Lamer et al.. 1994). Calcineurin is not the only intermediate molecule of the Ca 2+ signaling apparatus in muscle cells. Ryanodine receptor, calsequestrin, parvalbumin, calmodulin. $100 proteins, annexin, beta-catenin and calpain are Ca2+-dependent molecules which modulate protein metabolism, differentiation and growth of muscle cells (Berchtotd et al., 2000) and alterations in their function have been shown to be associated to muscle diseases, such as dystrophinopathies. Recently, the CaZ+-catmodulin dependent protein kinase family has been implicated in stimulation of muscle differentiation through disruption of HDAC-MEF2 complexes and activation of MEF2-mediated transcription (Lu et al., 2000; Olson and Williams 2000). In summary, it is clear that dysfunction of the system~ controlling Ca 2+ homeostasis cart be detrimental for the normal" function and survival of muscle fibers (Figure 5).

42


Figure 5. Signal transduction pathways in skeletal muscle survival and wasting : (Modifiedfrom Muscat and Dressel, 2000).

C.

The PI3-kinase pathway

Another important mediator of extracellular signaling is the phospho-inositol kinase (PI3K), composed of two subunits: a 110 kDa catalytic subunit and two alternate isoforms of an 85 kDa regulatory subunit. Activation of the PI3-kinase pathway evokes specific responses in skeletal muscle, including protein translation, differentiation and glucose metabolism. In addition, the ability of growth factors to promote survival has been attributed, at least in part, to the PI3-kinase activation (Datta et al., 1999). The survival stimulus starts with the binding of trophic factors to a transmembrane receptor. This protein-protein interaction stimulates the recruitment of PI3K to the inner surface of the plasma membrane and the activation of the serine-threonine protein kinase Akt, which in turn determines the inactivation of death-promoting molecules (Lawlor and Rotwein, 2000a-b). Although most of our knowledge of this pathway in skeletal muscle derives from in vitro culture systems, the same signals are presumably activated in vivo to maintain the integrity of skeletal myofi bers phenotype and function (Zhu et al., 2001). The differentiation program of skeletal muscle cells can be recapitulated in in vitro culture as two functionally distinct stages, proliferation and differentiation. Proliferating myoblasts can undergo either terminal differentiation or apoptosis under conditions of mitogen deprivation (Figure 6). Post-mitotic myoblasts committed to the differentiation pathway express proteins that protect them from programmed cell death. Specifically,

43


p2t Myogeni n

chromatin condenses ceil shrinkage

Bd2

GM Figure 6, Differentiationand apoptosisin skeletalmuscle.

the apoptosis-resistant phenotype has been correlated with the induction of the cyclindependent kinase inhibitor p21 cipuwafl in these cells (Wang and Walsh, 1996). Among the many growth factors involved in muscle cell survival. IGF-1 has been specifically implicated in PI3K-mediated signal transduction. Chakravarthy et al. (2000) reported that IGF-1 extends the replicative life span of skeletal muscle satellite cells in v~tro via the activation of the PI3-kinase pathway. As previously discussed, IGF-1 also stimulates myogenic differentiation and generates hypertrophy of myocytes in vivo and in vitro (Florini et al.. 1986: Rosenthal and Cheng 1995; Engert et al., 1996; Musar6 and Rosenthal, 1999), suggesting that this growth factor can regulate both proliferative and differentiative responses in muscle cells. Experimental evidence suggests that IGF-1 exerts its functions by activating two different intracellutar signal transduction pathways, conveying proliferative and differentiative signals, respectively. The proliferative response is mediated by the MAP-kinase pathway (Coolican et al.. 1997), whereas the pathway leading to differentiation involves the activation of PI3K (Kaliman et al., 1996; Coolican et al., 1997). The majority of biological actions of IGF-1 are mediated by the type 1 IGF-1 receptor (IGF1R) and by several intermediate molecules known as Insutin-Receptor-Substrates (IRSs) (Whitehead et al., 2000). Once activated, IGF1R undergoes a conformational change leading to autophosphorylation of tyrosine residues that serve as recruiting

44


sites for cytoplasmic proteins, including IRS proteins, containing Src-Homology domain 2 (SH2). IRS molecules are associated with the IGF1R in clathrin-coated pits at the cell surface, creating a scaffold for attracting downstream molecules, such as PI3K. The interaction of IRS proteins with the cytoskeleton provides a controlled way of delivering intermediate molecules to their appropriate site of action. Among IRS molecules, the subunit IRS-1 appears to be involved in the transduction of the survival signal though direct association with PI3K (Conejo and Lorenzo, 2001). It has recently been demonstrated that IGF-1 promotes muscle celt survival through PI3K induction of the myogenic regulatory protein MyoD. Ectopic expression of MyoD induces p21, whereas inhibition of p21 blocks MyoD-mediated survival (Lawlor and Rotwein, 2000). MyoD also plays a pivotal role in muscle survival during vertebrate embryogenesis, where the signals required to determine presomitic mesoderm are important to control cell growth, differentiation and survival. In particular, Borycki et al. (1999) reported that the defect in MyoD activation by surface ectoderm and neural tube/notochord in Pax3 deficient mice is associated with programmed cell death of somitic cells. These studies implicate MyoD as the mediator of least one PI3K-dependent survival pathway in skeletal muscle. On the other hand, loss of MyoD expression does not block the effects of IGF-1 on myocyte survival in culture (Lawlor and Rotwein, 2000), suggesting a second pathway involving PI3K-mediated activation of Akt, with subsequent induction of p21. D.

MAP-Kinase pathway

The crosstalk between signaling pathways also involves the interaction of Ca2+ with mitogen-activated protein kinase (MAPK) signaling. The members of the MAPK family are predominantly activated by mitogenic stimuli (Nebreda and Porras, 2000). However, a growing body of evidence has implicated MAPK proteins in non-mitogenic stimuli, including muscle differentiation and survival. Among the members of the MAPK family, p38 plays a critical role in the transduction of non-mitogenic stimuli in different cell types (Nebreda and Porras, 2000; Ono and Han, 2000). Four isoforms of p38-MAPK (c~-8) have been identified. The p38~/ isoform is preferentially expressed in skeletal muscle and induced during muscle differentiation (Lechner et al., 1996; Li et al., 1996). In muscle cell cultures, p38 stimulates myogenic differentiation, regulating MyoD activity via phosphorylation and increasing the transcriptional activity of the co-regulator MEF2C (Zetser et al., 1999). Moreover, treatment of C2C12 myoblasts with the inhibitor of p38, SB203580, delays elongation and alignment of myoblasts, prevents fusion and inhibits the induction of musclespecific genes (Cuenda and Cohen 1999). These observations suggest that p38 activates common targets of several signal transduction pathways to induce a specific survival response. The involvement of the MAPK pathway in myogenic differentiation functions is further supported by the report that the expression of caveolin-3, a membrane protein that is essential for the myoblast fusion, requires p38 activation (Galbiati et al., 1999). Caveolins are the principal components of the caveolae, vescicular invaginations of the plasma membrane, which participate to trafficking events and signal transduction


45

processes (Smart et al., 1999). Caveolin-3 is localized to the sarcolemma where it forms a complex with cytoplasmic signaling molecules (G-proteins and Src-like kinases), dystrophin and its associated glycoproteins, such as alpha-sarcoglycan and betadystroglycan (Song et al., 1996). The fact that autosomal dominant limb girdle muscular dystrophy (LGMD) has been associated with a severe deficiency of caveolin-3 in muscle fibers (Minetti et al.. 1998) further highlights the importance of caveolins in the pathogenesis of muscular dystrophy. In this context. MAPKs could play a pivotal role in the maintenance of the differentiated state and in myofiber survival.

IV.

The proteolytic systems in skeletal muscle tissue

Most muscle pathologies are characterized by the progressive toss of muscle tissue due m chronic degeneration combined with the inability of regeneration machinery to replace damage muscle tissue. The persistent protein degradation observed in muscle diseases reflects a pathological muscle catabolism, known as muscle wasting or cachexia. The continual synthesis and degradation of cell proteins is the result of normal intracellular metabolism mad represents an important homeostatic function of muscle tissue. Altering the homeostatic set point is detrimental for myofibers survival and muscle functional integrity. Several signaling pathways that promote cellular survival are active in muscle fibers. Mutations m one or more component of these survival-signal transduction pathways can lead to muscle wasting. Two proteolytic systems, calpains and proteasome-mediated ubiquination, have been associated with muscle wasting and are activated when the survival program does not work properly, thereby compromising myofiber survival. A.

The calpain pathway

Calpains are Caicium~activated cysteine proteases that partiCipate in various intracellular signal transduction pathways mediated by Ca 2+ (Sorimachi et al., 1997): On the basis of Ca 2+ concentration, two ubiquitous isoforms, ~t and m, are welt characterized and several other isoforms are selectively expressed in mammalian tissues. Calpain 3 is predominantly expressed in skeletal muscle and maintains proteolytic activity at physioiogical Ca 2+ (Sorimachi et a l , 1989) Alteration in Calpain3 function is associated with limb-girdle muscular dystrophy type 2A (Baghdiguian et al., 1999). Calpalns consist of two subunits, a 80kDa large subunit which contains protease activity and a 30kDa smaller subunit, functioning as a regulator of catpain activity: Calpmns are activated by several stimuli in which intracetlular Ca 2+ homeostasis is affected, causing disruption of the contractile tissue (Cannon et al., I99t), mitochondriat swelling, sarcoplas~c reticulum vacuolization (Armstrong et al., 1991; Frid6n et aI,, t989) and sarcomeric alterations (Belcastro et al., 1998). Calpains are preferentiaUy localized in the Z disk of the sarcomere (Kumamoto et al., 1992) where they initiate the proteolytic cleavage o f muscl e protein, causing the complete disassemNy of myofibrils and loss of the Z diSk (Huang and Forsberg, 1998)i The activity of calpains is regulated by the endogenous inhibitor calpastatin which

46


prevents both enzyme activation and expression of catalytic activity. The interactions between calpastatin and calpains are modulated by intracellular Ca 2+ concentration. The calpain isozymes are associated with organelles and molecules involved in metabolic and signal transduction pathways, such as glycolytic enzymes, protein kinase A and C, phospholipase C and cytoskeletal protein, such as desmin, vimentin, integrin, cadherin, c~-actinin (Saido et al., 1994), The non-erythroid spectrin protein known as fodrin, which is a multifunctional protein and a major component of the cortical cytoskeleton of most eukaryotic cells including muscle cells, is another molecular target of calpains (Huang and Forsberg, 1998). Cleavage of fodrin accompanies apoptosis induced by ligation of the CD3/T cell receptor complex, Fas ligation, or treatment of cells with staurosporine, glucocorticoid, or synthetic ceramide (Martin et al., 1995; Planey and Litwack, 2000), all factors that affect muscle physiology and myofiber survival. In contrast to other proteolytic systems, calpains cleave target proteins at specific sites, leaving large polypeptide fragments with altered physiological properties. In addition, post-translational modifications provide a mechanism of "marking" specific proteins for calpain degradation. Phosphorylation of Troponin T and I by protein kinase A and C dramatically alters the sensitivity of the protein to calpain degradation (Di Lisa et al., 1995). In addition, calpain is associated with neutrophil accumulation during exercise (Kunimatsu et al., 1995), suggesting a role for calpains in muscle injury. The recruitment of neutrophil represents the first line of defense against pathogen agents and their numbers increase in injured muscle. Collectively these studies suggest that calpains play a key role in muscle homeostasis and comprise an addition mechanism whereby alteration in Ca 2+ concentration triggers cellular damage. B.

The ubiquitin-proteasome pathway

The ubiquitin-proteasome pathway plays a key role in the turnover of muscle protein and is activated in several catabolic process, such as cancer, fasting, acidosis, sepsis, and glucocorticoid treatment, leading to muscle wasting (Mitch and Goldberg, 1996; Mitch et al., 1999). Protein substrates are first marked for degradation by conjugation to small protein cofactor, ubiquitin. The pathway involves an enzymatic cascade starting with the ubiquitination of muscle protein to be degraded by the 26S proteasome in a process that unfolds the protein, releases ubiquitin, and degrades the protein to small peptides and amino acids (Mitch and Goldberg, 1996; Zamir et al., 1992). The ubiquitin conjugation to protein substrates requires ATP. The first step of this process involves the activation of the ubiquitin molecule by an ATP-requiring enzyme, El. Activated ubiquitin is then transferred by E1 to a carrier protein E2; in the last step the ubiquitin-protein ligase E3 catalyzes the covalent interaction between the carboxyl group of ubiquitin and the e-amino group of lysine in the protein substrate. These reactions are repeated to form a ubiquitin chain. The ubiquitin-conjugated proteins are then transferred, in an ATP-dependent reaction, in the 26S proteasome complex containing multiple proteolytic sites where the proteins are degraded to amino acids by pepfidases. The ubiquitin itself is released and reused for a new enzymatic reaction.


47

Among hormones and cytokines that affect muscle homeostasis, glucocorticoids play a pivotal role in muscle degeneration by stimulating proteotysis and inhibiting protein synthesis, and are important factors in the development of muscle atrophy in various catabolic conditions (Mitch and Goldberg, 1996). Glucocorticoids also induce an increase in ubiquitin transcript levels and enhance the conversion of amino acids to glucose in the liver. Glucocorticoid-stimulated muscle protein breakdown is therefore primarily mediated by ubiquitin-proteasome-dependent proteolysis, although calciumdependent protein degradation may also be involved. In certain catabolic conditions. including sepsis, an interaction between glucocorticoids and proinflammatory cytokines is also involved in the exacerbation of muscle protein breakdown.

V.

Therapeutic strategies to attenuate muscle wasting and promote myofiber survival

The prolongation of skeletal muscle strength in aging and neuromuscular disease has been the objective of numerous approaches employing a variety of delivery systems. The continued study of factors and molecular pathways that participate in the prevention of muscle catabolism will be important for the attenuation of muscle wasting in a clinical setting. The generation of appropriate experimental models is particularly critical for the development of therapeutic strategies m attenuate muscle wasting and promote myofiber survival. Although to date a mouse model of muscle cachexia is lacking, currently available animal models of muscular dystrophy, neuromuscular degeneration and disuse atrophy can be employed to test new interventions for muscle atrophy. Cell-trased therapies, stalled by the difficulty in obtaining sufficient numbers of autologous myoblasts and by inefficient incorporation into host muscle (reviewed in Grounds, 2000) have recently enjoyed a renaissance following the observation that bone marrow stem cells were competent to contribute to skeletal muscle (Ferrari et al., 1998). More recently techniques have been refined for isolating totipotent stem cells from murine skeletal muscle, that were capable of reconstituting the bone marrow stem cell population in lethally irradiated hosts (Gussoni et al., 1999). Since the small number of muscle stem cells that can be isolated from normal muscle is a limiting factor, viable therapeutic strategies based on autologous material depend upon the development of methods to increase the muscle stem cell population. The persistence of regenerative potential in the muscles of MLC/mIGF-1 transgenic mace, as well as the reversal of age-related muscle atrophy by infection of mature muscles with virally delivered MLC/mIGF-1. provides a promising venue for expanding mature stem cell populations by administration of mIGF-1 genes, either in situ or ex vivo, Other aspects of myocyte survival tactics may prove useful for devising therapeutic strategies to combat muscle degeneration. Blocking proteotytic pathways or inducing muscle regeneration with pharmacological intervention will require a detailed knowledge of the signaling mechanisms involved, and the extent to which blocking or enhancing these pathways will be detrimental to other ph) siologieal parameters such as motor innervation. Finally, the potential feedback between skeletal muscle and other tissue types such as heart, adipose tissue and bone is a major aspect of physiology that merits

48


consideration, and which will undoubtedly play an important role in promoting muscle plasticity and survival in the context of the clinic.

VI.

References

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Rudnicki M. A. and Jaenisch R. (1995). The MyoD family of transcription factors and skeletal myogenesis. Bioessaysl 7, 203-9. Saido T. C., Sorimachi H., and Suzuki K. (1994). Calpain: new perspectives in molecular diversity and physiological-pathological involvement. FASEB J 8, 814-22. Schiaffino S., and Reggiani C. (1996). Molecular diversity of myofibrillar proteins: gene regulation and functional significance. Physiol Rev 76, 371-423. Semsarian C., Wu M. J., Ju Y. K., Marciuiec T:, Yeoh T., Allen D., Harvey R. P., and Graham R. M. (1999). Skeletal muscle hypertrophy is mediated by a Ca2+-dependent caleineurin signalling pathway. Nature 400, 576-81 Smart E. J., Graf G. A., McNiven M. A., Sessa W. C., Engelman J. A., Scherer P. E., and Okamoto T., Lisanti MP (1999). Caveolins, liquid-ordered domains, and signal transduction. Mol Cell Biol 191, 7289-304. Song K. S., Scherer P. E., Tang Z., Okamoto T., Li S., Chafel M., Chu C., Kohtz D. S., and Lisanti M. P. (1996). Expression of caveolin-3 in skeletal, cardiac, and smooth muscle cells. Caveolin-3 is a component of the sarcolemma and co-fractionates with dystrophin and dystrophin-associated glycoproteins. J Biol Chem 271, 15160-5. Sorimachi H., Imajoh-Ohmi S., Emori Y., Kawasaki H., Ohno S.0 Minami Y., and Suzuki K. (1989). Molecular cloning of a novel mammalian calcium-dependent protease distinct from both m- and ran-types. Specific expression of the mRNA in skeletal muscle. J Biol Chem 264, 20106-11. Sorimachi H., Ishiura S., and Suzuki K. (1997). Structure and physiological function of calpains. Biochem J 328,721-32. Sutherland C. J., Esser K. A., Elsom VL., Gordon ML., and Hardeman EC. (1993). Identification oi' a program of contractile protein gene expression initiated upon skeletal muscle differentiation. Dev Dyn 196, 25-36. Wade R., Sutherland C., Gahlmann R., Kedes L., Hardeman E., and Gunning P. (1990). Regulation of contractile protein gene family mRNA pool sizes during myogenesis. Dev Biol 142, 270-82. Wang J., and Walsh K. (1996). Resistance to apoptosis conferred by Cdk inhibitors during myocyte differentiation. Science 273, 359-61. Whitehead J. P, Clark S. F., Urso B., and James D. E. (2000). Signalling through the insulin receptor. Curr Opin Cell Biol 12, 222-8. Yaffe, D. (1968). Retention of differentiation potentialities during prolonged cultivation of myogenic cells. Proc. Natl. Acad. Sci. USA 61,477-483. Zamir O., Hasselgren P. O., O'Brien W., Thompson R. C., and Fischer J. E. (1992) Muscle protein breakdown during endotoxemia in rats and after treatment with interleukin-1 receptor antagonist (IL-lra). Ann Surg 216, 38~1-5. Zetser A., Gredinger E., and Bengal E. (1999). p38 mitogen-activated protein kinase pathway promotes skeletal muscle differentiation. Participation of the Mef2c transcription factor. J Biol Chem 274, 5193-200 Zhu W. Z., Zheng M., Koch W. J., Lefkowitz R. J., Kobilka B. K., and Xiao R. P. (2001). Dual modulation of cell survival and cell death by beta(2)-adrenergic signaling in adult mouse cardiac myocytes. Proc Natl Acad Sci USA 98, 1607-12.

CHAPTER

3

INTERACTIONS B E T W E E N THE CELL CYCLE AND THE MYOGENIC PROGRAM

JING HUANG

and MATt

J. T H A Y E R

Table of contents I. II.

III. IV.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cell cycle overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Regulation of cdk activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. The retinoblastoma susceptibility protein (RB) . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Phosphorylation of RB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Cell cycle checkpoints . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . M y o D family and muscle formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Terminal cell-cycle arrest of myoblasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Regulation Of MyoD activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. The role of RB during terminal growth arrest of myoblasts . . . . . . . . . . . . . . . . . . C. Interaction of MyoD with RB mediates terminal cell cycle arrest D. E. F.

of myoblasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . MyoD-mediated p21 induction during terminal myoblast growth arrest . . . . . . . . . Direct inhibition of cyclin kinase activity by MyoD . . . . . . . . . . . . . . . . . . . . . . . . p300/CBP cooperates with MyoD during muscle differentiation . . . . . . . . . . . . . .

V.

Maintenance of the differentaited state . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. A role for RB and p21 for maintaining G1 arrest . . . . . . . . . . . . . . . . . . . . . . . . . . . B. A role for cyclin D3 in preventing D N A synthesis in myombes . . . . . . . . . . . . . . . VI. A role for M D M 2 and ATR in muscle tumor formation . . . . . . . . . . . . . . . . . . . . . . . . . A. Amplification of M D M 2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Duplication of ATR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Cell cycle checkpoints and differentiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

L

53 54 54 55 55 56 57 59 59 60 61 61 61 62 62 62 63 64 65 66 67 68

Introduction

P r o l i f e r a t i n g m a m m a l i a n cells c a n e x i t t h e cell c y c l e a n d e n t e r a q u i e s c e n t s.tate k n o w n as GO w h e n g r o w t h f a c t o r s are r e m o v e d . C e l l s i n GO c a n e x i t f r o m this r e s t i n g state a n d r e i n i t i a t e c e l l u l a r p r o l i f e r a t i o n i n r e s p o n s e to g r o w t h stimuli. I n contrast, skeletal m y o b l a s t s in t h e p r o c e s s o f d i f f e r e n t i a t i o n w i t h d r a w p e r m a n e n t l y f r o m t h e celt cycle. 53 Advances in Developmental Biology and Biochemistry Ed, David Sassoon 53 -- 74 © 2002 Elsevier Science. Printed in the Netherlands

54

J. Huang and M. J. Thayer

This permanent cell cycle withdrawal is characterized by a loss in their ability to reenter the cell cycle in the presence of growth factor stimulation (Endo and Nadal-Ginard, 1986; Gu et al., 1993; Konigsberg et al., 1960; Stockdale and Holtzer, 1961). Thus, in the transition from cycling myoblasts to terminally differentiated myotubes, and during the maintenance of the differentiated state, these cells must develop additional blocks to prevent them from reentering the cell cycle. The process of muscle differentiation is characterized by three orderly events: 1) the terminal cell cycle arrest of myoblasts; 2) the coordinated activation of musclespecific gene expression and; 3) myotube formation. The regulatory factors involved in these orchestrated events can be divided into two groups: 1) muscle-specific transcription factors, such as MyoD, myogenin, myf-5, and MRF4 and; 2) growth inhibitors such as p21 and RB. Interactions between these two classes of regulatory proteins play a major role throughout the entire sequence of events that ultimately leads to the fully differentiated muscle. The goal of this chapter is to review coordinated interactions between these regulatory factors from the terminal cell cycle arrest to muscle gene activation of the differentiated state.

II.

Cell cycle overview

The cell cycle is the process through which cells duplicate themselves. The cell cycle is divided into four phases--G1, S, G2, and M. The two major events that occur during each cycle are: S phase, in which the DNA is replicated, and M phase, in which ceils divide to generate two daughter cells. In eukaryotic cells, cell cycle progression involves a series of tightly regulated and coordinated events. A.

Regulation of cdk activity

The driving force for cell cycle progression is the activation of cyclin dependent kinases (cdks). As its name implies, the activity of cdks is dependent upon interactions with cell cycle regulated proteins known as cyclins. Cyclins are a family of proteins that share a conserved region of about 100 amino acids known as the cyclin box, each expressed during a specific phase of the cell cycle (Sherr, 1994). The cyclin box serves as a docking site that recruits cdk inhibitors or cdk substrates to cyclin-cdk complexes. Each cyclin binds and activates specific cdks at specific times during the cell cycle, which ensures the activation of specific cyclin-cdk complexes only at the appropriate time during the cell cycle. Cyclin levels are regulated by ubiquitin-mediated proteolysis (Slingerland and Pagano, 1997), and at the level of transcription (Cheng et al., 1998; Matsushime et al., 1991; Muller et al., 1994), this dual regulation ensures that each cyclin is present only during specific cell cycle stages. During the early stages of G1, D-type cyclins accumulate within the cell where they interact with cdk4 and cdk6. During late G1, E-type cyclins accumulate and interact with cdc2 (Sandhu and Slingerland, 2000). Together, the D and E type cyclins promote cell cycle progression from G 1 into S phase.

The Cell Cycle and Myogenic Differentiation

55

In addition to tight control over expression timing, cyclin-cdk complexes are regulated by cyctin kinase inhibitors (cki)~ The ckis can be divided into two families: INK4, which inhibits cdk4 and cdk6 t Sandhu and Slingerland. 2000: Sherr and Roberts. 1995), and KIP, which appear to inhibit all cdk family members {Sandhu and Slingerland. 2000). The INK4 family includes pt5, pl6, pl 8, and plg. and is functionally restricted to the G1 phase, where cdk4 and cdk6 function. INK4 family members are thought to compete with D-type cyclins during G1. forming independent binary complexes with cdk4 or cdk6 and thereby inhibiting the activity of the cyclin D/cdk4 or cdk6 complexes. In comparison with the 1NK4 family, the KIP family includes three broadly acting members: p21 (cipl), p27 (kipl), and p57 (kip2). KIP family members are able to interact with multiple cyclin-cdk complexes to inhibit their activities throughout the entire cycle (el-Deiry et al., 1993: Harper et al., 1993; Harper et al.. 1995). B.

The retinoblastoma susceptibility protein (RB)

The retinoblastoma susceptibility protein (RB) was first identified as the product of the retinoblastoma susceptibility gene RB-1 (Wang et al., 1994). This gene was the first rumor suppresser gene identified, and it has growth inhibitory functions. The RB protein ts a member of a family of pocket proteins, including RB. p107. and p130, which are conserved through evolution (Wang et al., 1994). These pocket proteins are not considered as classic transcription factors, due to their lack of sequence-specific DNA-binding activities. Instead. they are recruited to promoters through interactions with other transcription factors, such as E2F family members. E2F-1 was the first cellular protein shown to interact with RB. Interactions between E2F family members and RB occur through the C-terminal activation domain of E2F. This interaction is thought to mask the E2F transactivation domain. In addition, because RB can interact simultaneously with both E2F and certain cellular and viral proteins. including transcriptional repressors such as RBP1 (Lai et al.. 1999), HBP1 (Tevosian et al., 1997), RIZ (Buyse et al., 19951. and the histone deacetylases HDAC1 and HDAC2 (Dunaief et al,, 1994: Fattaey et al., 1993; Lee et al., 1998; Trouche et al.. 1997: Welch and Wang, 1993). Therefore. it has been proposed thai the RB-E2F. p130-E2F, and pl07-E2F complexes actively repress E2F-dependent transcription by two mechanism, one involving the masking of the activation domains of E2F members and the other by recruitment of transcriptional repressors. The recruitment of HDACs is thought to result in the deacetylation of histories at or near the E2F dependent promoters. Histone deacetylation is thought to strengthen the interactions between the negatively charged DNA backbone and positively charged histories, which in turn interferes with transcription. C.

Phosphorylation of RB

The interactions between RB and the E2F family members are regulated by the phosphorylation status ofRB. Phosphorylation of RB is mediated by the GI cyclin/cdk complexes (Buchkovich et al.. 1989; Chen et al., 1989: DeCaprio et al.. 1989). RB is

56


under- or hypo-phosphorylated during early G1 as well as in GO, but becomes hyperphosphorylated during the G1-S transition. Phosphorylation of RB is mediated by the D-type cyclins (cyclin D1, D2, and D3) in association with either cdk4 or cdk6 (Sandhu and Slingerland, 2000), or by the E-type cyclins (cyclin E1 and E2) in association with cdk2 (Ewen et al., 1993; Kato et al., 1993). Inhibition of the cyclin D/cdk4 or cdk6 complexes by the INK4 family (p15, p16, p18, and p19), or inhibition of the cyclin E/cdk2 complexes by the KIP family (p21, p27, and p57) would prevent phosphorylation of RB, thus inhibiting the G1-S cell cycle transition resulting in G1 cell cycle arrest. The phosphorylation state of RB is directly associated with its growth arrest function. The E2F family members are found associated only with the hypo-phosphorylated form of RB, and the transition from G1 into S phase requires 'free' and fully functional E2F. Thus, phosphorylation of RB results in the accumulation of free E2F during late G1, resulting in the transcriptional activation of genes required for entry into S-phase (Morgan, 1995; Sheaff et al., 1.997; Willems et al., 1996). Because only hypo-phosphorylated RB binds to E2F, hyper-phosphorylation of RB is essential for E2F-mediated S phase entry. Hence, the repression of E2F by RB is a critical regulatory mechanism during the G1-S phase transition. D.

Cell cycle checkpoints

Unlike differentiating cells, which terminally arrest their cell cycles from the G1 phase of the cell cycle in a state known as GO, mammalian cells with DNA damage arrest in G1, S, and G2 phases of the cell cycle. Primary among mammalian G1 checkpoint genes is p53. p53 is required for the G1 checkpoint, in which cells arrest in G1 in response to DNA damaging agents such as ionizing irradiation (Kastan et al., 1991). Utilizing cells from knockout mice it has become clear that the p53 gene is critical for G1 arrest following ,/-irradiation (Donehower et al., 1992). In addition, p53 has been shown to be required for maintenance of the G2 arrest following IR (Bunz et al., 1998). Following DNA damage, p53 protein levels are significantly increased in vertebrate cells grown in vitro (Cox et al., 1994; Kastan et al., 1991; Lu and Lane, 1993; Maltzman and Czyzyk, 1984), as well as in vivo (Hall et al., 1993; Midgley et al., 1995). Although p53 mRNA levels do not change in response to DNA damage in many cell systems, the levels of p53 protein increases rapidly. The half life of p53 protein increases substantially after DNA damage (Maki and Howley, 1997; Maltzman and Czyzyk, 1984; Price and Calderwood, 1993), and increased translation of p53 mRNA also contributes to p53 induction (Fu and Benchimol, 1997; Kastan et al., 1991; Mosner et al., 1995). The relative contributions of increased half-life and enhanced translation remain largely undefined. In addition to an increase in the levels of p53 protein, DNA damage is thought to result in activation of p53's ability to bind sequence-specific DNA and consequently transactivate gene expression (Siliciano et al., 1997). Furthermore, the ATM (Ataxia Telangiectasia mutated) gene has been implicated in regulation of p53, because cells from AT patients do not activate p53 normally in response to DNA damage [reviewed in (Elledge, 1996)]. Activation of p53 DNA binding activity is thought to involve phosphorylation of serine 15 of p53

57

The Celt Cycle and Myogenic Differentiation

by ATM (Banin et al.. 1998; Canman et al.. 1998) and by ATR (Canman et al., 1998). In addition, dephosphorylation of p53 by an unknown phosphatase and subsequent association with 14-3-3 proteins leads to an increase in the affinity of p53 for sequencespecific DNA (Waterman et al.. 1998). In response to IR, G1 arrest is mediated at least in part. through induction of the CDK inhibitor p21/WAF1/Cipl by p53 (el-Deity et al., 1993). The p21/WAF1/Cipl gene encodes a protein, which forms a complex with cyclin/cdks, and represents one of the most studied p53 response genes (Elledge and: Harper. 1994: Harper and Elledge. 1996: Ko and Prives, 1996; Sanchez and Elledge, 1995). In normal cells p21 is found associated with a variety of cyclin/cdk complexes, including the G1 cyclins CDK4/Cyclin D (Xiong et al., 1992; Xiong et al.. 1993). In addition, p21 can inhibit the kinase activity of all of the cyclin/cdk complexes (Xiong et al.. 1993). A current model for the G1 checkpoint is illustrated in Figure 1.

Mitogemc signals DNA Dams§e

E2FIDPI Figure l.

.~_ DNA Synthems

Model for DNA damage induced G1 checkpoint. Damage to DNA initiates a cascade of events

which results in the induction of the cyclin kinase inhibitor p21 which in turn inhibits the G1 phase cyclins. Inhibition of the G1 phase cyclins results in accumulation of hypo-phosphorylated and therelore activated RB. Activated RB prevents the G1-S phase transition by binding and inhibiting the S-phase promoting transcription factor E2F/DP1. Once the damage has been repaired MDM2 functions to inhibit RB as welt as p53 and to promote E2F/DP1 function, thus promoting re-entry into the cell cycle.

HI.

MyoD family and :muscle formation

Differentiating muscle cells fuse to form multinucleated myotubes; thereby Withdrawing permanently from the cell cycle. In mice, specification and differentiation of skeletal

58


muscle cells depend on four muscle-specific basic helix-loop-helix transcription factors, the MyoD family of muscle determination genes. Targeted disruption of each family member (MyoD, Myf-5, myogenin, and MRF4) i n mice has demonstrated the importance of these factors for vertebrate myogenesis. Inactivation of Myf-5 results in defects in early myotome formation with presumptive muscle precursors adopting non-muscle cell fates (Braun and Arnold, 1995; Tajbakhsh et al., 1996; Tajbakhsh et al., 1996). However, this early myotomal defect is compensated :for by MyoD, leading to apparently normal muscle at birth. Targeted disruption of MyoD alone also does not dramatically affect muscle specification or differentiation, since these mice have grossly normal muscle (Rudnicki et al , 1992). The functional overlap between Myf-5 and MyoD was demonstrated by generating mice with disruption of both genes, resulting in a complete absence of proliferating myoblasts as well as differentiated muscle fibers (Rudnicki et al., 1993). In contrast, mice with disruption of the myogenin gene contain normal numbers of proliferating myoblasts, but these cells fail to differentiate, indicating that myogenin has a unique role in the transition from determined myoblasts to fully differentiated myotubes (Hasty et al., 1993; Nabeshima et al., 1993). Furthermore, inactivation of the MRF4 gene results in a very mild muscle phenotype, consistent with the notion that MRF4 may have functions that are redundant with the other MyoD family members (Braun and Arnold, 1995; Patapoutian et al., 1995; Zhang et al., 1995). The results of these experiments have led to a simple model for vertebrate skeletal myogenesis that involves the early functioning of Myf-5 and MyoD to establish or determine myoblast cell fate, and myogenin mediating the terminal differentiation of myoblasts. Vertebrate skeletal muscle is derived from cells in the prechordal and somitic mesoderm that give rise to committed myogenic cells of the somite, which eventually become the skeletal muscle of the head, trunk and limbs. The different myofiber and myogenic constituents are thought to be formed from different lineages of myogenic cells (Miller, 1992). tn the mouse, primary myofibers develop first at 8.5 days of gestation, followed by secondary myofibers at day 14. Recent experiments have suggested that MyoD and Myf-5 do have distinct roles for determining these myogenic lineages during development (Kablar et al., 1997). While MyoD(-/-) embryos display normal development of paraspinal and intercostal muscles in the body, proper muscle development in limb buds and brachial arches is delayed by about 2.5 days. By contrast, Myf-5 (-/-) embryos display normal muscle development in limb buds and brachial arches, but markedly delayed development of paraspinal and intercostal muscles. Taken together, these observations strongly support the hypothesis that Myf-5 and MyoD play unique roles in the development of epaxial and hypaxial muscle, respectively. In addition, satellite cells, the muscle stem cells of adult muscle, arise around day 17 as a unique myogenic lineage [for review see (Bischoff, 1994)]. Satellite cells are mitotically quiescent but are induced to enter the cell cycle in response to stress induced by exercise or injury. The activated satellite cells undergo multiple rounds of division before fusing with existing myofibers resulting in repair or hypertrophy, or both. Recently, MyoD has been shown to play a novel role in satellite cell function (Megeney et al., 1996). MyoD (-/-) mice interbred with the mdx mouse (a model for Duchenne and Becker muscular dystrophy) exhibit increased penetrance of the mdx

The Ceil Cycle and Myogenic Differentiation

59

phenotype, characterized by reduced muscle hypertrophy and increased myopathy, This is thought to arise due to a defect in muscle regeneration. Consistent with this hypothesis, the single MyoD (-/-) knockout mice show defects in skeletal muscle regeneration following injury (Megeney et al.. 1996).

IV.

Terminal cell-cycle arrest of myoblasts

During muscle cell differentiation, skeletal myoblasts permanently withdraw from the cell cycle during the G1 phase. At the onset of myoblast differentiation, two events occur simultaneously: activation of myogenic regulatory factors and inhibition of the G1-S phase transition. Activation of the M'~oD family of transcription factors, induction of the p21 gene (one of the KIP family members), inhibition of cdk activity, and hypo-phosphorylation of RB are all necessary for myoblast cell cycle arrest during G1. A large body of evidence demonstrates that direct or indirect interactions between MyoD and these cell cycle regulators leads to terminal cell cycle arrest of myoblasts. A.

Regulation of MyoD activity

The MyoD family of muscle regulatory proteins is transcriptional activators of muscle-specific genes. The MyoD family, MyoD, Myf5, myogenin, and MRF4, belong to the b-HLH class of transcriptional activators and, as discussed above. are thought to play unique roles during muscle differentiation (Braun et al.. 1989: Davis et al.. 1987; Edmonson and Olson. 1989; Rhodes and Konieczny, 1989; Wright et al.o 1989). The b-HLH domain mediates both heterodimerization with ubiquitously expressed bHLH proteins, such as El2 and E47 (E proteins), and binding to specific DNA sequence. CANNTG, known as an E box. which in turn activates muscle-specific gene expression. Although constitutively expressed in proliferating myoblasts. MyoD becomes active only at the onset of myoblast differentiation, suggesting that MyoD activity is highly regulated in proliferating as well as differentiated cells. MvoD activity is regulated at multiple levels. In proliferating myoblasts. MyoD activity is inhibited by direct interaction with negative regulatory factors and by phosphorylation mediated by cell cycle regulatory kinases. During myoblast proliferation. MyoD activity is held in check by interactions with the Id family of proteins (Neuhold and Wold. t993). The Id family of proteins contains HLH domains but lack the basic DNA binding domain, and consequently fail to bind DNA. Therefore. the HLH domain of the Id proteins allows heterodimerization with other HLH proteins, such as the MyoD and E proteins, thus sequestering MyoD and/or E proteins into complexes that fail to bind DNA (Benezra et al., 1990). However, at the onset of differentiation, Id gene expression is down-regulated, resulting in MyoD and E protein dimerization, which leads to activation of muscle-specific genes (Neuhold and Wold. 1993). In addition to negative regulation by td proteins, phosphorylation of MyoD has been shown to inhibit MyoD activity. Site-directed mutagenesis of MyoD revealed that serine residue 200 (Ser200), located within a cdk consensus phosphorylation site. is phosphorylated by cdkl and cdk2 (Kitzmann et al.. 1999). This hyper-phosphorylated

60


form of MyoD has a reduced half-life, resulting from ubiquitin-mediated degradation of MyoD (Song et al., 1998). Set200 phosphorylation of MyoD may also reduce the interaction of MyoD with its associated factors, such as RB (Gu et al., 1993), muscle enhancer factor (MEF)-2 proteins (Molkentin et al., 1995), and the co-activator p300 (Eckner et al., 1996; Puri et al., 1997). In addition, MyoD is highly phosphorylated in growing myoblasts and undergoes substantial dephosphorylation at the onset of muscle differentiation (Kitzmann et al., 1999). In support of this observation, increasing the levels of cyclin D 1 results in an elevation of the phosphorylated form of MyoD, as well as inhibition of terminal myogenic differentiation (Rao et al., 1994; Skapek et al., 1995). Therefore, these data suggest that the unphosphorylated form of MyoD mediates the transition of myoblasts to differentiated myotubes. B.

The role of RB during terminal growth arrest of myoblasts

Genetic studies from patients with retinoblastoma as well as from knockout mice, combined with experiments that elucidate the mechanism of cellular transformation by DNA tumor virus, have defined a central role for the RB family of tumor suppressors in the regulation of the mammalian cell cycle (Sidle et al., 1996). This family of proteins, RB, p107, and p130, regulates the activity of the S-phase transcription factors of the E2F family, which in turn regulate expression of genes required for cell cycle progression. RB is not only a critical regulator of G1 cell cycle progression, but also plays a significant role during muscle differentiation. First, both RB mRNA and protein are dramatically induced during terminal differentiation of mouse skeletal muscle cells (Endo and Goto, 1992), suggesting that the accumulation of RB protein is likely to be required for growth arrest during differentiation of myogenic cells. Second, changes in the RB phosphorylation state, with an accumulation of the growth-suppressing hypo-phosphorylated form of RB, is one of the early events that occurs during muscle differentiation (Gu et al., 1993). It has been demonstrated that most of the RB is hyperphosphorylated in proliferating myoblasts, but rapidly becomes hypo-phosphorylated during muscle differentiation (Gu et al., 1993). As expected, that hypo-phosphorylated form of RB in association with E2F would ensure myoblast G1 cell cycle arrest. RB, in associated with MyoD, is thought to mediate muscle-specific gene expression. In fibroblasts that lack RB, ectopically-expressed MyoD induces an aberrant skeletal muscle differentiation program characterized by normal expression of early differentiation markers, such as myogenin and p21, but attenuated expression of late differentiation markers such as myosin heavy chain (MHC) (Novitch et al., 1996). However, similar defects in MHC expression were not observed in either pl07-deficient or p130-deficient cells, indicating that the defect is specific to the loss of RB. Furthermore, MyoDmediated transactivation of muscle specific genes in SAOS-2 osteosarcoma cells, which lack functional RB, requires co-expression of RB (Gu et al., 1993) or p107 (Schneider et al., 1994), indicating that RB cooperates with MyoD during differentiation. Taken together, these observations indicate that RB mediates myoblast terminal withdrawal from the cell cycle as well as cooperates with MyoD to regulate muscle specific gene expression.


C.

61

Interaction of MyoD with RB mediates terminal cell cycle arrest of myoblasts

The functional interactions between MyoD and RB during muscle differentiation suggest that they act in the same pathway governing the terminal cell cvcle arrest and muscle differentiation. A direct interaction between MyoD and RB was demonstrated by co-immunoprecipitation assays. Gu et al. (1993) reported that MyoD directly and specially binds hypo-phosphorylated RB. in vitro and in vivo. This binding occurs through the bHLH domain of MyoD and the c-terminal half of RB. One potential consequence of this interaction is that bound MyoD may prevent RB from being phosphorylated by cdks during the early stages of muscle differentiation which would potentiate the growth inhibitory role of RB. In addition. MyoD induces RB expression as early as 48 hours after induction of muscle differentiation (Cenciarelli et al., 1999), again reinforcing the RB mediated inhibition of the G1-S phase transition. D.

MvoD-mediated p21 induction during terminal myoblast growth arrest

p21. a member of cell cycle inhibitor Kip family, plays a critical role during cell cycle arrest induced by many forms of cellular stress, including DNA damage, p21 is up-regulated at the level of transcription by the tumor suppresser protein p53 following DNA damage. However, during muscle cell differentiation of myoblasts, p21 induction appears to be mediated by MyoD (Halevy et aI., 1995; Otten et al.. 1997), because forced expression of MyoD in non-muscle cells causes a dramatic increase in the levels of p21 (Cenciarelli et al.. 1999: Halevy et al.. 1995). In addition, induction of the cdk inhibitors p18 and p21 have been observed to couple cell cycle arrest to myogenic differentiation (Franklin and Xiong, 1996; Guo et al., 1995: Halevy et al.. 1995; Missero et al., 1996; Parker et al., 1995). These data indicate that p21 induction during muscle cell differentiation is mediated by MyoD and is associated with muscle differentiation. The increased levels of p21 protein during differentiation of muscle cells is associated with a deCrease in cdk2 activity (Guo et al., 1995: Missero et al., 1996). In vitro, a single p21 monomer is sufficient to inhibit a cyclin/cdk2 complex (Hengst et al., 1998). During the early stages of muscle differentiation, RB assumes a hypo-phosphorylated or active form° the evidence indicates that p21 takes an active role in this process. Indeed. neither p21 nor p16 is able to prevent RB-null myocytes from entering S-phase when ectopically expressed in these cells (Novitch et al.. 1996). Therefore. the role that p21 plays during muscle differentiation is related to its ability to inhibit cyclin/cdk activity, and thus maintain or induce the hypo-phosphorylated state of RB. E

Direct inhibition of cyclin kinase activity by MyoD

In addition to inducing p21 and RB during the early stage of muscle differentiation. MyoD is also involved in the inhibition of cdk activity (Zhang et al., 1999). In vitro, co-immunoprecipitation studies indicate that MyoD can directly interact with cdk4 through a conserved 15 amino acid domain in the C-terminus of MyoD. These studies indicate that the: cdk4-dependent phosphorylation of RB can be inhibited by full-length

62


MyoD, its c-terminus lacking the bHLH domain, or the 15 amino acid cdk4-binding domain (Zhang et al., 1999). Significantly, cell gowth and RB phosphorylation in 10T1/2 mouse fibroblasts is also inhibited by forced expression of full-length MyoD, the c-terminus of MyoD, or the Cdk4 binding domain of MyoD (Zhang et al., 1999). Moreover, the cdk4-binding domain of MyoD fused to GFP can induce differentiation of C2C12 muscle cells in growth medium and the defective myogenic phenotype in MyoD-negative BC3H1 cells can be rescued completely by the cdk4-binding domain of MyoD (Zhang et al., 1999). These studies demonstrate that MyoD can directly inhibit cyclin kinase activity, which would in turn lead to an increase in the hypophosphorylated form of RB and terminal cell cycle arrest of myoblasts. F.

p300/CBP cooperates with MyoD during muscle differentiation

Several lines of evidence suggest that p300/CBP mediate the transacfivation activity of MyoD. First, by using a transient transfection assay, Sartorelli et al. (1997) found that p300 and the related CBP protein increase the ability of MyoD and myogenin to transactivate several muscle-specific promoters and thus promote myogenesis. Second, the results from both physical binding and functional transcription assays indicate that two domains of p300, N-terminal (amino acids 1-596) and C-terminal (amino acids 1572-2370), can act independently to contact and facilitate MyoD activity via the N-terminal activation domain of MyoD. Third, Cenciarelli et al. (1999) have shown that p300 activity is required for MyoD-mediated induction of early differentiation markers such as MyoD itself, myogenin, p21, and RB. Furthermore, acetylation of MyoD by either p300/CBP or pCAF on two lysines, located at the boundary of the MyoD DNAbinding domain, increases MyoD DNA-binding activity. In addition, microinjection of acetylated MyoD increases its transactivation activity on the muscle-specific promoter of muscle creatine kinase. In contrast, MyoD mutants that were not able to be acetylated in vitro by p300 are not activated by p300 in the same functional assay-. These studies indicate that p300/CBP can facilitate MyoD activity through direct acetylation. A current model for muscle cell differentiation is shown in Figure 2.

V.

Maintenance of the differentaited state

Unlike quiescent cells, myotubes cannot be induced to reenter S-phase in response to growth factor stimulation. This permanent withdrawal from the cell cycle is thought to be critical for structure and function of mature muscle. Although the mechanisms preventing myotubes from reinitiating DNA synthesis are still unclear, there are several studies that indicate that RB, p21, and cyclinD3 are involved. A.

A role for RB and p21 for maintaining G1 arrest

An active role for RB during terminal cell cycle arrest of myoblasts has been described above. However, a large body of evidence suggests that RB also functions to prohibit the replication of DNA in differentiated muscle cells. Differentiated myotubes, with


63

Mitogenic signals

Cyclin Dependent Kinases

p')l _

J/

p300/CBP/PCAF

E2FIDPI

~

DNA Synthesis

Figure 2. Model for muscle cell differentiation. Mitogenic signals function to promote G1-S phase progression by stimulating expression of the GI phase cyclins, which in turn inhibit RB and MyoD activities. During MyoD mediated cellular differentiation MyoD induces expresmon of the cyclin kinase inhibitor p2I and inhibits the G1 phase cyclin dependent kinases directly. Inhibition of the G1 phase cyclin kinases results m accumulationof hypo-phosphorylatedand activated RB. which in mrn inhibits the S-phasepromoting factor E2F/DP1. thus promoting G1 cell cycle arrest and the activation of muscle-specific genes.

nonfunctional RB, are capable of reinitiating D N A synthesis (Gu et al., 1993: Novitch et al.. 1996; Schneider et al., 1994). In addition, viral proteins such as SV40 large T antigen and adenoviral E1A, both of which share the ability to bind and inactivate RB, can act alone in inducing D N A synthesis in differentiated muscle cells. In addition. myocytes lacking p21 can synthesize D N A indicating that p21 is also involved in preventing DNA synthesis in differentiated muscle cells (Andres and Walsh. 1996). Indeed, it was found that reactive cyclin E-cdk2 complexes bound to p21 persists m differentiated muscte cells, and purified E1A, targeting an inhibitory activity of p21, restores cdk2-associated kinase activity in differentiated myotubes, thus inducing DNA synthesis in these cells (Mal et al., 2000). Moreover. restoration of kinase activity m differentiated muscle cells by E1A leads to phosphorylation o f RB. These observations suggest that both p21 and RB are involved in preventing differentiated muscle cells from reinitiating D N A synthesis and that RB is functioning downstream o f p21. B.

A role for cyclin D3 in preventing D N A synthesis in myotubes

Binding of cyclin with cdk is required for cdk activity, which is essential for cell cycle progression. There are at least nine different types o f cyctins discovered so far. Different cyclins bind and activate specific cdks in a cell cycle-dependent manner (Sandhu and Slingerland, 2000). The progression from G1 to S is promoted by a group of G1 cyclins, the D-type cyclins functioning in mid G1. and the E-type cyclins

64

J, Huang and M. J. Thayer

functioning in late G1 (Pestell et al., 2000). As expected, the expression of cyclins in muscle cells is down-regulated at the onset of terminal differentiation, as cells arrest in the G0/G1 phase of the cell cycle (Kiess et al., 1995; Rao and Kohtz, 1995; Wang and Walsh, 1996), with the exception of cyclin D3, whose expression is induced during terminal differentiation (Cenciarelli et al., 1999; Kiess et al., 1995; Rao and Kohtz, 1995). By using a hormone-activated MyoD cell line, cyclin D3 was induced as well as stabilized following MyoD expression (Cenciarelli et al., 1999). Furthermore, Cenciarelli et al. (1996) found that the ability of microinjected E1A to reinitiate DNA synthesis in terminally differentiated myotubes is counteracted by cyclin D3 co-injection. Moreover, cyclin D3 is found in inactive complexes with cdk4, cdk2 and PCNA (Cenciarelli et al., 1999; Kiess et al., 1995; Riley et al., 1994). Therefore, the mechanism by which cyclin D3 prevents differentiated muscle cells from reinitiating DNA synthesis is likely due to the ability of cyclin D3 to trap proliferating cell nuclear antigen (PCNA), a positive regulator for DNA synthesis, into inactive complexes with cdk, cdk2, and RB.

VI.

A role for M D M 2 and ATR in muscle tumor formation

Rhabdomyosarcomas are skeletal muscle tumors and are one of the more common solid tumors of childhood, representing 4-8% of all malignancies in humans under 15 years of age. Tumors arise de novo from skeletal muscle and are grouped by histologic and cytogenetic criteria as either emhryonal or alveolar rhabdomyosarcomas. A balanced translocation between chromosomes 2 and 13, t(2:13) (q35;q14), is associated with alveolar rhabdomyosarcomas (Barr et al., 1993). The PAX3 gene has been shown to be fused to a forkhead gene family member (FKHR) in the t(2:13) translocation (Barr et al., 1993; Shapiro et al., 1993). In some cases of alveolar rhabdomyosarcoma, the PAX7 gene, located at lp36, has been shown to be translocated to the FKHR gene as a 1:13 translocation (p36;q14) in some cases of alveolar rhabdomyosarcoma (Davis et al., 1994). Loss of heterozygosity on the short arm of chromosome 11 encompassing llp15 is associated not only with embryonal rhabdomyosarcomas (Scrable et al., 1990), but also with several other solid tumors (Newsham et al., 1991), suggesting the location of one or more tumor suppressor genes for multiple tumor types in this region. In addition, gene amplifications have been observed in both embryonal and alveolar rhabdomyosarcomas. Utilizing Comparative Genomic Hybridization (CGH) on primary alveolar rhabdomyosarcomas, the most frequent amplicons have been localized to 2p24 and 12q13-14, with both amplifications occurring in 4 out of 10 tumors (Weber-Hall et al., 1996). The 2p24 amplicon had previously been shown to involve the MYCN gene (Dias et al., 1990), while the genes involved in the 12q13-14 amplicon have not yet been fully defined. Two distinct chromosome 12q13-14 amplification units have been described in other types of sarcomas (Suijkerbuijk et al., 1994) as well as in gliomas (Reifenberger et al., 1994). Mapping of these two ampticons implicates MDM2 or CDK4 and SAS as likely targets of the amplification events (Reifenberger et al:, 1994). The frequency of these two different amplicons in primary rhabdomyosarcomas is currently unknown.


A.

65

Amplification of MDM2

One obvious phenotype of tumor cells is a lack of terminal differentiation. We have conducted a series of somatic cell genetic experiments designed to identify genetic loci present in rhabdomyosarcoma cell lines that are capable of inhibiting muscle differentiation (Fiddler et al.. 1996; Smith et al., 1998; Tapscott et al., 1993). Initially, we showed that rhabdonayosarcomas could be classified as either dominant or recessive with respect to MyoD activity and terminal differentiation (Tapscott et al.0 1993). Subsequent analysis of two different dominant types of minors indicated thin the loss of differentiation could be mapped to individual loci. First, microcell mediated chromosome transfer of a derivative chromosome 14 from the rhabdomyosarcoma cell line Rhl8 into the differentiation competent myoblast cell line C2C12 inhibits muscle differentiation and the ability of MyoD to transacfivate reporter constructs. The derivative chromosome 14 contains a region of amplified DNA originating from chromosome 12q13-14, and contains several genes often amplified in sarcomas (Fiddler et al.. 1996). Testing the amplified genes for the ability to inhibit muscle-specific gene expression indicated that forced expression of MDM2 inhibits MyoD function, and consequently inhibits muscle differentiation. Thus. amplification and over-expression of MDM2 in rhabdomyosarcomas inhibits MyoD function, resulting in dominant inhibitior~ of muscle differentiation (Fiddler et al., 1996). The oncogenic properties of MDM2 have been postulated to result from direct interaction with several cell cycle regulatory proteins. MDM2 interacts directly with p53 (Oliner et al., 1992), and blocks p53 mediated transactivation by inhibiting the activation domain of p53 (Chen et al.. 1993: Haines et al., i994; Momand et a12 1992: Oliner et: al.. 1993; Wu et al., 1993; Zanherman et al.. 1993). In addition, MDM2 interacts with the activation domain of E2F1, resulting in stimulation of E2F1/DP1 transcriptional activity (Martin et al.. 1995). Furthermore. MDM2 has been shown to interact directly with RB, resulting in stimulation of E2F/DP1 transcriptional activity and inhibition:of RB growth regulatory function (Xiao et al., 1995 I. Taken together. these results suggest that MDM2 not only relieves the proliferative block mediated by either p53 or RB. but also promotes proliferation by stimulating the S-phase-inducing transcriptional activity of E2F/DP1. Thus, the most obvious role for amplification of MDM2 in tumorigenesis would be to inactivate p53 or RB or both. In addition, we have shown that amplification of MDM2 inhibits MyoD activity and consequently inhibits normal muscle differentiation (Fiddler et al., 1996). These studies have identified a previously un-described activity for the MDM2 protein, and broadens the role of MDM2 in cell growth control to include inhibition of differentiation. Because MDM2 is expressed in proliferating myoblasts, and MDM2 can inhibit muscle differentiation (Fiddler et al., 1996), it is possible that MDM2 inhibits differentiation of myoblasts during cellular proliferation. Furthermore, activation of muscle-specific genes to high levels reqmres members of the RB family ~Gu et aL, 1993: Novitch et al.. 1996; Schneider et al.. 1994). During muscle differentiation. RB expression increases (Coppola et al.. 1990: Endo and Goto. 1992: MarteUi et al.. 1994) and assumes a hypo~phosphorylated, activated state (Gu et al.. I993: Thorburn et al., 1993). Because MDM2 interacts with RB and inhibits the growth regulatory functions of RB

66


(Xiao et al., 1995), it is possible that RB in turn regulates the growth-stimulatory effects of MDM2. Consistent with this possibility, we find that forced expression of RB can restore MyoD activity in cells with amplified MDM2 (unpublished observations). Because RB and MDM2 interact, these results suggest that an additional role for RB during muscle cell differentiation is to bind and inactivate MDM2. B.

Duplication of ATR

In a second screen for inhibitors of differentiation, we found that microcell mediated chromosome transfer of an isochromosome 3q [i(3q)], from a different rhabdomyosarcoma cell line, into the differentiation competent myoblast cell line C2C12 also inhibits differentiation. I n addition to inhibition of differentiation, the i(3q) causes abnormal centrosome amplification, resulting in aneuploidy, and abolishes G1 arrest following DNA damage (Smith et al., 1998). We have extended these observations by showing that forced expression of ATR (ataxia-telangiectasia and rad3 related; located at 3q24) results in a phenocopy of the i(3q) containing hybrids. Itis possible that these findings could have implications for non-muscle tumors as well. CGH indicates that 3q is a hotspot for increased DNA copy number in several different types of cancers. In head and neck squamous cell carcinoma (HNSCC), the most frequently observed increase in DNA copy number was from chromosome 3q, occurring in 10 of 13 primary tumors (Speicher et al., 1995). Karyotypic analysis of HNSCC cells showed the presence of an i(3q) in 30-40% of tumors (Carey et al., 1993; Jin et al., 1993; Rao et al., 1994). Similarly, the most frequent increase in DNA copy number in primary small cell lung carcinomas was also on 3q, occurring in 10 out of 13 cases (Ried et al., 1994). Furthermore, the formation of an i(3q) has been observed in small cell lung carcinomas (Rabbitts et al., 1990). Prior to our study, the most compelling argument for an involvement of 3q in tumorigenesis came from analysis of cervical carcinomas (Heselmeyer et al., 1996). Gain of 3q is the most consistent chromosomal aberration in cervical carcinomas and was present in 9 out of 10 tumors. Furthermore, this alteration occurs during the progression from severe :dysplasia to invasive carcinoma (Heselmeyer et al., 1996). Interestingly, the presence of the gain of 3q correlated with the conversion of tetraploid dysplastic cells to aneuploid carcinoma cells. While it remains possible that different genetic alterations of 3q occur in different tumors, our data indicate that the i(3q) present in Rh30 rhabdomyosarcoma cells induces aneuploidy. Therefore, the aneuploidy observed in cervical carcinoma cells may be a direct result of increased copy number on 3q resulting in ATR over-expression. Taken together, these studies implicate alterations in the ATR locus as causing loss of differentiation and cell cycle abnormalities in several different types of tumors.

67


VII.

Cell cycle checkpoints and differentiation

Damage to genomic DNA occurs spontaneously in all living ceils, and represents a significant and constant problem. In addition, chemical or physical mutagens can cause a variety of DNA lesions, including base modifications, intrastrand and interstrand crosslinks as well as single or double strand breaks (Friedberg et at., 19951. If left unrepaired, these DNA lesions can lead to mutations or loss of viability. Thus, cell cycle checkpoints and DNA repair mechanisms have evolved to ensure cellular survival in the face of DNA damage. Multicellular organisms have additional issues to deal with. including differentiation programs, as well as the existence of a limited number of stem ceils required for renewal and repair of differentiated tissues. Because cell cycle checkpoints and differentiation utilize the same key cell cycle regu.latory factors to mediate cell cycle arrest, multicellular organism must integrate these two processes simultaneously. Because both cell cycle checkpoints and muscle differentiation use the same cyclin kinase inhibitor, p21, to arrest the cell cycle, and because forced expression of p21 in muscle cells leads to premature differentiation (Halevy et at., 1995). a mechanism to protect the myoblast population from premature differentiation induced b y DNA damage must exist. We propose that the inhibitory interactions between MDM2 and MyoD and between ATR and MyoD fulfill this role. so that if a myoblast sustains DNA damage, premature differentiation dose not occur. A model to integrate these two processes is shown in Figure 3.

Mitogenic slgnals

',i,°°

I

~

~

E2F/DPI

~--~ DNA Synthesis

elffebentiat~n

Figure 3. :Model for integration of the DNA damage response and the muscle cell differentiation:pathway, If proliferating muscle cells incur DNA damage, the increase in p21 expression would potentiate M y o D activity and promote differentiation. Therefore, we propose the DNA damage response has evolved to inactivate MyoD during the G1 phase checkpoint. This would allow myoblasts that experience DNA damage to re-enter the cell cycle; thus protecting the myoblast population from premature differentiation.

68

VIII.


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CHAPTER 4

FIBER TYPE SPECIFICATION IN VERTEBRATE SKELETAL MUSCLE

STEFANO SCHIAFFINO, C A R L O R E G G I A N I and G E E R T R U I J TE K R O N N I E


Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Signals responsible for muscle fiber type specification during embryonic development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Growth factors and hormones affecting muscle fiber type specification . . . . . . . . . . . . . IV. Muscle fiber type specification by innervation and contractile activity. . . . . . . . . . . . . A. Triggering signals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Transduction pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Transcription factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Conclusions and open issues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

I.

75 75 79 81 82 84 86 87 89 89

Introduction

The diversity of skeletal muscles has long been recognized and a large number of studies during the last 40 years have described in detail the biochemical, structural and functional properties of the various muscle fiber types, as well as the role o f myoblast predetermination, innervation and hormones in modulating these properties (see Bottinetfi and Reggiani, 2000; Gunning and Hardeman, 1991: Hughes and Salinas. 1999: Pette and Staron. 1997; Schiaffino and Reggiani, 1996; Stockdale. 1992). However, a search for the signals, transduction pathways and regulatory genes responsible for muscle cell diversification has started only during the last few years. Progress in this field will be briefly reviewed here with specific reference to vertebrate skeletal muscle.

II.

Signals responsible for muscle fiber type specification during embryonic development

In vertebrates, muscle cell specification starts early in development with the emergence of distinct populations o f myogenic precursors. Recent work in zebrafish and chick 75 Advances in Developmental Biology and Biochemistry Ed. David Sassoon 75 © 2002 Elsevier Science. Printed in the Netherlands.

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S. Schiaffino, C. Reggiani and G. te Kronnie

embryos suggests that some of the signals from the notochord and neural tube that are known to influence myogenesis in the somites may also directly affect the development of specific fiber types. Zebrafish embryos have three distinct muscle fiber types: muscle pioneer slow muscle fibers, non-pioneer slow muscle fibers and fast muscle fibers (see Stickney et al., 2000). Slow muscle fibers develop from adaxial cells, which are adjacent to the notochord in the segmental plate. These cells are the first to express MyoD and specific MyHCs (see Raamsdonk et al. 1978; Devoto et al., 1996). Lineage studies have demonstrated that after formation of t h e somite borders, adaxial cells begin to elongate and then migrate radially to the surface of the somite where they form a layer of embryonic slow muscle fibers (Devoto et al., 1996). A subset of these fibers express Engrailed protein and are therefore named pioneer muscle cell in analogy to pioneers in Drosophila. After adaxial cell migration to the surface of the myotome, fast muscle fibers develop from ceils that are initially lateral to the adaxial cells in the segmental plate. The notochord-derived members of the Hedgehog family of secreted proteins play a central role in muscle cell differentiation. In particular, Sonic hedgehog (Shh) signaling is required for the early formation of adaxial cells and for the subdivision of adaxial cells into slow fibers and slow fiber pioneers. Zebrafish mutants lacking Shh expression fail to form slow muscle but do form fast muscle, while overexpression of Shh is sufficient to induce slow muscle cells in the paraxial mesoderm and ectopic expression of Shh leads to formation of slow at the expense of fast muscle (Blagden et al., 1997; Duet al., 1997). The role of Shh has been confirmed using an in vitro culture system in which Shh was found to induce the slow phenotype in embryonic zebrafish myoblasts (Norris et al., 2000). In Drosophila, transduction of the Hedgehog signal is closely associated with the product of the segment polarity gene Patched (Ptc), a multipass membrane-spanning protein, and with the activity of cAMP-dependent protein kinase A (PKA). This appears to be true also in zebrafish. Mis-expression of Shh results in ectopic activation of Ptc-1 transcription and expression of Ptc-1 is regulated by protein kinase A activity, suggesting that the mechanism of signalling by Hedgehog family proteins has been highly conserved during evolution (Concordet et al., t996). A recently identified mutant slow-muscle-omitted (Smu) seems also to act downstream Hh. Mutations in the zebrafish Smu gene, which like the Ptc gene, encodes a multipass membrane-spanning protein, disrupt Hedgehog signaling: Smu (-/-) embryos have a 99% reduction in the number of slow muscle fibers and a complete loss of Engrailedexpressing muscle pioneers (Barresi et al., 2000). An emerging view is that Ptc and Smu act as subunits of a putative Hh receptor complex. In this model, Smu acts as the transducing subunit, and can be blocked by a direct interaction with the ligandbinding subunit, Ptcl Thus, when Hh binds to Ptc, Smu is released from Ptc-mediated inhibition and activates the intracellular signaling pathway which is subjected to negative regulation by PKA. Overexpression of dominant negative PKA (dnPKA) in wild-type embryos causes all somitic cells to develop into slow muscle fibers. Overexpression of Shh does not rescue slow muscle fiber development in Smu (-/-) embryos, whereas overexpression of dnPKA does (Barresi et al., 2000). The zinc finger

Zebrafish.

Fiber Type Specification in Vertebrate Skeletal Muscle

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transcription factors of the Gli family (Glil, Gli2 and Gli3) are maior final effectors of Shh signaling in vertebrates and mutations, affecting the zebrafish Gli2 gene, interfere with somite formation and patterning during embryogenesis (Karlstrom et al., 1999; van Eeden et al.. 1996). The dissection of Shh signaling pathways is complicated by the fact that the zebrafish, like all other vertebrates, have multiple family member of most genes involved in Hedgehog signaling, e.g. three Hh genes and two Ptc genes. with overlapping expression patterns and redundancy between different homologues. Thus, the loss of activity of one signal, Shh, can be partially compensated for by other Hedgehog family proteins (Lewis et al.. 1999). Furthermore, other signals such as those mediated by members of the TGF-beta and Wnt families are involved in zebrafish muscle cell specification. For example, ectopic expression of Dorsalin-1, a member of the TGF-beta superfamily, in the notochord inhibits the formation of muscle pioneer cells, demonstrating that TGF-beta signals can antagonize the induction of muscle pioneer cells by Hedgehog (Du et al., 1997). Chick. The notochord and neural tube are crucial in patterning myogenic cell lineages during avian and mammalian somitic myogenesis. Studies based on bead implantation and antisense inhibition experiments in chick embryos show that that Shh is both sufficient and necessary for MyoD activation in somites and that genes of the Shh signal response pathway, such as Ptc and Gli, are activated in coordination with somite formation (Borycki et at., 2000). The notochord and neural tube not only promote myogenesis but also determine the slow phenotype of the primary fibers and purified Shh protein mimics this action in the chick embryo (Cann et at., t999). Newly formed somites placed in organ culture predominantly form primary fibers that express slow myosin in response to Shh and this effect can be blocked by disrupting the Shh signaling pathway through increased activity of somitic PKA. It wilt be of interest to determine whether Shh signaling is also involved in the cell lineage determination of limb myoblasts that originate from somites and migrate into the periphery during limb bud formation. These Pax3 positive precursors of the hypaxial musculature that derive from the ventro-lateral portion of the dermomyotome do not express MyoD and Myf5 genes untill they have reached the limb bud. However, elegant in vivo surgical transplantation studies in chick embryos indicate that limb myoblast diversity arises prior to the entry of myoblasts into the limb (Van Swearingen and Lance-Jones, 1995). When mvoblast migration is interrupted by transplanting limb bud tissue to the coelomic cavity of a host embryo early in the migratory period, the muscles form a majority of slow myotubes. In contrast, in limbs transplanted at slightly later stages, only muscles that normally contain the highest numbers of fast myotubes were missing. Parallel results were obtained in chimeric limbs made by transplanting a quail limb bud to a chick host at different times during the migratory period. These findings suggest that the earliest myoblast migrants give rise mainly to slow primary myotubes whereas the later migrants give rise to fast myotubes. A significant depletion of stow myosin-positive profiles was found within slow muscles after transplantation of young limb bud tissue to older hosts, suggesting that the fate of migrating myoblas~s is not defined by the early limb bud environment. Thus, it appears that limb myoblaSt diversification o f fast and slow muscle cell lineages arises in the somites or early in the migratory period.

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On the other hand, muscle patterning in the developing limb is localised within the limb mesenchyme and not in the somitic myogenic precursor cells themselves and Shh is known to regulate the patterning of anterior/posterior muscles (Duprez et al., 1999). Ectopic application of a localised source of Shh to the anterior margin of the wing is able to transform anterior forearm muscles into muscles with a posterior identity and to induce a mirror image duplication of the normal posterior muscles fiber types in the new posterior muscles. Interestingly, the reorganisation of the slow fibers can be detected before muscle mass cleavage started, suggesting that the appropriate fiber type arrangement is in place before the splitting process can be observed. Two members of the Gli family from chick, called Gli and Gli3, appear to be differentially regulated by Shh in the developing chick limb bud: Shh up-regulates Gli transcription, while down-regulating Gli3 expression (Marigo et al., 1996). The existence of distinct fast and slow myogenic cell lineages in the developing avian embryo has been clearly established by the classical studies of Stockdale and coworkers (see Stockdale. 1992). The crucial experiment was the demonstration that clones of cultured muscle cells expressing either fast and slow myosin heavy chain (MyHC) or exclusively fast-type MyHC can be derived from early chick embryos. Muscle cells grown from embryos at later stages of development tend to form predominantly fast MyHC. Motor nerves are not required initially for the differentiation of fast and slow muscle fibers, although innervation is subsequently essential for muscle growth and survival (Butler et al., 1982). Motor innervation is also involved in fiber type differentiation during fetal development. Muscle ceils from fetal slow muscle express slow myosin only when co-cultured with neural tube, whereas muscle cells from fetal fast muscle do not express slow myosin even when co-cultured with neural tube, showing that the formation of slow muscle fiber types is dependent on both myoblast lineage and innervation (DiMario and Stockdale, 1997). Satellite cell populations also differ in adult chick and quail muscles (Feldman and Stockdale, 1991). Primary satellite cells isolated from fast muscles formed only fast fibers in culture, while up to 25% of those isolated from slow muscles formed fibers that expressed both fast and slow MyHCs. Different populations of myogenic precursors showing differential expression of the transcription factors, Myf-5 and MyoD, have been identified in the mouse embryo, however there is no evidence that these cells correspond to distinct myogenic lineages nor, for that matter, that they are related to the precursors of fast and slow muscle cells (Tajbakhsh and Buckingham, 2000). Mammalian myogenic cells grown from early and late embryonic stages display different properties in culture, including differential expression of slow myosin, which is only found in cultures from embryonic myoblasts (Vivarelli et al., 1988). Heterogeneity of myogenic precursors in term of slow MyHC expression was detected in the somitic myotome of E l l . 5 rat embryos (Dhoot, 1994). There are contrasting results as to whether satellite cells from mammalian muscles show differential expression of slow myosin (Dusterhoft and Pette, 1993; Edom et al., 1994). Retroviral labeling of myoblasts clones showed that rodent myoblasts labelled at early embryonic stages (El5) contributed to primary fibers only, the majority of which expressed slow myosin (Dunglison et al., 1999). In contrast, myoblasts labelled

Mammals.


79

at fetal or early postnatal stages fused with both fast and stow fibers (Dunglison et al.. 1999; Hughes and Blau, 1992). Muscle fiber types can differentiate in the absence of the nervous system in rat embryos, supporting the notion of intrinsic differences between myogenic precursors (Condon et al.. 1990). At present, there is no direct evidence about a role of Shh signaling on the diversification of fast and slow muscle cells in the mammalian embryo.

HI.

Growth factors and hormones affecting muscle fiber type specification

Muscle growth and fiber type specification are modulated by growth factors and hormones during development and in the adult. We will focus here on thyroid hormones and insulin-like growth factors 1 (IGF-1), that have been more intensively investigated in relation to signaling mechanisms affecting fiber type specification. Hyperthyroidism causes a slow-to-fast fiber type conversion in skeletal muscle. whereas hypothyroidism causes a fast-to-slow conversion (see Muscat et al., 1995: Pette and Staron. 1997). In the rat, the lack of thyroid hormones during the first weeks of postnatal development causes a delay in the transition from the immature fiber phenotype, characterized by the presence of embryonic and neonatal MyHCs. to the adult fiber phenotype, characterized by the presence of the fast-type MyHC-2A, -2X and -2B, and affects especially the appearance of MyHC-2B (Muscat et al., 1995). In the slow soleus muscle, hypothyroidism delays the disappearance of neonatal MyHC. reduces the expression of fast MyHC-2A and increases the expression of MyHC-slow (Adams et al.. 1999). In the fast plantaris muscle hypothyroidism inhibits both the decrease of neonatal MHC and the expression of fast MyHC-2X and -2B (Adams et al.. 1999). The effect on the transition from neonatal to fast 2B fibers is particularly evident: at the age of one month the plantaris of hypothyroid rats is virtually devoided of type 2B fibers and still dominated by neonatal fibers (Adams et aL, 1999). The effects of thyroid hormones are mediated by different thyroid receptors (TRs) and recent studies show that TRc~I-, TR[~-, or TRal/~-deficient mice show changes in fiber type profile and MyHC isoform expression resembling those found in hypothyroid animals (Yu et al., 2000). However, the absence of embryonic and fetal MyHC isoforms in the TR-deficient mice indicates that the transition from developmental to adult MyHC isoforms is not solely mediated by TRs. Thyroid hormones might also affect indirectly the muscle phenotype. For example, beta-adrenergic receptor density in skeletal muscle fibers is increased by hyperthyroidism (Martin et al., 1992), and beta adreno-receptor activation reduces the expression of the slow phenotype and increases the expression of the fast phenotype including a shift from aerobic to anaerobic metabolism IPolta et al.. 2001: Zeman et al., 1988). Thyroid deficiency also depresses plasma IGF-I levels (Adams et al.. 2000b). Several growth factors, including fibroblast growth factors and insulinqike growth factors, are essential for muscle cell proliferation and differentiation both in vitro and in vivo (Hauschka, 1994). Some of these growth factors, such ad IGF-1, are also produced by muscle cells themselves and appear to affect muscle development through autocrine/paracrine mechanisms. Body growth and muscle growth are dependent on

80


growth hormone (GH) and IGF-1 and most cells, including muscle cells, contain receptors for both growth factors. The relative role of the two factors has been recently investigated using transgenic mice with mutations in GH, IGF-1 and their receptors. Whereas the lack of GH is compatible with life and causes pituitary dwarfism in both mice and humans, mice bearing a null mutation for the IGF-1 gene have a high neonatal mortality rate and marked body growth and muscle growth retardation, and mice bearing a null mutation for the IGF-1 receptor gene invariably die shortly after birth (see Le Roith et at.. 2001). The finding that surviving IGF-1 null mice do not display accelerated growth after injection of GH during postnatal development suggests that the effect of GH on postnatal body growth is mostly mediated by IGF-1. On the other hand, the growth retardation of mouse double mutants lacking GH receptor and IGF-1 is more severe than that observed with either class of single mutant, showing that GH and IGF-1 promote postnatal growth by both independent and common functions (Lupu et al., 2001). The original "somatomedin hypothesis" proposed that circulating IGF-1 produced by the liver, the primary site of IGF-1 production, mediate the effects of GH on various tissues. However, in transgeuic mice that lack IGF-1 specifically in the liver, no defect in growth or development was detected although circulating and serum levels of IGF-1 were decreased by approximately 75% (Le Roith et al., 2001; Sjogren et al., 1999; Yakar et al., 1999). These results can be explained by assuming that local production of IGF-1, acting in a paracrine/autocrine fashion, mediates GH-induced somatic growth. Muscle fibers can indeed produce IGF-1 and muscle-specific splice variants of IGF-1 have been identified (Goldspink, 1999; McKoy et al., 1999; Musaro et al.. 2001: Yang et al., 1996). The effect of overexpression of these variants in muscle cells has been the object of recent investigations. Viral mediated expression of IGF-1 in muscle has been found to induce muscle hypertrophy in young animals and prevent muscle atrophy and loss of the fastest, most powerful type 2B fiber types seen in old muscles (Barton-Davis et al., 1998). Injecting rat muscle with a plasmid encoding IGF-1 causes a switch to glycolytic metabolism (Semsarian et al.. 1999). Transgenic mice with a rat muscle-specific IGF-1 cDNA driven by a fast muscle promoter show marked muscle hypertrophy but essentially unaltered distribution of slow and fast fibers, with a tendency toward faster fiber types (Musaro et al.. 2001). Which are the intracellular signalling pathways activated by IGF-1 in muscle cells? There is evidence that the PI3K-Akt acts as a major downstream mediator of IGF signaling (see Lawlor and Rotwein, 2000). On the other hand. IGF-1 has also been reported to induce hypertrophy of cultured muscle cell through calcineurin-mediated signalling (Musaro et al., 1999" Semsarian et al.. 1999). It is not clear whether this is true also in vivo, because the calcineurin inhibitor cyclosporin A does not prevent muscle hypertrophy in transgenic mice with a rat muscle-specific IGF-1 cDNA driven by a muscle-specific promoter (Musaro et al., 2001). Indeed, calcineurin has been implicated in the differentiation of slow fibers (see below), a finding not easily reconciled with the tendency for IGF-1 to induce a shift towards a fast-glycolytic phenotype (Semsarian et al., 1999).


IV.

81

Muscle fiber type specification by innervation and contractile activity

Skeletal muscle is dependent on innervation since early stages of development. Though many primary generation fibers can form and differentiate into fast and slow lineages even in the absence of functional innervation, the survival of most primm'y fibers and the formation of most secondary generation fibers is dependent on innervation. Motor innervation is essential for the maturation of the muscle fibers during postnatal development and for the maintenance of fiber type specificity throughout life. The matching of diverse populations of motoneurons to their appropriate muscle targets has been recently reviewed (Hughes and Salinas. 1999). Cross-reinnervation experiments have clearly shown that motor neurons can re-specify the gene programs of the muscle fibers. However, the re-specification is often incomplete, possibly reflecting intrinsic differences between fiber types. Motor neurons can affect the muscle fiber phenotype through different types of signals, including signals that act locally at the neuromuscular junction, such as agrm or neuregulin, and signals that act throughout the muscle fiber, such as specific trains of action potentials. A special case of local signaling is represented by the effect of sensory nerve terminals on spindle muscle fibers (intrafusal fibers). In contrast with the rote played by efferent (motor~ innervation which drives muscle fiber growth and differentiation, afferent (sensory) innervation of muscle spindles seems to block fiber growth and differentiation. Intrafusal fibers, which receive afferent as well as efferent innervation, retain several immature features such as small size. expression of neonatal myosin and lack of adult-type excitation-contraction coupling (see Watro and Kucera, 19991. Recent work on mice with knock-out of neurotrophin 3 (Ernfors et al., 1994: Farinas et al., 1994; Wright et ah. 1997), a muscle-derived growth factor required for the growth and survival of afferent nerve fibers, has brought support to earlier evidence that sensory innervation is essential for the formation of muscle spindles (Walro and Kucera. 1999). The differential distribution of myosin isoforms, with neonatal MyHC being present in the equatorial region and adult type MyHCs in the polar regions, appears to reflect the action of afferent innervation in the equatorial region and efferent innervation in the polar regions (Walro and Kucera. 1999). On the other hand. developing muscle fibers destined to become intrafusal fibers can be identified before the contact with sensory nerve endings by the expression of Egr3, a muscle transcription factor that has been shown to be essential for intrafusal fiber formation (Tourtellotte and Milbrandt, 1998). Therefore the differentiation of intrafusal muscle fibers is apparently dependent on both intrinsic mechanisms, mediated by unknown signals prior to innervation, and extrinsic mechanisms, related to sensory innervation. The contractile activity mediated by motor nerves plays a major role in muscte fiber type specification. This has been clearly shown by electrostimutation experiments. including direct electrostimulation of denervated muscles (Pette and Vrbova. 1992~.. A major open issue is how the information encoded in the activity patterns of motor neurons is translated into fiber type-specific transcriptional programs. Surface excitation is coupled to contraction of muscle fibers and contraction is in turn coupled to metabolic activation. However, the relative role of electrical, mechanical and metabolic signaling on muscle gene regulation remains to be established. The transduction pathways

82


activated by these signals and the transcription factors that act as final effectors of the signaling cascades are also poorly defined. A.

Triggering signals

Electrical activation and Ca 2+ signaling. The effect of electrical signals on the muscle phenotype has been extensively investigated by electrostimulation via nerves or directly with electrodes implanted onto the denervated muscles. Various impulse patterns, resembling the firing patterns of fast and slow motor neurons, have been shown to affect differentially the muscle fiber type profile (Pette and Vrbova, 1992). However, it is not clear how the muscle cell is able to decode the information contained in specific depolarization patterns. Voltage-dependent ionic channels could act as sensors of electrical signals and intracellular Ca 2+ changes are known to mediate the effect of membrane depolarization patterns on gene regulation in different cell systems (Buonanno and Fields, 1999). Variations in the amplitude, frequency and compartmentalization of calcium signals are decoded by calcium/calmodulin-dependent enzymes, such as calcineurin and CaMK (see below) and thus regulate gene expression in cardiac and skeletal muscle (Frey et al.. 2000a; Olson and Williams, 2000). Free calcium is higher in slow-twitch compared to fast-twitch skeletal muscle fibers (Carroll et al., 1997) and changes in resting calcium concentration were detected after continuos low frequency stimulation of rabbit (Sreter et al., 1987) and rat muscles (Everts et at.. 1993). A more detailed analysis has been recently carried out on isolated muscle fibers using sensitive calcium indicators (Carroll et al., 1999). Low frequency stimulation of fast muscles from hypothyroid rats was found to lead to an increase in resting free calcium that is significant as early as 12 h after beginning of stimulation, attains a 2.5-fold elevation at 1 day and persists in this range after 10 days. A direct role for Ca 2+ changes in inducing a slow muscle-specific gene program is supported by studies in cultured muscle cells (Kubis et al., 1997; Meissner et al., 2000). Primary cultures from newborn rabbit muscle were grown on microcarriers in suspension and acquired after several weeks an adult pattern of fast myosin light and heavy chains. When Ca 2+ ionophore was added to the culture medium, raising intracellular Ca 2+ about 10-fold, the myotubes switched to a slow muscle gene program, expressing slow myosin light and heavy chains, elevated citrate synthase, and reduced lactate dehydrogenase. This effect was reversed after withdrawal of the Ca 2+ ionophore. Mechanical signals. The role of mechanical signals on muscle gene regulation and fiber type specification is illustrated by different experimental models, such as mechanical overloading produced by elimination of synergistic muscles or unloading produced by hindlimb suspension or microgravity (Pette and Staron, 1997; Thomason and Booth, 1990). Overloading is known to induce a fast-to-slow transformation, in addition to muscle hypertrophy, whereas unloading causes a slow-to-fast transformation, in addition to muscle atrophy. Unloading applied during early development in a microgravity environment reduced the expression of the slow MyHC gene and augmented the expression of the fast MyHC-2X and -2B in the slow soleus muscle (Adams et al., 2000a). However, in these experimental models it is difficult to discriminate the direct effect


83:

of mechanical unloading from concomitant effects produced by electrical hyperactivity or silencing of motor neurons due to altered reflex activity. The finding that passive stretch can affect muscle growth even in denervated muscles point to the existence of biomechanical sensors that can regulate muscle gene expression independently o f neural reflex activity (Goldspink et al., 1992). A possible location for a biomechartical sensor is at the level of the muscle cytoskeleton and in particular at the interface of specialized regions of the sarcomere, the Z-discs. and specialized regions of the subsarcolemmal cytoskeleton, the costameres (Chien. 2000). New proteins identified at these sites may be involved in mechanosignaling. For exmnple. the muscle L1M protein (MLP) has been shown to promote myogenic differentiation (Arber et al., 1994) and MLP (-/-) mice develop dilated cardiomyopathy :with hypertrophy and hem-t failure after birth l.Arber et al., 19971. MLP exhibits a dual subcellular localization, being present in both the nucleus and in the cytoplasm (Arber et al., 1997: Kong et al.. 1997). In the cytoplasm MLP co-localizes with beta-spectfin at the sarcolemma overlying the Z- and M-lines of myofibrils, namely at the costameres, in both cardiac and skeletal muscle cells and interacts directly with alpha-actinin (Flick and Konieczny, 2000). Nuclear MLP functions through a physical interaction with the muscle basic helix-loop-helix (bHLH) transcription factors MyoD, MRF4. and myogenin (Kong et al.. 1997). Another potential candidate for mechanosignaling is representedby the newly discovered FATZ/calsarcins/myozenin proteins (Fanlkner et al.. 2000: Frey et al.. 2000b: Takada et al., 2001), a novel family of sarcomeric proteins that link ealcineurin with the contractile apparatus, thereby potentially coupling muscle activity to calcineurin activation. Calsarcin-t is expressed specifically in adult cardiac and slow-twitch skeletal muscle while Calsarcin-2 is restricted to fast skeletal muscle. ZASP/Cypher/Oracle (Faulkner et aI.. 1999; Pussier et al., 2000; Zhou et al., 1999) is a novel PDZ-LIM domain protein located a: the Z-discs that binds different types of PKC proteins and thus could function as an adapter to couple PKC-mediated signaling to the sarcomere. Sarcolemmal sensors at the level of the costameres, such as integrins, could also be involved in the mechanotransduction process by which cells transduce mechanical stresses into biochemical signals. Metabolic signals. Muscle contraction elicits a variety of immediate metabolic responses, such as those involving glucose transport and carbohydrate and fat metabolism. as well as long-term, transcription-dependent changes in metabolic enzymes. Metabolic changes precede contractile protein changes following low frequency electrostimulation of fast muscles. Intense exercise and low frequency electrostimulation cause a decrease in the ATP/ADP ratio and it has been suggested that an imbalance between energy requirement and energy supply might represent a signal for glycolyticto-oxidative and fast-to-slow switches in the patterns of muscle gene expression (Pette and Staron. 1997). This view is supported by the finding that similar responses are induced by creatine depletion, leading to ATP depletion, and by exercise under conditions of impaired blood flow. also leading to ATP depletion (see Pette and Starom 1997 and references therein). But how does, the muscle fiber sense ATP depletion m order to activate the appropriate signaling pathways? A good candidate to mediate this effect might be the AMP-activated protein kinase (AMPK), an enzyme that plays a key role in orchestrating

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many of the acute changes in carbohydrate and fat metabolism induced by exercise (Winder and Hardie, 1999). AMPK is activated by ATP depletion and consequent elevation of 5'-AMP via the adenylate kinase reaction (2ADP ---, ATP + AMP). The AMP-AMPK system can thus be considered as a "cellular fuel gauge" that monitors the energy status of the muscle fiber. In skeletal muscle AMPK is activated by hypoxia, exercise and electrostimulation and its level of activation parallels the intensity of the exercise. The effect of AMPK on muscle gene transcription is the object of active investigation, It will be interesting to determine whether transcriptional regulation of specific muscle genes, in particular those involved in fiber-type-specific metabolic profile (oxidative/glycolytic) are elicited by metabolic demands rather than by events occurring early during the excitation-contraction coupling, e.g. Ca2+ signaling. Interestingly, the targets of A M P K include not only metabolic enzymes but also components of transduction pathways potentially involved in muscle gene regulation such as Raf, a key intermediate in the pathway from Ras to MAPK(ERK) (Hardie et al., 1998). B.

Transduction pathways

Several signaling pathways, including calcineurin, CaMK and Ras, have been implicated in fiber-type-specific muscle gene regulation. Calcineurin. Activation of calcineurin, a calcium/calmodulin-regulated serine/threonine phosphatase, in cultured muscle cells selectively up-regulates slow-fiber-specific gene promoters and, conversely, inhibition of calcineurin activity by administration o f cyclosporin A promotes slow-to- fast fiber transformation in vivo (Chin et al., 1998). Calcineurin signaling appears to regulate a set of multigenic protein families, including myosin heavy chain (MyHC), sarcoplasmic reticulum ATPase (SERCA) isoforms and metabolic enzymes involved in the transition from slow oxidative (type I) to fast oxidative (type IIa) phenotype in soleus muscle (Bigard et al., 2000). Activated calcineurin was reported to promote the slow fiber type program both in cultured muscle ceils and after adenovirus-mediated gene transfer in vivo (Delling et al., 2000). However, the role of the calcineurin pathway is still controversial. Some groups found no change in fiber type distribution after treatment with cyclosporin A in adult rat muscles (Biting et al., 1998). Overexpression of active calcineurin induced both fast and slow muscle-specific promoters in cultured myotubes but did not activate either slow or fast reporter genes injected into rat soleus muscles (Swoap et al., 2000). Transgenic models have also been used to explore the role of calcineurin. Transgenic mice that expressed activated calcineurin under control of the muscle creatine kinase enhancer exhibited an increase in slow muscle fibers (Naya et al., 2000). However, although calcineurin expression from the transgene was at least 10-fold higher than the level of expression of the endogenous gene in skeletal muscle, the proportion of slow fibers was still extremely small. We have reinvestigated the role of calcineutin using a muscle regeneration model in which fiber type specification is dependent on slow motor neuron activity (Serrano et al., submitted). We found that the calcineurin inhibitors, Cyclosporin A and FK506, prevent the induction of MyHC-slow in the regenerating rat soleus muscle. An issue not

Fiber Type SpeCification in Vertebrate Skeletal Muscle

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addressed in previous studies in vivo is that. since calcineurin is expressed athigh levels in neural tissues (Klee et al., 1998) and is known to affect neurotransmittev release by nerve terminals. Cyclosporin A could affect primarily motor neuron function and only secondarily muscle gene expression. To distinguish between direct and indirect effects of calcineurin inhibitors, we took advantage of the observation that the up-regulation of MyHC-slow by slow motor neurons can be reproduced by electrostimulation of denervated soleus with a continuous 20 Hz pattern resembling the firing pattern of slow motor neurons. The finding that Cyclosporin A inhibits the up-regulation of MyHC-slow produced by direct etectrostimulation points to a primary role of calcineurin in muscle fibers. This interpretation was confirmed by the finding that the induction of MyHC-slow was also prevented in regenerating muscle fibers transfected with a ptasmid coding for the peptide cain/cabin-1, a physiologic peptide inhibitor of calcineurin. A large number of studies during the last few years has shown that Ras-dependent MAPK pathways, including ERK, JNK and p38 MAPKs, are transiently activated by contractile activity in both rat (Goodyear et al., 1996; Ryder et al., 2000; Sherwood et al.. 1999; Wretman et al., 2000) and human skeletal muscle (Aronson et al., 1998; Boppart et al., 2000; Krook et al., 2000: Osman et al,. 2000; Wideg~-en et al., 1998). MAPK stimulation is also found in isolated muscles, thus reflects contractile activity per se and not systemic influences (Ryder et al., 2000: Wretman et al., 2000). Based on these findings, it has been postulated that MAPK activation may serve as a link between contractile activity and transcriptional responses in skeletal.muscle fibers. However. no direct connection between MAPK activation and transcription of specific muscle genes has been demonstrated in these studies. Direct evidence for a role of Ras/MAPK pathways in activity-dependent muscle gene regulation was obtained using a different approach, namely by selective perturbation of this pathway with constitutively active and dominant negative mutants in vivo (Murgia et al., 2000). The experimental model used in these experiments is the regenerating rat soleus muscle that differentiates in a few days into a slow muscle in the presence of the slow motor neurons whereas it undergoes a default fast-like differentiation in the absence of the nerve. The rationale of this in vivo transfection approach is that a signal transducer involved in mediating the effects of nerve activity should be able to mimic the effects of innervation and induce a slow phenotype when transfected in a constitutively active form in denervated muscle. Conversely, the transfection of a dominant negative mutant of the same signal transducer into innervated muscle should inhibit the induction of the slow phenotype and maintain the default fast phenotype. Using this approach we found that a dominant negative Ras mutant (RasN17) prevents tile induction of MyHC-slow by slow motor neurons, while a Ras mutant IRasVI2S35) that selectively stimulates the MAPK/ERK pathway is able to induce MyHC-slow in the absence of innervation (Murgia et al.. 2000). On the other hand, another Ras double mutant (RasV12C40), that selectively stimulates the H 3 K pathway, was found to regulate muscle fiber size in a manner analogous to the action of electrical activity but didnot affect the expression of fiber type-specific muscle genes (Murgia et at., 2000).

Ras/MAPK.

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CaMK. Calcium/calmodnlin-dependent protein kinase IV (CaMKIV) has been recently

shown to enhance the calcineurin-induced stimulation of the transcriptional activity of myocyte enhancer factor-2 (MEF2) in cultured muscle cells (Wu et al., 2000). CaMK appears to promote myogenesis by relieving MEF2 from the repression of the class II histone deacetylases HDAC4 and HDAC5: in particular. CaMK was found to disrupt MEF2-HDAC complexes and induce nuclear export of HDAC5 (McKinsey et al., 2000a; McKinsey et al., 2000b). However. there is no direct evidence so far that CaMK can also affect fiber type-specific muscle gene transcription (see below). C.

Transcription factors

The MyoD family of bHLH transcription factors consists of four members: MyoD, Myf-5, myogenin and MRF4. The central role of these factors in myogenesis is well established, however their role in fiber-type-specific gene regulation remains to be established. All four bHLH myogenic transcription factors and especially myogenin are strongly up-regulated after denervation (Eftimie et al., 1991; Voytik et al., 1993) and have been implicated in the up-regulation of nicotinic acetylcholine receptor subunit genes in extrajunctional regions of denervated muscle fibers (Duclert and Changeux, 1995). Myf5 and MRF4, but not myogenin are up-regulated by low frequency stimulation combined with stretch in rat fast muscles (Jacobs-E1 et al., 1995). Myogenin is expressed at higher levels in slow oxidative adult muscles while MyoD is more abundant in fast muscles (Hughes et al., 1993; Voytik et al., 1993). However, fiber type distribution is unchanged in MyoD -/- mice. When overexpressed in fast muscle myogenin causes a switch to an oxidative metabolism but not to a slow myosin gene expression (Hughes et al., 1999). On the other hand, changes in expression of MyoD/myogenic transcripts do not correlate with changes in fast and slow myosins seen in skeletal muscle under conditions of hypothyroidism and chronic low-frequency stimulation (Krans and Pette, 1997).

MyoD/myogenin.

The myocyte enhancer factor-2 (MEF2) family consists of four members: MEF2A, MEF2B, MEF2C and MEF2D, each comprising several variants generated by alternative splicing. Using a MEF2-dependent transgene, Naya et al. found that the transcriptional activity of MEF2 is high in muscle tissues during mouse embryogenesis and declines after birth (Naya et al., 1999). Careful analysis of different transgenic lines with this MEF2 sensor showed that transcriptional activity of MEF2 is occasionally expressed in slow but not fast muscles (Wu et al., 2000). Thus it was suggested that MEF2 may act as one of the effectors of calcineurin signaling in inducing the slow fiber gene program (Wu et al., 2000). However, when the same construct was injected in adult fast and slow rat muscles, the level of activity was much higher in fast compared to slow muscles (our unpublished observations). Accordingly, the transcriptional activity of MEF2 was higher in regenerating denervated rat soleus, characterized by a default fast-like pattern of gene expression, compared to innervated soleus with a typical slow phenotype. It will be interesting to determine whether the forced expression of activated or dominant negative MEF2 mutants can affect muscle gene expression and fiber type profile.

MEF2.


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NF-AT. The family of NF-AT transcription factors consists of four members: NF-AT1 (NF-ATp), NF-AT2 (or NF-ATcl, NF-AT3 and NF-AT4 (or NF-ATx), each comprising several variants because of alternative splicing. NF-AT factors are characterized by a highly conserved DNA binding domain (Rel similarity domain) and a calcineurin binding domain and act as downstream effectors of calcineurin in different cell systems. The role of NF-AT transcription factors in muscle fiber type specification is still controversial. Transcriptional activation of slow-fiber-specific gene promoters in cultured muscle cells is dependent on the integrity of NF-AT sites (Chin et al., 19981. On the other hand. a troponin-slow promoter lacking the NF-AT site is specifically expressed in slow-twitch muscles of transgenic mice like the wild-type promoter (Calvo et al.. 1999). Slow MyHC expression was preferentially specified by activated calcineurin, but not by NF-ATc3 (Delting et al., 2000). Overexpression of NF-AT2 expression plasmids, that did strongly activate a NF-AT reporter, did not activate muscle-fiber-type-specific genes both in vitro and in vivo (Swoap et al., 2000).

A slow-muscle-fiber enhancer binding protein, called MusTRD1. was identified based on its binding to a specific sequence (USE B1) present in the human troponin I slow upstream enhancer (USE) (O'Mahoney et al., 1998). Mutations that disrupts USE B1 binding activity indicate that the USE B1 element is essential for high-level expression in slow-twitch muscles. The open reading frame described for MusTRD1 is identical to that of a gene (GTF3/GTF2IRD1) deleted in WilliamsBenren syndrome, a microdeletion syndrome caused by haploinsufficiency of genes at 7ql 1.23 and shows significant homology to the helixqoop-helix repeat domains o f the transcription factor TFII-I (Franke et al., 1999: Tassabehji et al., 1999). However, the putative MusTRDl-protein is 486 amino acids shorter than the predicted protein encoded by GTF3/GTE2IRD1. At the moment, there is no evidence that MusTRD1 is differentially expressed in fast and slow muscle fibers and can affect the fiber phenotype.

MusTRD1.

V.

Conclusions and open issues

For several years research on muscle fiber type specification has been mainly involved with the identification of the various fiber types present in skeletal muscles, their molecular composition with particular reference to myofibrillar protein isoforms, and their developmental and adaptive changes with particular reference to the role of innervation and electrostimulation. During the last few years the focus has switched to the study of the molecular signals responsible for myoblasts lineage determination in the embryo and activity-dependent muscle gene regulation in the adult animal, Future studies should clarify how intrinsic and extrinsic mechanisms interact to detei~nine the muscle fiber phenotype through the coordinated action of multiple signaling pathways. Several lines of evidence indicate that both intrinsic and extrinsic mechanisms dictate fiber type specificity. For example, denervated fast and slow rat muscles differ in their response to electrostimulation: slow myosin is not induced in fast muscles after two

88


months of electrostimulation with a low frequency impulse pattern that is able to maintain slow myosin expression in the denervated slow soleus muscle (Ausoni et al., 1990). The response of muscle fibers to hormonal regulation, e.g. the changes in MyHC profile induced by hyper- or hypo-thyroidism, are likewise dependent on the specific nature of each fiber type (Izumo et al., 1986). Intrinsic differences between the various fiber types also affect their response to intracellular signaling pathways involved in fiber type specification. For example, the role of Ras and calcineurin signaling in the induction of the slow phenotype appears to be restricted to "competent" or "responsive" muscle fibers. Constitutively activated RasV12 and the double mutant RasV12S35 are able to induce slow MyHC in t h e absence of the nerve in the regenerating slow soleus muscle but not in the regenerating fast extensor digitorum Iongus muscle (Mm'gia et al., 2000). Transgenic mice with overexpression of activated calcineurin show limited fast to slow fiber transformation, in that most fast fibers do not undergo this transition (Naya et al., 2000). One possibility is that there is heterogeneity among fiber types with respect to their sensitivity to calcineurin signaling and that only a small subset of fast fibers (possibly the type 2A muscle fibers?) are competent to respond to calcineurin. The cooperation of multiple signaling pathways is likely involved in activitydependent muscle fiber type specification. Recent results point to an interaction between calcineurin and CaMK (Wu et al., 2000) and it is possible that similar interactions take place between calcineurin and Ras-MAPK, as shown in other cell systems. Cooperation between Ras and calcineurin signaling is responsible for T cell activation (Aramburn et al., 2000) and functional interactions between Ras- and calcineurin-regulated pathways have been recently described also in cardiac muscle cells. Increased NF-AT activity, which is blocked by calcineurin inhibitors, is induced by constitutively active Ras in cultured primary cardiomyocytes (Ichida and Finkel, 2000) and angiotensin II-induced MAPK activation in vitro and in vivo is blocked by calcineurin inhibitors (Murat et al., 2000). It will be interesting to determine whether cross-talk between the two pathways is also present in skeletal muscle fibers. In addition, it remains to be established whether these various pathways are stimulated by distinct signals triggered by contractile activity or by a common triggering signal, for example intracellular Ca 2+. Ca2+-calmodulin is the physiological activator of both calcineurin and CaMK pathways, and Ras signaling can also be activated by changes in intracellular Ca 2+ concentration (Finkbeiner and Greenberg, 1996). The activation of a fiber type-specific gene program is a relatively long process that presumably requires several intermediate steps. For example, the up-regulation of slow MyHC in the regenerating rat soleus muscle takes place after 24-48 hours after the re-establishment of neuromuscular connections with slow motor neurons (Jerkovic et al., 1997) or after electrostimulation of the denervated regenerating soleus with a tonic low frequency pattern of impulses (our unpublished observations). The delay between the start of electrostimulation and the change in MyHC gene expression may be even longer (several days) with adult muscles undergoing fast-to-slow or slow-to-fast transformation (Williams and Neufer, 1996). Therefore it is unlikely that fiber type transformation induced by activity patterns is the result of a direct "excitationtranscription coupling", namely a signaling cascade impinging directly onto specific MyHC gene promoters. It seems more likely that the switch in muscle gene regulation


89

requires still unknown intermediate steps, possibly involving the up/down-regulation of other (regulatory?) muscle genes, ha addition, discrete steps in fiber type specification m a y be differentially affected by distinct signaling pathways. F o r e x a m p l e , the changes M y H C g e n e e x p r e s s i o n i n d u c e d by e l e c t r o s t i m u l a t i o n or h y p e r - / h y p o - t h y r o i d i s m generally f o l l o w the s e q u e n c e M y H C - s l o w o M y H C - 2 A o M y H C - 2 X ~ M y H C - 2 B (Pette and Staron. 1997; Schiaffino and R e g g i a n i . 1994). It is not k n o w n w h e t h e r these m y o s i n switches are all regulated b y the s a m e m e c h a n i s m s . W e h a v e recently o b s e r v e d that in a regenerating m u s c l e s y s t e m in w h i c h slow m o t o r neurons induce a fast-to-slow switch, inhibition o f calcineurin activity by c y c l o s p o r i n A b l o c k s the M y H C - 2 A --~ M y H C - s l o w but not the M y H C - 2 X --~ M y H C - 2 A switch (Serrano et al., submitted).

VI.

Acknowledgments

Our research was supported by grants f r o m the Ministero d e l l ' U n i v e r s i t 5 e della R i c e r c a Scientifica e T e c n o l o g i c a o f Italy, A g e n z i a Spaziale Italiana, Telethon-Italia, the E u r o p e a n C o m m i s s i o n (contract n. Q L K 6 - 2 0 0 0 - 0 0 5 3 0 ) , and the G i o v a n n i A r m e n i s e H a r v a r d F o u n d a t i o n for A d v a n c e d Scientific Research.

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patterning in the zebrafish. Danio rerio. Development 123. 153-64. Van Raamsdonk. W,. Pool, C. W.. and Te Kronnie, G. (1978). Differentiation of muscle fiber types in the teleost Brachydanio rerio. Anatomy and Embryology 153, 137-155. Van Swearingen, J., and Lance-Jones. C. (1995). Slow and fast muscle fibers are preferentially derived from myohlasts migrating into the chick limb bud at different developmental times. Dev B M 170. 321-37. Vivarelli, E.. Brown. W. E.. Whalen, R. G., and Cossu. G. ~1988). The expression of slow myosin during mammalian somitogenesis and limb bud differentiation. J Cell Biol 107. 2191-7. Voytik. S. L.. Przyborski, M., Badylak. S. F.. and Konieczny, S. F. (1993). Differential expression of muscle regulatory factor genes in normal and denervated adult rat hind]imb muscles. Dev Dyn 198. 214-24. Walro. J. M.. and Kucera. J. (1999). Why adult mammalian intrafusal and extrafusal fibers contain different myosin heavy-chain isoforms. Trends Nem'osci 22. 180-4, Widegren, U.. Jiang, X. J.. Krook, A., Chibalin, A. V., Bjomholm. M.. Tally, M., Roth, R. A., Henriksson, J., Wallberg-henriksson. H.. and Zierath. J. R. (1998). Divergent effects of exercise on metabolic and m~togenic signaling pathways in human skeletal muscle. Faseb J 12. 1379-89. WiUiams. R. S.. and Neufer, P. D. (1996). Regulation of gene expression in skeletal muscle by contractile activity. In Handbook of Physiology:Integration of Motor. Circulatory, Respiratory and Metabolic Control During Exercise.. L. B. Rowell, and J. T. Shepherd, eds. (American Physiological Society, Bethesda. M. D.), pp. 1124-1150. Winder. W. W., and Hardie, D. G. (1999). AMP-activated protein kinase, a metabolic master switch: possible roles in type 2 diabetes. Am J Physiol 277. El-10. Wretman. C., Widegren. U.. Lionikas. A.. Westerblad, H.. and Henriksson. J. (2000). Differential activation of mitogen-activated protein kinase signalling pathways by isometric contractions in isolated slow- and fast-twitch rat skeletal muscle. Acta Physiol Scand 170, 45-9. Wright, D. E., Zhou, L., Kucera, J., and Snider, W. D. (1997). Introduction of a neurotrophin-3 transgene into muscle selectively rescues proprioceptive neurons in mice lacking endogenous neurotrophin-3. Neuron 19, 503-17. Wu, H., Naya, F. J., McKinsey, T. A., Mercer, B., Shelton, J. M., Chin, E. R., Simard, A. R., Michel, R. N., Bassel-Duby, R., Olson, E. N., and Williams, R. S. (2000). MEF2 responds to multiple calcium-regulated signals in the control of skeletal muscle fiber type. Embo I 19, 1963-73. Yakar, S. Liu, J. L., Stannard, B., Butler, A., Accili, D., Saner, B., and LeRoith, D. (1999). NormaI growth and development in the absence of hepatic insulin-like growth factor L Proc Natt Acad Sci USA 96, 7324-9. Yang, S., Alnaqeeb, M., Simpson, H., and Goldspink, G. (1996). Cloning and characterization of an IGF-1 isoform expressed in skeletal muscle subjected to stretch. J Muscle Res Cell Motil 17, 487-95. Yu, F., Gothe, S., Wikstrom, L., Forrest, D., Vennstrom, B., and Larsson, L. (2000). Effects of thyroid hormone receptor gene disruption on myosin isoform expression in mouse skeletal muscles. Am J Physiol Regul Integr Comp Physiol 278, R1545-54. Zeman, R. J., Ludemann, R., Easton, T. G., and Etlinger, J. D. (1988). Slow to fast alterations in skeletal muscle fibers caused by clenbuterol, a beta 2-receptor agonist. Am J Physiol 254, E726-32. Zhou, Q., Ruiz-Lozano, P., Martone, M. E., and Chen, J. (1999). Cypher, a striated muscle-restricted PDZ and LIM domain-containing protein, binds to alpha-actinin-2 and protein kinase C. J Biol Chem 274, 19807-13.

CHAPTER 5

ROLE OF C Y T O K ~ S

IN SKELETAL MUSCLE G R O ~ H

AND

DIFFE~NTIATION

B.B. O L W I N , Y. B R E N - M A T T I S O N , D.D.W. C O R N E L I S O N , Y.V. F E D O R O V , H. F L A N A G A N - S T E E T and N.C. JONES


III.

IV.

V. VI.

I.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Historical perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A Development of cell lines and effects of conditioned media . . . . . . . . . . . . . . . . . . B. Identification of specific growth factor-mediated activities . . . . . . . . . . . . . . . . . . . Growth factors and their effects in myogenic cell culture: . . . . . . . . . . . . . . . . . . . . . . . A. Muscle cell proliferation, differentiation and apoptosis . . . . . . . . . . . . . . . . . . . . . B, Receptor tyrosine kinases and their signaling cascades . . . . . . . . . . . . . . . . . . . . . C. Receptor serine kinase signaling cascades . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Nuclear targets of myogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Growth factors and myogenesis in vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Muscle induction: A repressive mechanism? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Somitic muscle induction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Regulation of limb muscle development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusion and the future questions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

97 98 98 98 99 100 101 I07 108 110 110 110 111 114 115

Introduction

The problem of growth control is still one o f the largely unsolved puzzles in animal development. The organization of skeletal muscle is dependent on the proper integration of cell proliferation, differentiation and death. While the majority of myoblasts (conunitted muscle precursor cells) in the developing organism will undergo terminal differentiation and fuse into mulfnucleated myotubes, a relatively small number of ceils form a reservoir of quiescent, undifferentiated cells used for growth and repair of late pre-natal and postnatal skeletal muscle tissue. These quiescent but undifferentiated myoblasts, also called satellite cells, are typically activated following injury or exercise (Mauro, i 9 6 t ) . They are attached to the muscle fiber plasma membrane and surrounded by a common basement membrane. The coordinate regulation of myoblast cell proliferation and differentiation provides organism with a mechanism to control the deposition and patterning o f muscle 97 Advances in Developmental Biology and Biochemistry Ed. David Sassoon © 2 0 0 2 Elsevier Science. Printed in the Netherlands.

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mass during embryonic development, growth and regeneration. These events are most likely controlled by specific growth factors that are produced locally by muscle cells, motor neurons, or inflammatory cells. Growth factors, also referred to herein as cytokines, are a diverse family of extracellularly positively or negatively acting signaling proteins that affect these different aspects of myoblast cellular physiology.

II.

Historical perspectives

A.

Development of cell lines and effects of conditioned media

The influence of culture conditions on skeletal muscle differentiation has been an intense subject of investigaton since the early part of the 20th century (reviewed in Konigsberg, 1964). Much initial effort was focused on the origin of the multinucleated myotube and whether the nuclei arose from amotitic nuclear divisions or from proliferation of myoblasts followed by fusion of postmitotic cells into multinucleated myotubes (Konigsberg, 1964). This controversy was not resolved until skeletal muscle cells could be effectively cultured clonally. As an aside to these studies in the early 1960's, it was noted that serum concentrations affected differentiation of skeletal muscle cultures and noted that conditioned media significantly enhanced differentiation (Konigsberg, 1963). These observations were among the first to suggest that exogenous factors present in serum may be involved in regulating the proliferation or differentiation of skeletal muscle cells. Later that decade and in the early 1970's these observations were extended and it was elegantly demonstrated that medium conditioned by myoblasts was likely to be lacking specific factors necessary for repressing skeletal muscle differentiation and maintaining myoblast proliferation (Hauschka, 1972; White and Hanschka, 1971). Moreover, myoblasts obtained from different developmental stages exhibited distinct requirements for conditioned media, establishing that not all myoblasts are created equally (White et al., 1975). The molecular basis for this observation is not yet understood. B.

Identification of specific growth factor-mediated activities

The search for specific factors that would regulate skeletal muscle proliferation and differentiation spanned the 1970's and early 1980's; impure preparations of FGF were shown to promote myoblast proliferation and repress terminal differentiation (Gospodarowicz et al., 1975; Hauschka, 1981; Linkhart et al., 1980), however, these activities could not be definitively assigned to the FGFs until the late 1980's (Olwin and Hauschka, 1986). A hypothesis was established suggesting that serum mitogens repressed terminal differentiation of skeletal muscle cells, which was proposed to be a default state in culture (Clegg et al., 1987b). In the late 1980's, three groups established that members of the TGF~ family inhibited skeletal muscle cell differentiation, further strengthening the theory that the default state for skeletal muscle cells is to differentiate (Florini et al, 1986c; Massague et al., 1986a; Olson et al., 1986b). In conflict with these data, reports were published in the early 1980's suggesting insulin and IGFs are involved in promoting myoblast proliferation (Florini and Ewton, 1981). The concentrations

Role of Cytokines in Skeletal Muscle Growth and Differentiation

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of insulin required suggested that the actions of insulin were primarily mediated via the IGFs tsomatomedins). Unexpectedly, a role for this factor in stimulating differentiation was also seen (Florini et al.. 1986b). How one factor is capable of both stimulating proliferation and stimulating differentiation, two mutually exclusive events, is not clear and has not yet been resolved. However, recent data indicates that the IGFs may be most effective as hypertrophic factors, possibly resolving this conundrum (Musaro and Rosenthal, 1999). Hypertrophy of adult skeletal muscle in vivo would require activation and proliferation of satellite cells followed by differentiationl While it is not clear that IGFs can mediate all of these events, they clearly can induce hypertrophic responses when overexpressed in vivo (Barton-Davis et al.. 1998: Musaro et al.. 2001 ). In the late 1980's and 1990's identification of factors affecting proliferation, differentiation and survival of skeletal muscle cells increased almost exponentially: Concurrent with the explosive growth in identifying factors affecting myogenesis was a growth in the number of different skeletal muscle celt lines developed to analyze skeletal muscle proliferation and differentiation. A summary of known celt lines and factors affecting proliferation and differentiation illustrates the challenge to research groups attempting to understand the extracellular events that control myogenesis. The number of cell lines used to study myogenesis have added further to the overalt confusion regarding which factors are necessary for proliferation, differentiation, cell survival, etc. The most commonly used cell lines are from diverse origins that likely reflect the differences in their responsiveness to different factors: the L6/L8 rat myogemc cell lines were derived from neonatal rats (Yaffe, 1968); the: C2 celt line was derived from adult mouse tissue (Yaffe and Saxel, 1977) as was the MM14 cell line (Hauschka. t972"): the Rmo cell line was derived from adult rat skeletal muscle (Merrill, 1989); the Sol 8 cell line was derived from mouse soleus muscle: and. the BC3H1 cell line was derived from an induced brain tumor in adult :mice (Schubert et al.. 1974). Among other cells commonly used for the study of muscle proliferation a n d differentiation are C3H10T1/2 cells, derived from embryonic mouse mesoderm, which can be converted into muscle cell lines by treatment with 5-aza-cytidine or by ectopic overexpression of any of the MRFs (Konieczny and Emerson. 1984; Lassar et at., 1986: Taylor and Jones, t982).

IlL

Growth factors and their effects in myogenic cell culture

The large number o f factors and their differential effects on myoblast proliferation, differentiation and survival and the variety of cell lines used, makes inte~retation o f the effects of individual growth factors difficult if not circumspect. A commonl problem among all of the groups involved is to assume that an activity observed in one cell line o r in culture is an activity involved in '!skeletal muscle proliferation, differentiation or survival" when the effect may be: specific for a myoblast subtype or possibly a tissue culture artifact.

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Muscle cell proliferation, differentiation and apoptosis

A large number of growth factors, including FGFs (Chen and Quinn, 1992; Clegg et al., 1987a; Johnson and Allen, 1990: Kelvin et al., 1989; Kudla et al., 1995; Layne and Falxner, 1999; Milasincic et al., 1996; Olwin and Hauschka, 1986; Peng et al., 1997; Pizette et al., 1996; Rat and Kohtz, 1995; Rapraeger et al., 1991: Schofield and Wolpert, 1990; Schweigerer et al., 1987), PDGFs (Jin et al., 1993: Jin et al., 1991; Webb and Lee, 1997; Yablonka-Reuveni et al., 1990; Yablonka-Reuveni and Rivera, 1997; Ye et al., 1996), IGFs (Canicio et al., 1998: Chen and Quinn. 1992; Coolican et al., 1997b; Engert et al., 1996a; Ewton et al., 1994; Florini et al., 1986a; Johnson and Allen, 1990; Vandenburgh et al., 1991) inflammatory cytokines (Austin et al., 1992: Austin and Burgess, 1991; Cantini et al., 1995; Ogilvie et al., 2000; Okazaki et al., 1996) and hormones (Chert and Quinn, 1992; Cossu et al., 1989; De Angelis et al., 1992) positively regulate proliferation of myoblast cell lines and primary myoblasts in culture, Establishment of a role for these factors in vivo is a time consuming and difficult process; data implicating a small number of these factors in skeletal muscle development or regeneration is discussed later in this review. In addition to identifying growth stimulatory factors, a number of growth inhibitory factors have also been studied in muscle cell culture. These include TGF~s (Cusella-De Angelis et al., i994; De Angelis et al., 1998b; Filvaroff et al., 1994; Florini and Ewton, 1988: Florini et al.. 1986d; Johnson and Allen, 1990; Massague et al., 1986b: Olson et al., 1986a: Rat and Kohtz, 1995; Schofield and Wolpert, 1990; Spizz et al., 1987; Yamaguchi. 1995; Zentetla and Massague, 1992), TNFct (Layne and Farmer. 1999; Meadows et al.. 2000; Miller et al., 1988; Szalay et al., 1997), myostatin (Thomas et al.. 2000; Zhn et al., 2000), IFNg (Kalovidouris et al., 1993; Lough et at., 1982; Multhauf and Lough, 1986), IL-15 (Quinn et al., 1997), and PF-4 (Peng et al., 1997). Proliferation and differentiation of skeletal muscle cells are often considered mutually exclusive events. However, satellite cells in vivo exist in a quiescent, undifferentiated state that can be maintained for years, with satellite cell activation occurring on demand. Frequently, induction of myogenic differentiation in culture is initiated via removal of the majority of serum or by allowing the cultures to grow to a high density and self condition the media. Since the 1960's it has been recognized that factors present in serum act to prevent myogenic differentiation. However, it was not until the 1980's that specific factors were isolated that possessed the capacity to prevent terminal differentiation in the absence of cell division. FGFs were the first factors identified, which clearly repressed terminal myogenic differentiation in the absence of cell proliferation (Olwin and Hauschka, 1986; Clegg et al., 1987b) followed by TGF~s (Florini et al., 1986c; Massague et al., 1986a; Olson et al., 1986b). Of particular interest is the apparent opposite effect of these growth factors depending on serum conditions, where FGF-6 (Pizette et al., 1996) or TGF~ (Zentelta and Massagu6, 1992) can stimulate differentiation depending on the concentrations of factors or culture conditions. Insulin-like growth factors (IGFs), acting through a single receptor have also been reported to first stimulate proliferation and then stimulate differentiation of myoblasts (Coolican et al., 1997b; Engert et al., 1996a; Ewton et al., 1994; Florini et al., 1986a). Whether or not these factors can affect


101

multiple and potentially mutually exclusive events in skeletal muscle development or regeneration is not clear and will require further analysis of their actions in vivo. Programmed cell death, or apoptosis, is involved in many physiological processes occurring during embryonic development and tissue repair. It is also a process highly regulated by growth factors. Wasting of muscles in various physiological conditions may result from an imbalance in the growth factor environment. Simple withdrawal of factors may induce apoptotic cell death of muscle cells in vitro. Addition of FGF-2 (Stewart and Rotwein, 1996), IGFs (Meadows et al., 2000; Stewart and Rotwein. 1996), insulin (Stewart and Rotwein, 1996), PDGF-BB (Stewart and Rotwein, 1996) or thrombin (Chinni et al., 1999) protects myoblasts from apoptosis, while incubation in the presence of the inflammatory cytokine TNFc~ stimulates myoblast apoptosis (Meadows et al.. 2000). The role of cytokines in regulation of specific myoblast subtypes has not been carefully addressed. Elegant cell and tissue culture work has provided support for an inhibitory effect of TGF[3 on differentiation of primary fetal but not embryonic myoblasts (Cusella-De Angelis et al., 1994). Recent data from studies during chicken muscle development in vivo support a role for TGFI3 in fetal but not embryonic myoblasts (Flanagan-Steet et al.. submitted). Finally, a role for HGF in myoblast migration (Bladt et al., 1995a) as well as secondary myogenesis in vivo has been shown (Maina et al., 1996). Together, these data show that cytokine actions are likely to be specific for myoblast subtypes and that multiple cytokines will play critical roles directing skeletal muscle development at specific time points. Much additional work is needed to identify the important factors involved and to delineate their roles in myogenic development and regeneration. B.

Receptor tyrosine kinases and their signaling cascades

Signals from the cell surface to the nucleus are transduced by the binding of peptide growth factors to their respective transmembrane receptors resulting in the activation of specific intracellular signaling cascades. At least four different growth factor receptors are known to be involved in the regulation of skeletal myogenesis: the receptor tyrosine kinases (IGFRs, PDGFRs, and FGFRs (Figure 1)), the receptor serine kinases (TGF[3R superfamily (Figure 2)), hedgehog receptors (Figure 3), and wnt receptors. Although demonstrated to be involved in induction of myogenesis or myoblast proliferation and differentiation in vivo. the signaling pathways involving writ factors will not be discussed in detail. Dimerization of receptor tyrosine kinases (RTKs) through the binding of their respective figands turns on the kinase activity of the receptor (Schlessinger, 2000). The activated kinase then transphosphorylates the other receptor monomer and then can phosphorylate specific tyrosine residues on cytosolic membrane localized proteins (Lemmon and Schlessinger, 1994). Phosphorylated tyrosine residues provide a recognition site for protein/protein interactions mediated through Src homology 2 (SH2)and PTB (phosphotyrosine binding) domains allowing for the recruitment of downstream signaling molecules to the activated RTK (Pawson and Scott, 1997; ~Schlessinger, 2000).

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FGF Receptor-1 Signaling HSPG

Heterotrimeric G-protein Ras?

Proliferation

Differentiation

Figure 1. FGF receptor signaling. More is known concerning FGF receptor signaling in regard to myogenesis than any other growth factors. FGF receptor-1 can transduce intracellular signals that function independently to promote proliferation and repress terminal differentiation. Heparan sulfate provided by heparan sulfate proteoglycans (HSPGs) is necessary for FGF receptor dimerization, FGF binding and FGF activation of FGF receptors. Intracellular signaling pathways involving both proliferation and repression of myogenesis are mediated in part through the actions of Ras and unidentified heterotrimeric G-proteins. FGF activates the ERK1,2 and p38ct,13 pathways in proliferating myoblasts; both pathways are necessary for cell proliferation. Activation of the atypical PKCs appears necessary to repress myogenic differentiation and involves as yet unidentified MAPKs. Finally, cell proliferation indirectly inhibits myogenic differentiation so that direct repression of myogenesis and indirect repression of myogenesis via cell proliferation together contribute to maintain the undifferentiated state of proliferating myoblasts. How these intracellular signals inhibits the activities of the myoD or MEF2 family are unknown.


103

TGFS Type I receptor

( Figure 2. Receptor serine kinase signaling. These TGF[3familyreceptors can repress myogenic differentiation and inhibR myoblast proliferation upon ligand binding. The type II receptor binds ligand and associates as a dimer with a Type I receptor dimer forming a heterotetramer. In this active signaling complex, the type II receptor phosphorylates the type I receptor, which is then activated. The phosphorylated and active type I receptor then in turn phosphorylates receptor Smads. The phosphorylated receptor Smads associate with co-Stands, are translocated to the nucleus and act with co-activators (lef/TCF) or co-repress0rs (ski/snoN) to regulate target genes. Cdk inhibitors are among the target genes that are involved in TGFl3-mediatediflbibition of cell proliferation. How TGFI3signaling represses myogenic differentiation is not understood.

RTK-dependent activation of mitogen activated protein kinase ( M A P K ) signaling c a s c a d e s ~ e a well-defined mechanism for the regulation of mitogenesis and differentiation o f many varied cell types (Lewis et al., 1998; Robinson ~ d C o b b , 1997). A t l e a s t three M A P K families have b e e n identified: the extracellular signal regulated k i n a s e s (ERKs), the c-jun NH3-terminal kinases/stress activated p r o t e i n kinase (JNK/SAPKs), a n d the p 3 8 M A P K s (Lewis et al., 1998; Robinson and Cobb, 1997) i(Figure 4). C o m m o n to alI identified M A P K cascades is th e sequential ph0sphorylatio n o f signaling intermediates generally referred to as M A P K ~ ( M K K K ) , M A P K K (MKK), and finally

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HH

( PKA

Cofactors

~

Muscleinduction InhibitDifferentiation

HhTarget Genes

Figure 3.

Shh signaling. Shh is an important regulator of muscle induction in the myotome and represses

differentiation of myoblasts in the limb bud. The signaling mechanisms utilized by Shh are poorly understood. Shh is known to bind to patched (ptc), a 12-membrane-spanning receptor and stimulates smoothened (smo) to signal. This releases the transcription factor gli from an inactive complex in the cytoskeleton involving costal, fused (a kinase) and suppressor of fused. The active gli is then translocated to the nucleus and can act on target genes. A second mechanism for regulating gli activity involves protein kinase A (PKA). Phosphorylation of gli by PKA promotes proteolysis of gli, removing the transactivation domain. The proteolytic fragment of gli binds to gli sites on DNA and further inhibits hedgehog target gene transcription. It is not understood how Shh Signaling affects myogenesis.


105

Myogenesis Growth Factors

MKKK

Raf

MKK

MKKI/2

MAPK

MEKKs 1-4

?

ERKII2 ERK3 Promotes: Proliferation Differentiation

MAPKKK5

MLK3

DLK

?

MKK6

MKK5

1 MKK4

MKK7

JNK%3 Promotes: D~Iferentiation

MKK3

p38 ot~ p38 "° Promotes: Differentiation

p38

ERK5

Proliferation (c~J~)

Differe~tiatio~

Figure 4. MAPK cascades regulating myogenesis. Identified MAPK cascades and their reported activities on skeletal muscle myoblasts. ERK1,2 MAPKs have been reported to promote differentiation, promote proliferation, and inhibit differentiation. The majority of these studies were performed on cells where serum withdrawal is required to induce myogenesis. Serum reduction is likely to induce a cellular stress response and alter signaling in a number of the MAPK pathways. FGFs have been shown to activate both ERK[.2 and p38%13 in proliferating MM14 cells as well as primary myoblasts. In these experiments, both MAPKs have been shown to be necessary for myoblast proliferation. Additional data from a number of laboratories has shown a role for p38c~,[3 in promoting induction of muscle genes. Inhibition of these MAPKs prevents myogenic differentiation. The nuclear targets for these MAPKs are not clearly identified, however, p38 MAPKs have been shown to directly phosphorylate members of the MEF2 family of transcription factors,

M A P K ( L e w i s et al., 1998: R o b i n s o n and Cobb. 1997). O n c e phosphorylated, the M A P K translocates to the nucleus w h e r e it regulates transcription through phosphorylation o f defined transcription factors. In skeletal muscle, activation o f several M A P K s has b e e n i m p l i c a t e d in both inhibition and induction o f m y o g e n i c differentiation. T h e available data on the role o f the M K K 1 / 2 - E R K 1 / 2 cascade in regulation o f skeletal m y o g e n e s i s is conflicting. D e p e n d i n g u p o n cell type used, M K K 1 / 2 - E R K 1 / 2 activation either p r o m o t e s ( G r e d i n g e r et al.. 1998) or inhibits m y o g e n i c differentiation (Cootican et al.. 1997b; M i l a s i n c i c et al., 1996: W e y m a n and W o l f m a n . 1998). A d d i t i o n a l l y , p38 M A P K s (c~, 13 and y) h a v e b e e n identified as p o s i t i v e m e d i a t o r s o f skeletal m y o g e n e s i s

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(Cuenda and Cohen, 1999; Lechner et al., 1996; Wu et al., 2000; Zetser et al., 1999). Recent data from our laboratory suggests that p38c~/[~ acts in the Gl-phase of the cell cycle as a competence factor necessary for either proliferation or differentiation depending on the local cellular environment (Jones et al.. submitted). We propose that satellite cells require activation of p38c~/[~ MAPKs to exit from a quiescent state. Discrepancies within the cited results can be attributed to differences in cell specific signaling, to the use of serum as opposed to use of a single growth factor in experimental conditions, or to the correlation of the steady state activities of signaling intermediates with biological responses at time points far beyond the critical signaling event. Unlike satellite cells in vivo, which are maintained in a qmescent, undifferentiated state, the skeletal muscle cell lines used in the studies above differentiate upon cessation of proliferation and thus, inhibition of their proliferation is likely to indirectly induce myogenesis. Negative regulation of MAPK activity is modulated by a family of dual-specificity protein tyrosine phosphatases (dsPTPs). The dual-specificity phosphatases inactivate the catalytic activity of MAPKs through the removal of phosphates from both the Thr and Tyr residues in kinase regulatory domain (Neel and Tonks, 1997). Two dsPTPs have been implicated in regulating myogenic differentiation in skeletal muscle cell lines. MKP-3 (MAPK phosphatase-3) is specific for dephosphorylation of ERK1,2 and ERK 5 (Kamaknra et al., 1999). This dsPTP is dynamically expressed in MM14 cells; within 6h of FGF withdrawal the protein and mRNA is lost suggesting a role in maintenance of the proliferation or in early conversion to a terminally differentiated state (Mourey et al., 1996). Temporally, the loss of MKP-3 protein correlates well with the acquisition of a post-mitotic phenotype. A role for MKP-3 in myogenesis is not yet established, however, overexpression of MKP-3 prevents MM14 proliferation but does not induce muscle gene expression or overt differentiation (Jones et al., unpublished data). A second dsPTP, MKP-1 is a broad specificity phosphatase and was initially identified as an immediate early gene (Charles et al., 1993). Ectopic overexpression of MKP-1 in C2C12 cells inhibits myoblast fusion (Bennett and Tonks, 1997), suggesting that MAPK activity may be required for fusion of myoblasts into multi-nucleated myotubes. In addition to activation of MAPK signaling pathways, RTKs also activate phosphoinositide 3-kinase (PI3K) (Pawson and Scott, 1997). PI3Ks are activated either by recruitment to activated receptors via adapter molecules or through an undefined Ras-dependent mechanism. Initially identified through their ability to phosphorylate phosphatidylinositol, PI3Ks have recently been shown to activate both p38c~/~ and Akt (PKB). A role for p38ct/~ in promoting myogenesis has been described; inhibiting the activity of these kinases with the drug SB208030 prevents expression of skeletal muscle genes and myoblast fusion, suggesting that these kinases are necessary to promote myogenic differentiation (Cuenda and Cohen, 1999; Lechner et al., 1996; Wu et al., 2000; Zetser et al., 1999). Since IGFs promote hypertrophy and activate PI3K, it is tempting to speculate that activation of p38a/~ occurs downstream of IGF receptor activation and is required for IGF-mediated hypertrophy. However, MM14 cells and primary mus cle cells are blocked from proliferation by the SB208030 inhibitor, suggesting that the role(s) of p38a/~ are more complicated than previously thought (unpublished data).


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An additional p38 family member also known as p387 or ERK6 has also been implicated in regulating myogenesis. Overexpression of ERK6 enhances C2C12 cell differentiation. while overexpression of a catalytically inactive mutant prevents differentiation (Lechner et al.. 1996). Although interesting, these data have not been replicated in other muscle cells or in primary cultures. A second kinase activated by PI3K is Akl. a ser/thr kinase that contains a pleckstrin homology domain and requires phospholipid binding for activation. A number of reports in C2C 12 cells and in L6 cells show that activation of Akt is associated with enhanced myogenesis, including induction of muscle-specific gene expression and enhanced fusion (Fujio et al.. 1999; Lawlor and Rotwein. 2000). Some of these actions are thought to be mediated via the MEF2 family of transcription factors (Han et al., 1997~. Thus, stimulation of the PI3K pathway appears to positively stimulate myogenesis through at least two independent pathways, the p38a/~ MAPK pathway and through activation of Akt. C.

Receptor serine kinase signaling cascades

Two distinct classes of receptor serine kinases (RSKs) have been identified (Type I and Type II). The ligand induced active signaling complex is a hetero-tetramer containing two each of Type I and Type II receptors. Type II RSKs. including T~RII. activin receptor II (ActRII), ActRIIB, and bone morphogenic protein receptor II (BMPRII), are constitutively active serine kinases which unidirectionally phosphorylate Type I RSKs. Phosporylation of Type I RSKs, identified as activin receptor-like kinase 1-7 (ASK 1-7). turns on kinase activity resulting in phosphorylation of discrete intracellular signaling molecules (Massague, 1998: Massague et al., 2000; Wrana, 2000). ASK 1-7 dictate both extracellular signaling specificity (through interactions with defined ligands) as well as intracellular signaling specificity (through phosphorylation of specific Smad proteins). To date, eight Smads have been identified and further classified into functional groups (Massague. 1998; Wrana. 20001. Smads 1,2,3,5. and 8 are receptor-regulated smads (R-Smads~ which upon phosphorylation by specific ASKs recruit Smad4. also referred to as common-partner Smad (Co-Smad), translocate to the nucleus and regulate t~ranscription (Massague et al., 20001. Smads 6 and 7. the inhibitory Smads (I-Smads), directly interact with Type I RSKs thereby competitively inhibiting R-Smad activation. Although there is no established role for any Smad in the regulation of skeletal myogenesis. TGF~'s ability to inhibit myogenic differentiation in culture implies a functional role of SmadS. Moreover, subcellular distribution of Smads is dramatically altered upon terminal differentiation (McDermott. personal communication). An alternate downstream mediator of TGF[3, 0 appears necessary for TGF~-dependent inhibition of myogenic differentiation in developing myoblasts (Zappelli et al., 1996). Of particular interest is data demonstrating the interaction of the ski oncoprotein with the TGF~ signaling pathway (Massague et al., 2000). The ski protein functions as a corepressor for Smads. The obvious muscle hypertrophy obtained from the sN transgenic mouse (Sutrave et al.o 1990), it is tempting to speculate that ski blocks TGF[3 or myostatin gene transcription. Additionally, recent in vivo work has established that inhibition of TGF[3 signaling results in an increase in myoblast proliferation and myoblast cell number as

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well as disruption of normal muscle patterning further implicating TGF~ as a mediator of myogenic differentiation (Flanagan-Steet et al., submitted). D.

Nuclear targets of myogenesis

Among ultimate targets of peptide growth factor signaling cascades in skeletal muscle cells are the cell cycle regulatory proteins and the transcriptional machinery that regulates muscle specific gene expression. Essential to the control of myogenic determination and differentiation is the Myogenic Regulatory Factor (MRF) and MEF2 families of transcription factors. The MRF family is made up of four related basic-helixloop-helix proteins (bHLH): MyoD (Davis et al., 1987), Myf-5 (Braun et al., 1989), myogenin (Wright et al., 1989), and MRF4 (Rhodes and Konieczny, 1989). The MEF2 family contains five related members that are widely expressed during embryogenesis (Black and Olson, 1998). Expression of any one of the MRFs in several non-myogenic cell types converts those cells into skeletal muscle (Weintraub, 1993). The importance of MRFs in regulating myogenesis was further reinforced by eliminating their expression via homologous recombination. Targeted disruption of either MyoD or Myf-5, although causing a delay in myogenesis, has no long-term effect on muscle development (Braun et al., 1992; Rudnicki et al., 1992). However, disruption of both genes simultaneously results in complete loss of skeletal muscle (Rudnicki et al., 1993), similar to that observed for myogenin null mice. Myf-5 and MyoD are the first MRFs to be expressed in proliferating myoblasts, followed by expression of myogenin in differentiating myoblasts and MRF4 in the late stages of differentiation (Hannon et al., 1992; Pownall and Emerson. 1992). Based on knock out and expression data, a model has been proposed in which MyoD and Myf-5 act to commit the muscle progenitor cells to the myogenic lineage and myogenin and MRF4 regulate expression of muscle specific genes required for early and late stages of differentiation, respectively. Because MyoD and Myf-5 are expressed in proliferating myoblasts post-translational regulatory mechanisms must exist to modulate their transcriptional activity preventing precocious myogenic differentiation. The identification of a conserved threonine m the basic domain of MRFs and the demonstration that this site was phosphorylated by protein kinase C (PKC) suggested a possible model for growth factor-dependent inhibition of MRF transcriptional activity (Li et al.. 1992). However, subsequent studies were unable to detect phospho-threonine on ectopically expressed MRF4 isolated from proliferating myoblasts (Hardy et al., 1993). In addition, mutation of the phosphorylation site had no effect on growth factor-dependent inhibition of muscle specific gene expression (Hardy et al., 1993). Other mechanisms that modulate MRF transcriptional activity are the sequestration of MRFs in non-functional heterodimers with other proteins (Id 1-4. Mistl. I-mr, OUT, or Stral3) (Benezra et al., 1990; Chen et al., 1996; Kurabayashi et al., 1993; Lemercier et al.. 1998; Narumi et al., 2000; Pagliuca et al., 1995) or competition with MyoR or ZEB for binding to the E-box sequences (Lu et al., 1999; Postigo and Dean, 1997). Additionally, MyoD is hyper-phosphorylated in proliferating myoblasts and displays a much lower degree of phosphorylation in differentiating myoblasts (Kitzmann et al., 1999). Although hyperphosphorylation of myoD induces rapid turnover via a


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proteosome-dependent pathway (Kitzmann et al.. 1999: Song et al., 1998), growth factor-dependent MRF phosphorylation has not been clearly established and a role for cytokine regulated inhibition of MRFs remains controversial. The myocyte enhancer factor 2 (MEF2) family, directly interact with MRFs and synergistically activate muscle specific gene expression (Black et al., 1998). A potential role for growth factor regulation of MEF2 family members is much more well developed than for the MRFs. Two members of the MEF2 family. MEF2A and MEF2C, have recently been shown to be substrates of MAPKs (Han et al.. 1997: Yang et al.. I998: Zhao et al., 1999) and phosphorylation of MEFs by MAPKs positively regulates MEF2-dependent transcriptional activity. IGFs are direct activators of MEF2 activit5 and appear to activate MEF2-dependent transcription through two distinct signaling pathways involving MAPKs and calmodulin-dependent kinases. Phosphorylation of MEF2 by MAPKs is thought to block MEF2 association with histone deacetylases (HDACI 4 and 5, inhibiting HDAC activity and activating muscle gene expression via MRFs and MEF2 (McKinsey et al., 2000b). CaM-dependent kinases also appear to be involved as phosphorylation of HDAC4 and HDAC5 by CaM kinases prevents formation of MEF2-HDAC complexes and induces nuclear export of HDACs (McKinsey et al., 2000a). Induction of muscle gene expression is widely regarded as the hallmark of myogenic differentiation. However. the first phenotypic change ocurring in the myogenic differentiation program appears to be an irreversible exit from the cell cycle. MyoD, in addition to being a primary regulator of muscle specific gene expression, is also a regulator of myoblast proliferation Inhibition of proliferation and induction of muscle specific gene expression are independent biological activities of MyoD. as inhibition of DNA binding does not affect the ability of MyoD to inhibit cell proliferation (Crescenzi et al.. 1990). Although the exact mechanisms by which MyoD inhibits celt cycle progression are not well understood, it is clear that cell cycle arrest during differentiation requires a direct interaction between MyoD and the retinoblastoma tumor suppressor protein (Rb) (Gu et al.. 1993). Hypo-phosphorylated Rb is the predominam form in skeletal muscle and disruption of Rb expression prevents cell cycle exit (Halevy et al., 1995: Schneider et al., 1994). Interestingly, cdk phosphorylation of MyoD induces rapid ubiquitin mediated degradation effectively inhibiting muscle specific gene expression in proliferating myoblasts (Kitzmann et al.. 1999). Another mechanism by which MyoD may mediate cell cycle withdrawal is by induction of the cyclin dependent kinase inhibitor p21 (Guo et al., 1995; Haievy et al., 1995~. Myogenic cell lines and primary cultures are often induced to differentiate by reducing or eliminating the serum, which is often accompanied by a loss of cell number. Apoptosis occurring during terminal differentiation can be prevented b y inclusion of IGFs and has been mistaken for myotube hypertrophy when accurate cell counts are not performed. Of particular interest are a number of observations that relate cell cycle progression and apoptosis with myogenic .differentiation. In these studies, cdk.cyclin D complexes inhibit myogenic differentiation, while prevention of apoptosis and enhancement of myogenesis is dependent on cdk inhibitors, potentially mediated via the actions of Akt (Fujio et al:. 1999; Guo and Walsh. 1997: Lawlor and Rotwein. 2000: Walsh et al.. 1996; Walsb and Perlman. 1997: Wang et al.. 1997: Wang and Walsh, 1996).

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While some of the targets for signaling cascades regulating myoblast proliferation and differentiation have been identified, the mechanisms involved in transducing and receiving specific signals remains to be determined. Unravelling the intricate relationships between myoblast proliferation, differentiation and survival will represent a continued challenge for future research efforts.

IV.

Growth factors and myogenesis in vivo

A.

Muscle induction: A repressive mechanism?

The mechanisms involved in muscle cell induction from mesoderm are not entirely clear and likely to involve both repressive and instructive events. In early embryonic development, gastmlation involves ingression of cells from the dorsal epiblast layer through the primitive streak and migrating between the epiblast and hypoblast layers to form the mesoderm (Bellairs, 86). Since endoderm cells also originate from the epiblast all embryonic tissues are derived from the epiblast (Rosenquist, 1971; Stern and Canning, 1990; Stem et al., 1988). Interestingly, cells from whole embryos or the primitive streak can differentiate into muscle in culture, but not without DNA synthesis (Baldwin et al., 1991; Holtzer et al., 1990). Epiblasts (prior to gastmlation, stage 3-5), dissociated into single cell suspension and grown in serum-free medium, form primarily muscle cells (George-Weinstein et al., 1996). However, when the epiblast tissue was cultured intact, or cells were plated at high density, myogenesis was largely repressed, suggesting that cell-cell interactions repress muscle-specific differentiation at this stage of development. While MyoD mRNA can be detected in the epiblast (MyoD mRNA can also be detected in the mesoderm/hypoblast layers, though cultures from these layers form primarily cardiac cells), MyoD protein is not detected in intact epiblasts and is only detected 2 hours after plating. Moreover, when epiblast cells are implanted into the limb bud they do not form muscle. These observations suggest that the mechanism of in vivo muscle induction may involve the de-repression of inhibitory signals generated be cell-cell interactions. The involvement of specific extracellular factors in these events has not yet been described. B.

Somitic muscle induction

The majority of muscles of the vertebrate body are derived from the somites and induction, proliferation and differentiation of skeletal muscle is likely to be under the influence of both positive and negative signaling molecules. Two distinct muscle lineages exist within the somites; the epaxial muscles which originate in the dorsal medial region of the somites and give rise to the deep back muscles and the hypaxial muscles which originate in the lateral region of the somite and give rise to body wall and limb muscle (Ordahl and Le-Douarin, 1992). Axial structures, (including the neural tube and notochord) and surface ectoderm express factors (e.g. members of the Wnt family and Sonic Hedgehog) that act positively to regulate somite segmentation and differentiation, the lateral mesoderm appears to negatively


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regulate myogenesis and inhibits terminal differentiation in part by secreting BMP-4 (Christ et al., 1992: Pourquie et al., 1995; Pourqnie et al., 1996). BMP-4 may inhibit myogenesis by repressing expression of MvoD and Myf-5 (Reshef et al., 1998). In the myotome, noggin inhibits BMP-4 and is thought to then allow MRF expression promoting myogenesis in the myotome (Hirsinger et al., 1997). Other growth factors involved in myotomal myogenesis include Shh, which in combination with members of the Wnt family, induce myotomal MRF expression (Munsterberg et al., 19951: FGF-2 in combination with TGF~I also appear capable of promoting myogenesis in single somite assays (Stern et al., 1995: Stern and Hauschka, 1995; Stern et al.. 1997). Finally, insulin and insulin-like growth factors I and II (IGF-I and IGF-II) may also act as somite myogenic signals acting synergistically with Shh. FGF-2 and TGF[31 (Kiefer et al., 1990). How these factors regulate myogenic induction is not understood and may involve regulation of MRF expression, increases in muscle progenitor cell proliferation, or inhibition of muscle progenitor cell apoptosis (Cann et al.. 1999). Clearly, the induction or derepression of the skeletal muscle myoblast phenotype is complicated and not well understood. Additional new data add to this complexity and suggest that the role(s) for the MRFs may differ in early and later embryogenesis (Kiefer and Hauschka, 2001). C.

Regulation of limb muscle development

Less information regarding the factors that regulate proliferation, differentiation and patterning of limb skeletal muscle is available. Some of the factors implicated in regulation of limb myogenesis are similar to those involved in myotomal myogenesis and include FGFs (Gospodarowicz et al., 1975; Kardami et al.. 1985a: Seed and Hauschka. 1988), IGFs (Allen and Boxhom. 1989: Florini et al.. 1994), TGF[3s (Allen and Boxhorn, 1989; Florini et al., 1986c; Olson et al., 1986b), PDGFs (Soriano et al., unpublished dataL BMPs (Amthor et al.. 1998), SF/HGF (Brand-Saberi et al.. 1996: Heymann et al., 1996: Scaal et al., 1999), and Shh (Duprez et al.. 1998b). In the limb, myogenesis begins when SF/HGF (Brand-Saberi et al.. 1996~. initiates de-epithelialization, detachment, and migration of muscle precursor cells fi'om the lateral dermamyotome (Bladt et al.. 1995b). The HGF receptor (c-met)is directly regulated by Pax-3 (Daston et al.. 1996: Epstein et al.. 1996) and loss of Pax-3. as evidenced by the Splotch mutant mouse, results in complete down regulation of c-met and loss of the migratory muscle populations of the diaphragm, tongue, and limb. This phenotype is similarly recapitulated by null mutations in the c-met receptor (Bladt et al., 1995a). Once reaching their destination in the limb bud mesenchyme, the myogenic precursors become restricted to two distinct masses, one located in the dorsal limb quadrant, the other the ventral. Maintenance of these cells as an undifferentiated pool of proliferating myoblasts is critical for continued muscle expansion and outgrowth. Coordinating the spatial position and appropriate growth of the pre-muscle masses is likely mediated by the activities of several growth factors, some of which may include SF/HGF, Shh, FGFs. BMPs. and noggin. SF/HGF is expressed throughout the limb bud mesenchvme with high levels restricted to the anterior portion (Myokai et al.. 1995: Thery et al.. 1995).

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Consistent with its motogenic properties, SF/HGF inhibits MyoD expression and prevents premature differentiation (Scaal et al.. 1999). This activity may in part be regulated by the actions of FGFs secreted by the apical ectodermal ridge (AER) and BMPs emitted from the zone of polarizing activity (ZPA) and the limb ectoderm. Expression of FGFs is thought to maintain SF/HGF, while BMPs-2. 4. and 7 in the anterior, posterior, and distal limb margins have been shown to inhibit its expression. In addition, BMPs aiso exhibit dose-dependent control of Pax-3 production (Amthor et al., 1998). Pax-3, which is generally considered a marker for undifferentiated myoblasts, is inhibited by high levels of BMPs but enhanced by low BMP concentrations. These activities are counter-balanced by the presence of noggin, a known BMP antagonist, in the limb core (Amthor et al., 1998). Appropriately, the pre-muscle masses are precisely situated mid-way between high levels of BMP-2. BMP-4. BMP-7, and noggin. In this way AER-produced FGFs, mesenchymal noggin, and ectodermallyderived BMPs may help coordinate the spatial position and appropriate growth of the pre-muscle masses. Inappropriate emigration of muscle precursor cells might be disastrous and therefore, factors expressed in these limb regions may function to precisely guard and maintain the boundaries of the pre-muscle masses. Establishment and maintenance of the dorsal and ventral masses does not. however, end here. Shh, which is believed to be restricted to the posteriorly-positioned ZPA. is essential to its fidelity. Ectopic Shh enhances Pax-3 and MyoD expression, eventually increasing overall limb mass (Duprez et al., 1998a) (Bren Mattison and Olwin, submitted). In agreement with these data additional information has shown that inhibition of Shh signaling decreases MyoD levels and resuks in a loss of skeletal muscle (Bren-Mattison and Olwin, submitted). These data suggest that Shh normally inhibits differentiation and/or stimulates proliferation. Therefore. it is conceivable that Shh maintains an early population of myoblasts in an undifferentiated state until it is appropriate for them to differentiate. The transition from the proliferative Pax-3-positive state to one that xs myogenically permissive is an important step in limb myogenesis. Differentiation and fusion may occur in this early muscle lineage as individual myoblasts gain distance from the limb tissues that repress myogenesis. This is supported by expression patterns of Pax-3 and MyoD, where cells closest to the ectoderm and ZPA express Pax-3, while those cells more centrally located, express MyoD (Amthor et al., 1998). Once myogenic transcription is initiated in these central cells, they are capable of undergoing terminal differentiation and fusing to form the scaffolding of primary myotubes. FGFs also play a significant role during primary myogenesis. In addition to promoting SF/HGF expression, FGFs directly regulate the life cycle of skeletal muscle myoblasts. FGFs have been shown in vitro (Kardami et al., 1985b; Seed and Hauschka, 1988) and in vivo (Flanagan-Steet et al., 2000) to stimulate proliferation and inhibit differentiation of skeletal muscle cell lines and primary myoblasts. Eliminating FGF signaling in developing myoblasts results in cell cycle withdrawal and precocious differentiation. While it is not entirely clear how FGF signaling is molecularly linked to its biological outputs, several elaborate signaling cascades are essential for its regulation of myogenesis. In particular, two distinct pathways mediate FGF-induced proliferation and inhibition of differentiation (Jones et al., 2001; Kudla et al., 1995; Fedorov et al., 2001). These findings provide possible insights as to how the protein effectors that transduce


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FGF signals may be linked to the molecular regulators of myogenesis. Although no direct connections have yet been drawn, it iS clear that FGFs play an important role during skeletal muscle development. Unlike the numerous molecules identified that affect primary myogenesis, relatively few factors are known to specifically regulate secondary myogenesis. Thus far, TGF[~ appears to be the major modulator of this process in vivo (F!anagan-Steet and Olwin, submitted), Addition of TGF[~ to cultured skeletal muscle cell lines inhibits their ability to both proliferate and differentiate. While TGF[3 likely controls proliferation through such cell cycle inhibitors as p15, p21, and p27, one study has shown that selective inhibition of MEF2C nuclear translocation may be responsible for TGFI3's effects on differentiation (De Angelis et al., 1998a). TGF~'s role in differentiation is somewhat controversial since addition of TGF[3 to muscle cell cultures in the presence of high levels of serum stimulated differentiation (Zentella and Massagu6, 1992). Interestingly, when TGF[3s are applied to primary myoblasts, their inhibitory effects are restricted to fetal cells isolated from animals undergoing secondary myogenesis (Cusella-De Angelis et al., 1994). This effect is thought to be mediated by PKC0, which is selectively expressed in fetal and post-natal muscle cells (Zappelli el al., 1996). Identification of this lineage-specific signaling molecule was highly significant. It provided:a mechanism for how two subpopulations of muscle precursor cells could exist within a single environment and respond in phenotypically opposite manners. As primary myoblasts differentiate and fuse, secondary myoblasts maintain active proliferation. Spatial and temporal regulation of myogenesis is likely to be critical for the generation of histologically distinct muscle fibers during skeletal muscle development. The late processes of myogenic maturation are characterized by skeletal muscle hypertrophy and incorporation of the satellite cells beneath the basement membranes of individual fibers~ One molecule in particular, IGF-I, has been associated with promotion of the :hypertrOphic response. Addition of IGF-I to primary satellite cells stimulates their proliferation while concomitantly promoting differentiation (Alien and Boxhom, 1989; Engert et al.. 1996b; Quinn et al., 1994). Studies by Allen et al. (1989) demonstrated that TGFI3 was able to attenuate the magnitude of cellular responses to IGF-I, while FGF exacerbated them. In culture, IGF-I mediates its effects on proliferation and differentiation through two temporally separate pathways. Initially, IGF-I stimulates proliferation through MAPK-induced expression of cell cycle progression factors (Coolican et al., 1997a). This is followed by stimulation of differentiation, which is controlled by the phosphatidylinositol (PI)-3-kinase/AKT pathway (Kaliman et al., 1996). Induction of this pathway leads to upregulation of the myogenic regulatory factors and muscle-specific gene expression (Musaro et al., 1995). Interestingly, recent studies in vivo suggest that IGF-I's effects on skeletal muscle may not require prier proliferation. In fact. post-mitotic expression of IGF-I potentiates and accelerates the myogenic program, inducing dramatic myotube hypertrophy (Musaro and Rosenthal, 1999). Further, virally expressed IGF-I rescues age-related effects of muscle degeneration (Barton-Davis et al., 1998; Musaro et al., 2001). Together these data suggest a dual role for IGF s i g n i n g during the myogenic differentiation. Several additional factors have been implicated in skeletal muscle: maturation and regeneration. These include FGFs (Olwin and Hauschka, t986), EGFs (epidermal growth

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factors), TGF]3 (Allen and Boxhorn, 1989), and SF/HGF (Comelison and Wold, 1997; Tatsumi et al., 1998). The FGFR1, FGFR4, and c-met tyrosine kinases are expressed on the quiescent satellite cells of mature myofibers (Comelison and Wold, 1997). Although, the precise roles of these factors during satellite cell activation and skeletal muscle regeneration are unclear, their presence in late stage muscle tissue and known functions during early myogenesis suggest potential roles in the activation of quiescent satellite cells. One can imagine how association of c-met and SF/HGF could release the inactive myoblast from its site of hibernation and stimulate movement to the point of injury. Further, FGFs may subsequently promote proliferation to enable restricted expansion and repopulation of this injured region. These activities may be counter balanced by the presence of TGF[3s, which are known in culture to inhibit proliferation and differentiation of primary rat satellite cells. In this way several growth factors likely function simultaneously to spatially and temporally maintain the balance of satellite cell activation, proliferation, and differentiation. The generation of diversity is central to the process of skeletal muscle myogenesis. Morphologically individual skeletal muscles differ from each other on the basis of contractile rate, metabolism, muscle-specific protein expression, and fiber type composition. Each of these properties is established during development as progressive waves of fibers form. presumably from different populations of progenitor cells (Bonnet and Hauschka, 1974; Rutz et al., 1982; Rutz and Hauschka, 1982; Rutz and Hauschka, 1983; Seed and Hansclaka, 1984; Van Swearingen and Lance-Jones, 1995; White et al., 1975). A number of distinct subsets of skeletal muscle myoblasts have been identified in the somite and the limb. These include the epaxial and hypaxial lineages of the early somite. the craniofacial lineage of the prechordal mesoderm, and the embryonic, fetal, and adult populations in the limb. The epaxial and hypaxial myoblasts could be considered the primary lineages as they generate the bulk of the body's skeletal muscle. The cells of embryonic, fetal, and adult origin are subclasses of hypaxial muscle that are believed responsible for the formation of primary, secondary, and adult myofibers in the limb. Little is known concerning the relationship of these cells to each other and the cytokines that may be involved in the regulation of growth and differentiation of these distinct cell populations. Future work will undoubtedly focus on some of the factors involved in generating skeletal muscle diversity and patterning during embryogenesis and regeneration.

V.

Conclusion and the future questions

The role of growth factors during myogenesis in vivo is less well understood than what has been observed for studies in vitro: not only does myogenesis occur in a much less controlled and defined environment in vivo, where there are many and diverse sources of equally numerous and diverse active factors, there is also the question of which myogenic cells are actually being observed. During the course of development and postnatal life, several more or less well-defined but overlapping types of myoblasts will arise: axial myoblasts will arise and differentiate in the somites to produce the body wall musculature; embryonic, fetal and possibly adult hypaxial (limb) myoblasts will

Rote of Cytokines in Skeletal Muscle Growth and Differentiation

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emigrate in waves from the somites into the prospective limb fields; myoblasts destined to produce some muscles of the head will derive from and migrate with the neural crest; and new evidence suggests that cells with at least myogenic potential arise from and reside in blood and marrow compartments. Additionally, all of these populations will eventually give rise to differentiated muscles expressing mixtures of defined suites of genes, for example slow vs. fast myosin heavy chains. Given the diversity of origins, fates, and forms found in the myoblast population, it would be naive to assume that all myoblasts would respond in the same manner to any given growth factor. Indeed. this would often not even be desirable, as many of these types of myoblast exist in overlapping environments, and the organism must have some means of restricting responses to environmental factors among each population. This, then. poses a major caveat when discussing the broad role of growth factors in myogenesis in vivo, especially when attempting to generalize results coming from tissue culture. For example, it appears that in C2C12 myoblasts, the most commonly used myogenic cell line, autocrine stimulation by IGF II is absolutely required for maintenance of MyoD expression (Montarras et al., t996) and successful differentiation (Florini et al., 1991), yet IGF signalling appears to be unimportant during embryonic myogenesis and its role during satellite myogenesis is as yet unknown. Another example. from the MM14 cell line is that these cells express only FGFR1 of the four FGF receptors, while primary satellite cells express all four at differing levels (Comelison et al., 2000; Sheehan and Allen. 1999) and embryonic myoblasts express mainly FGFR4 (Stark et al., 1991). A major challenge to groups interested in extracellular factor regulation of muscle development will be to unravel the complexities of the activities observed for these factors and establish the relevance of the activities observed in cell culture with their biological roles in skeletal muscle development or regeneration in vivo. We can ask to what extent, then, will intracellular signalling pathways worked out in these cell lines reflect myogenesis in vivo, and in what populations will these pathways be relevant?

VI.

References

Allen, R. E.. and Boxhom, L. K. (1989). Regulation of skeletal muscle satellite cell proliferation and differentiation by transforming growth factor-beta, instdin-like growth factor I. and fibroblast growth factor. J Cell Physiol 138, 311-5. Amthor, H., Christ, B , Weil, M.. and Patel, K. (1998). The importance of timing differentiation during limb muscle development. Curr Biol 8. 642-52. Austin, L., Bower, J., Kurek, J., and Vakakis. N. (1992). Effects of leukaemia inhibitory factor and other cytokines on murine and human myoblast proliferation. J Neurol Sci 112. 185-91 Austin, L.. mad Burgess. A. W. (1991). Stimulation of myoblast proliferation in culture by leukaemia inhibitory factor and other cytokines. J Neurol Sei 101, 193-7. Baldwin. H. S., Jensen. K. L., and Solursh, M. (1991). Myogenic cytodifferentiation of the precardiac mesoderm in the rat. Differentiation 47, 163-72. Barton-Davis. E. R.. Shomrma. D. I., Musaro. A., Rosenthal, N., and Sweeney, H. L. (1998). Viral mediated expression of insulin-like growth factor I blocks the aging-related loss of skeletal muscle function

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CHAFFER 6

SOMITIC AND NON S O . T I C PROGENITORS OF S ~ L E T A L MUSCLE GIULIO COSSU

Table of contents I. II. III. IV. V. VI. VII. VIII. IX. X.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Myogenic commitment: Signals from neighboring tissues . . . . . . . . . . . . . . . . . . . . . . Fate diversification in newly formed somites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The formation of primary and secondary muscle fibers . . . . . . . . . . . . . . . . . . . . . . . . . A role for PKC0 in myoblast diversification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Satellite cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Unorthodox myogenic progenitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

I.

Introduction

127 128 129 131 133 133 135 136 137 138

In the developing chordate, skeletal muscle differentiation begins shortly after gastrulation and persists, in some respects, for the entire life-span of the animal (Hausckha, 1994). Cells of the somitic mesoderm are committed to myogenesis by signals emanating from adjacent structures (Christ and Ordahl, 1994). In the last decade some of these signaling molecules and many of the responding genes have been identified. Wnts, Sonic hedgehog, Noggin and Bone morphogenetic protein (BMP) have been shown to act positively or negatively on this process (reviewed in Cossu and Borello. 1999; Borycki and Emerson, 2000). In the dorsal domain of newly formed somite. Shh and Wnts activate Myf5 and MyoD s that in turn control myogenic differentiation and myotome formation. It is not clear whether all myogenic cells are committed at once in the somites. It is however clear that only a minority o f the somitic cells wilt differentiate to form the myotome and synthesize contractile proteins that accumulate in the cytoplasm and selfassemble into sarcomeres. Once a myogenic cell has differentiated it cannot divide any longer. As a consequence, growth of the muscle fiber during fetal and post-natal development depends upon accumulation of new sarcomeres (hypertrophy) and upon addition of myoblasts (hyperplasia). These myoblasts must be instructed on when to divide and when to differentiate, by either fusing with pre-existing fibers or among themselves to generate a new fiber. It is therefore obvious that diversification of myogenic celt fate is as crucial 127 Advances in Developmental Biology and Biochemistry Ed. David Sassoon 127 © 2002 Elsevier Science. Printed in the Netherlands.

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as commitment. In fact it allows the production of post-mitotic skeletal muscle during early embryogenesis and at the same time the maintenance of a pool of mitotic progenitors that permits further growth of the tissue as well as regeneration in response to injury. This scenario may be complicated if we postulate that not all myogenic progenitors ceils are committed in the newly formed somites and moreover, that part of the progenitor cells may be derived from the incoming vasculature and not from the somite.

II.

Myogenic commitment: Signals from neighboring tissues

In newly formed somites, signals from axial structures (neural tube/notochord complex) and dorsal ectoderm are required to promote myogenesis, respectively in the epaxial domain (future back muscle) and in the hypaxial domain (future body wall and limb muscles) (reviewed in Cossu et al. 1996a). In mammals, axial structures activate myogenesis through a Myf5 dependent pathway while dorsal ectoderm acts through a MyoD dependent pathway. The latter is dependent upon previous expression of either Myf5 or Pax3 (Tajbakhsh et al. 1997). This suggested the existence of two myogenic lineages: epaxial and hypaxial, that differ for the nature of the inducing tissue, the regulatory gene activated and the mature muscles that will be generated. However, it should be noted that a significant level of compensatory plasticity must exist given the phenotype of the single MyoD or Myf5 knock-out mice. Myf5 null embryos have initially epaxial muscle defects whereas MyoD null embryos have predominantly hypaxial muscle defects (Braun et al. 1992; Kablar et al. 1997) but in both cases, the residual gene is sufficient to support almost normal skeletal muscle development throughout the body. A combination of Shh, normally produced by the notochord and members of the Wnt family, expressed in the neural tube, can replace the activity of axial structures in activating myogenesis (Musterberg et al. 1995). In the lateral myogenic progenitor cells, MyoD expression and subsequent terminal differentiation is transiently repressed in order to allow migration to the limb and body wall where muscle formation will take place. It was shown that the lateral mesoderm inhibits MyoD expression and that cells expressing BMP4 (Pourqui6 et al. 1996) could replace this inhibitory activity. The irLhibitory action of BMP is counteracted by Noggin, produced by dorsal neural tube. While BMP and noggin are likely to interact directly, Wnt and Shh act through classic membrane receptors and post-receptor pathways that are beginning to be elucidated (reviewed in Cossu and Borello, 1999; Borycki and Emerson, 2000). From the data discussed above, it appears that several molecules (and probably others yet to be identified) act on somites to instruct some of the cells in the dorsal domain to adopt a myogenic fate. Interestingly, Shh and Wnts are also known to promote cell proliferation and survival, suggesting that a combinatorial action of Shh/Wnt on transcriptional activation, proliferation and survival must account for the final number of differentiated cells in a given structure such as the myotome. This is relevant to the next issue discussed in this review, namely how different fates are chosen within contiguous and probably equivalent cells of the epithelial dermomyotome (Tajbakhsh and Cossu, 1997).

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Somitic and Non Somitic Progenitors of Skeletal Muscle

IH.

Fate diversification in newly formed somites

Most of the work discussed above on the activation of myogenesis refers to: the formation of the myotome i However we do not know whether commAtment of the progenitors responsible for later phases of myogenesis occurs through the:same m e c h a n i s m s and in the same spati~temporal context. If this is the case, then m y o g e n i c determination must b e compatible with sustained proliferation, a n d differentiation mus~ be repressed until the cells find themselves in the right time and place, and, more importantly, after a given number of divisions, as to attain the correct number of myobl~,_sts.

Commitment and differentiation in dorsal somJte Wntl, 3a

Wnt 7a, 6

Dorsal

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BMP4 Lateral

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,. ,

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Figure 1 Myogeniccommitmentand differentiation.The model represents cells in the dorsal domain of newly formed somites that respond to inductive signals (Shh and Wnts) and are committedto myogenesis. From in wvo gene expressionpatterns and dye labelingexperiment,it is assumedthat cells in the dorso-medial domain (the future dorso-medial lip) are the first to activate Myf5 and to differentiate into the epaxial myotome. Cells in the dorso-lateraldomainare committed (througha Pax3-MyoDpathway) tOform migratory myoblasts at the limb level and the hypaxial myotome in the inteflimb level that later will contributeto the inter-costal muscles. Cells located in the central region of the dorsal somite are presumed to give rise to later generationsof myoblasts and also to non muscle cells.

In the dermomyotome (Figure 1), only a fraction of committed myogenic precursors are induced to terminally differentiate and migrate through the dorso-medial lip and

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(in the inter-limb level) through the ventro-lateral tip to form the first differentiated, mononucleated myocytes of the myotome (Williams and Ordahl 1997; Currie and Ingham 1998; Kalcheim et al. 1999). The remaining cells in the dermo-myotome are kept as undifferentiated progenitors that ingress the myotome at later stages and through different pathways (Cinnamon et al. 2001; Denetclaw and Ordahl, 2000). A t the limb level, cells migrate from the lateral edge of the dermo-myotome into the growing limb bud (Chevalier et al. 1977; Jacob et al. 1977). In this case, migrating myoblasts are committed and undifferentiated (Tajbakhsh and Buckingham, 1994; Williams and Ordahl, 2000), the major difference being represented by the absence of a pre-existing muscle structure such as the myotome in the limb. In both the epaxial and hypaxial domains, myoblast proliferation results in a pool of progenitors that will differentiate into primary, secondary fibers (Kelly and Zachs, 1969) and perhaps satellite cells during later development. Therefore, at successive stages of development, each fiber may derive from fusion of myotomal myoblasts with later generations of myoblasts from the dermomyotome or even (see below) from non somitic sources (Figure 2).

The contribution of different progenitors to skeletal muscle fibers Somite

ogenitors of I* fibers ~r~ of ~g° fibe~ rived progenitors ?

P r i m a r y a n d s e c o n d a r y fibers

M a t u r e fibers

Figure 2. The contribution of different progenitors to skeletal muscle fibres. In the schematic diagram, dorso-medial progenitors (dark green) will form the myotome and later primary fibers. Progenitors of secondary myoblasts(light green) will form secondaryfibers and also contribute to growth of primaryfibers. Non-somitic, vessel-associatedmyogenicprogenitors (red) are presumedto contribute at later stages to both muscle fiber growth and to the pool of myogenic(satellite?) cells.


13t

In Drosophila lateral inhibition through Notch and Delta has been shown as the probable mechanism by which adult myogenic progenitors are selected in response to Wg signaling (Baylies et al. 1998). It thus appears likely that a similar mechanism may operate in the mammalian somite. Indeed several Delta and Notch isoforms are expressed in the somites (McGrew and Pourquie 1998)and Notch inhibits myogenesis, probably through different intra-cellular mechanisms (Wilson-Rawls et al, 1999: Nofziger et al. 1999, Hirsinger et al. 2001). However direct immunocytochernical or genetic evidence for a role of Notch in diversifying cell fate in mammalian somites is still missing. Receptors for growth factors may be pertinent targets for Notch sign~tling. In this context it has been proposed that the dorsal portion of the neural tube inhibits terminal myogenic differentiation through production of growth factors (Buffinger and Stockdale, t 995) and therefore some kind of heterogeneity may be invoked to explain the differential fate of myotomat precursors versus other precursors, similarly to what is observed between embryonic and fetal myoblasts in the developing limb bud (see below). From this point of view, it is interesting to note that the neural tube produces various FGFs (Kalcheim and Neufeld, 1990) and the first cells which form the myotome are the only myogenic cells which do not express the FGF receptor FREK (Marcelle et al. 1995). Whether a pre-selected myogenic population fails to respond to FGF, or local signaling prevents expression of FGF receptors in an homogeneous population is unknown. since the location of myotomal precursors in the dermomyotome is still unknown.

IV.

The formation of primary and secondary muscle fibers

Successive to the formation of the myotome, primary fibers begin to form both in the epaxial domain, where they probably incorporate the mononucleated myotomal cells (although this has not been demonstrated conclusively), and in hypaxial domain. where there are no pre-existing muscle cells. In both case. myogenic differentiation is not synchronous. A fraction of the myoblast population (embryonic or I ° myoblasts) withdraws from proliferation and fuse to form primary fibers, while another fraction (fetal of II ° myoblasts) does not. A possible mechanism to ensure that certain myoblasts wilt differentiate in an environment that is permissive for proliferation, may be based on the inability of these myoblasts to respond to growth factors and/or to molecules which inhibit differentiation. A few years ago, we proposed a model by which TGF[3 may influence the process of primary fiber formation in vivo (Figure 3). Committed myoblasts proliferate in the presence o f mitogens and differentiate in their absence (Hauschka, 1994). It is therefore conceivable that a gradient, of mitogen concentration is established throughout the proximo-distal axis of the g~0wing limb (reviewed in Blagden, and Hughes,1999), with the lowest concentration present at the base of the limb, just where primary fibers initiaUy form. Myoblasts proliferate in the growing limb bud, in response to different signaling molecules such as FGFs, SHH, SF/HGF (Bladt et aL 1995; Goldfarb, 1996; Duprez et al. 1998) and then differentiate at the base of the limb bud. where growth factor concentration is probably lower and/or growth factor receptors are down-regulated (Itoh et al. 1996: Flanagan-Steet et al. 2000).

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TGF 13 and muscle histogenesis = TGF I3 receptor = PKC 0 e

fetal ° fiber I ° fiber

TGFO FGF

1- truer

TGF[~ TOF~

Figure 3. A model describing fate diversification in embryonic and fetal myoblasts needed to generate primary and secondary fibers. Embryonic myoblasts 0ight green), that do not express PKC0 (left upper inset) are insensitive to TGFI3 and differentiate in the absence of mitogens. Fetal myoblasts (dark green), that express PKC0 (middle upper inset) are inhibited by TGFI3 and remain available as a pool of undifferentiated ceils that will later resume proliferation in response to growth factors (FGF and possibly several others) produced by newly formed primary fibers.

Embryonic myoblasts are insensitive to TGF~ (Cusella De Angelis et al. 1994) which is abundant in the embryonic limb (Heine et al., 1987) and differentiate into primary fibers. On the contrary, fetal myoblasts should be prevented from differentiation by TGF~ It is worth noting that TGFI3 is produced by the ectoderm and is supposed to act in a paracrine fashion on surrounding mesenchymal tissues (Francis-West and Tickle, 1996). Furthermore TGFI3 is an inhibitor of myogenesis which does not stimulate myoblast proliferation (Olson et al., 1986). Thus fetal myoblasts may enter a quiescent but undifferentiated phase, from which they must subsequently exit and begin a new wave of proliferation to generate the pool of precursor cells necessary to form secondary fibers. This proliferation may well be sustained by peptide growth factors whose relative messages are frequently localized within developing muscle fibers in the embryo (Goldfarb, 1996). Indeed co-culture experiments showed that both cultured myotubes (and explants of embryonic muscle) release into the medium mitogens which stimulate myoblast proliferation (De Angelis et al., 1992). The model above proposed has the advantage of reconciling previously unexplained data such as the presence of growth factors in newly formed muscle with the differential effect of TGF~ on embryonic and fetal myoblasts (Cusella De Angelis et al. 1994).


133

However this hypothesis still awaits in vivo functional analysis with dominant positive and negative TGF[~ signaling.

V.

A role for PKCO in myoblast diversification

The model discussed is mainly based upon the selective inhibition of differentiation caused by TGF~ or possibly other growth factors in fetal myoblasts. This may in turn be due to differential expression or activity of molecules involved in the relative transduction pathway. Protein kinase C 0 (PKCO), the PKC isoform predominantly expressed in muscle, is selectively expressed in fetal myoblasts and satellite cells. in vivo and in vitro (Zappelli et al. 1997) and not in embryonic myoblasts. If PKCO is exogenously expressed in embryonic myoblasts, that are insensitive to TGF[3. it restores the differentiation-inhibitory effect of molecule. Why differential expression of PKCO should dictate the differential fate of embryonic versus fetal myoblasts remains, at present, a matter of speculation. It is a fact, however, that only a fraction of the total myogenic precursor population differentiate at this time in the base of the limb bud. All myoblasts in this area are probably exposed to a low concentration of growth factors but to a high concentration of TGF[L At this time, embryonic myoblasts, which do not express PKCO and, thus. are insensitive to TGFfl may differentiate into primary fibers, while fetal myobtasts, which do express PKCO should be blocked b y TGF~ (Figure 3). Once primary fibers are formed, they begin to produce growth factors such as FGF and thus promote a new wave of proliferation in fetal cells. The identification of PKCO as a key component of this pathway permits the design of experiments where expression of this enzyme in vivo. driven by the promoter of genes egpressed early in the limb bud. should render all myoblasts sensitive to TGF~ and thus prevent formation of primary fibers. This possibility is currently under investigation.

VI.

Satellite cells

Satellite cells are classically defined as quiescent mononucleated cells, located between the sarcolemma and the basal lamina of adult skeletal muscle (Bischoff, 1994). They have been shown to contribute to post-natal growth of muscle fibers, whose nuclei cannot divide. At the end of longitudinal growth, satellite cell become quiescent but can be activated if the existing fibers are damaged or destroyed. In this case they undergo a number of cells divisions producing fusion competent cells, that can either fuse with damaged fibers or form new ones, and other cells that return to quiescence, thus maintaining a progenitor pool. This fact has led to the suggestion that they represent a type of stem cells (Miller et al. 1999). Although with some controversy, markers for quiescent, activated and proliferating satellite ceils have been identified (Allen et al. 1991; Irintchev et al. 1994; Allen et al. 1995; Cornelison and Wold 1997; Dominov et al. 1998: Beauchamp et al. 2000). These include desmin, M-Cadherin, c-Met, My(5, MyoD and CD34 that are expressed in quiescent cells or soon after activation. It is important to stress that all of these markers

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are also common to other classes of myogenic and some (CD34 and desmin) even to non-myogenic cells. In addition, previous work from our laboratory had identified specific features of satellite cells (i.e. resistance to phorbol esters and susceptibility to TGF[3-induced block of differentiation) that suggested they may represent a different class of myogenic cells with respect to embryonic and fetal myoblasts (reviewed in Cossu and Molinaro, 1987). Indeed, satellite ceils are the only relatively well-defined myogenic cell in post-natal life (Seale and Rudnicki, 2000). It is currently assumed but not experimentally proved that they represent a single cell type, with a common embryological origin. The origin of satellite cells is presumed to be somitic, but the evidence for this is also not conclusive due to technical difficulties in identifying quail nuclei in chick-quail chimeras at the ultra-structural level (Armand et al., 1983). Furthermore, it is surprising that despite a low number of resident, quiescent satellite cells (identified by their location and the expression of M-Cadherin) in adult healthy muscle, hundreds of activated (MyoD positive) satellite cells are seen hours after an injury to the tissue (Grounds et al., 1991). This suggests that cells are recruited to muscle regeneration from additional sources, either locally or systemically (Figure 4).

Possible origin of myogenic cells in regenerating skeletal muscle Regerated fiber Myogenic cell Satellite cell Endothelial cell Perithilial cell Circulating cell (~

Figure 4.

Dividing cell

A schematic representation of the possible origin of myogenic ceils in regenerating skeletal

muscle. Regenerated fibers (characterized by centrally located nuclei) are formed by fusion of myogenic cells (dotted green) that may derive from resident, activated satellite cells (green) of from cells derived from the micro-vasculature, either as circulating cells (red) or endothelial cells (light brown) or pericytes (violet) of the vessel wall. A green nucleus symbolizes the activation of a myogenic program.


VII.

135

Unorthodox myogenic progenitors

Why should non myogenic cells give origin to skeletal muscle? This concep!~ now accepted on the basis of recent compelling evidence, stems from old and largely ignored expe~mental evidence. One of the first example of this phenomenon was correction by fibrobtast=my0blast fusion of the genetic defect of the mdg.mouse mutant muscle fibers (Chaudad et al. 1989; Courbin et al. i989). More recently several laboratories found that cells from the dermis,: the neural tube, the thymus can: give rise tO skeletal muscle cells in vitro and in vivo (reviewed in Cossu, 1997). It is possible that exposure of a competent mesodermat cell to a myogenic community may recruit it to themyogeni c program, much as it happens during early myogenesis both in amphibians and mammals (Gurdon, 1993; Cossu et al. 1995), A search for donor tissues that may contribute myogenic cells for muscle regeneration identified bone marrow as a possible source. By tranSplanting genetically-marked bone marrow into immune-deficient mice, we showed that marrow;derived cells can migrate into areas of muscle degeneration, undergo myogenic differentiation, and participate to regeneration o f the damaged fibers. Because injury had been induced locally in the Tibialis Anterior, these myogenic progenitors must h a v e reached the site of regeneration via the general circulation (Ferrari et al. 1998). During the following years, several other reports appeared showing unpredicted plasticity of stem cells from different sources such as neural, hemopoietic and mesenchymal stem cells (Bjorson et al. 1999: Petersen et al. 1999; Gussorfi et al. 19991 Jackson et al. 1999: Galli et al. 2000: Lagasse et al. 2000; Mezey et al. 2000; Brazelton et al. 2000). These data suggest that most stem cells resident in different tissues have the same range of p!uripotency (from brain to blood and muscle; from muscle to blood; from blood to muscle and brain), and thus may be more similar to each other than previously believed. In the case of muscle, three additional and related observations appeared: i. skeletal muscle contain a fraction of poorly adhering cells that very efficiently reconstitute muscle but can also make bone and can Circulate (Lee et al. 2000: Torrente et al. 2001); ii. SP (side population) ceils and also cultured myogenic cells from muscle reconstitute the hemopoietic system (Gussoni et al; 1999; Jackson et al. 1999); iii the embryonic dorsal aorta generates clones with the typical morphology of mouse adult satellite cells that co-express myogenic and endothelial markers (De Angelis et al. 1999). Together these data suggest that at least a subset of post-natal satellite cells may be rooted in a vascular lineage (Bianco and Cossu, 1999). B y transplanting fragments Of dorsal aorta from quail donor embryos into chick embryo wing buds, we detected quail nuclei in the vessels of the connective tissue inside developing skeletal muscle as well as in and between newly formed muscle fibers (Figure 5). Because donor nuclei were also faoud in most mesodermal tissues of the host (Minasi et al in preparation). We thus propose that pluripotent, rather than myogenic progenitors may be associated with developing vascu!ature. When ingressing a developing muscle anlagen, these progenitors should find themselves into: a :muscle :field and thus adopt a satellite celt ~fatel Ir/deed, recent work on Pax7 null mice has shown that in these mice satellite cells are absent but the blood-born SP population in muscle is present: interestingly these SP cells fail to generate muscle but show an increased propensity to form Mood, suggesting 'that Pax7 restricts potential m

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Figure 5. Immunoperoxidasestaining with QCPN antibody of chick-quail chimeras sacrificedat E19-20. Quail nuclei (dark brown) ate detected within the wall of intramuscularblood vessels and in the muscle connectivetissue (black arrowheads). Quail nuclei are also seen inside cross-striatedmuscle fibers (white arrowheads).

skeletal muscle (Scale et al. 2000). When the vasculature develops inside a different tissue, vessel-associated progenitor cells may adopt the specific fate of that tissue and contribute to its histogenesis (Figure 6). Because of their origin, these cells may remain multipotent and responsive to local commitment, thus explaining the trans-determinafion potency described above. For example, cells from skeletal muscle, even in the adult, can give rise to osteoblast-like cells and even form bone. (Katagiri et al. 1994; Lee et al. 2000): Whether these vessel-associated, pluripotent vascular cells arise from a primordial pericyte, from an angioblast or from an hemangioblast, is not currently known. In the adult pluripotency has been demonstrated for both purified hemopoietic stem cells and for pericyte-derived, mesenchymal stem cells, that can give rise to osteoblasts, chondroblasts, adipocytes and, under special circumstances, even skeletal muscle (Caplan, 1991; Prockop, 1997).

VIH.

Conclusions

Because of the attention that researchers, science policy makers and media pay to stem cells and to their pluripotency, the field is already flooded with papers, reviews and commentaries. It is obvious to say that the enormous number of ongoing experiments will produce a massive amount of data documenting the complete scenario of this pluripotency, and, in the case of skeletal muscle how many different cell types are

137


The origin of pluripotent cells from the vessels

Endothelial cells

.m•l•-

1/

/

/

Hemopoietic Circulating cells: blood (muscle, brain,

~----Pericyte cel]s

(Smooth muscle, Bone, Cartilage. Fat. Striated muscle ?~

Endothelial derived pluripotent cells: (Smooth& striatedmuscle, Bone Cartilage. Tendon)

Figure 6.

A model explaining the origin of pluripotenr cells. A growing vessel, comprised of an endothelial

layer, surrounding pericytes and circulating hemopoietic cells penetrates into a developing tissue. Asymmetric mitosis, both in the endothelial and in the perithelial layer, may generate cells that leave the vessel and adopt the fate of the tissue {e.g. muscle) where the vessel has entered. They may differentiate into muscle fibers or remain as undifferentiated progenitors (i.e. satellite cells).

endowed with the potential to produce muscle. A real challenges for the future will be the identification of the signaling molecules, produced in this case by neighboring differentiating tissues and not by the embryonic organizing centers (even though the same molecules may be involved). Equally important will be the elucidation of the intracellular responding machinery that defines the competence of a given cell to respond to such molecules and thus adopt a given cell fate. New technologies should help in this identification, especially when comparison between purified and homogenous populations will be possible. However, to reach a complete understanding of the contribution of somitic and non somitic (from vessels or other structures) to skeletal muscle histogenesis and regeneration in quantitative terms an in vivo approach, through the Cre-lox technology or some form of in vivo labeling will be necessary. The outcome of such work may change our current ideas on the embryonic origin of mesodermal tissues.

IX.

Acknowledgements

The work in the Authors ~ laboratory was supported by grants from Telethon, European Communi~, Fondazione Istituto-Pasteur Cenci Bolognetti, Ministry o f UniVersity ( M ~ S T ) and Agenzia Spaziale Italiana (ASI).

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CONTRIBUTOR ADDRESSES

David A, Sassoon Associate Professor Dept. Biochemistry and Molecular Biology, Mount Sinai Medical School, 1 G. Levy Place, Box 1020, New York, NY 10029 USA Phone: 212 241-9678 Fax: 212 860-9279 e-mail: [email protected]

Giulio Cossu Stem Cell Research Institute. Dibit, H. San Raffaele, Via Olgettina 58, 20132 Milan Phone: 3902 2156 0250 Fax: 3902 2156 0220 e-mail: [email protected] and Dept. of Histology and Embryology, II Medical School, University of Rome "La Sapienza'. Via A. Scarpa 14, 00161 Rome Phone: 3906 4976 6757 Fax: 3906 446 2854 e-mail: giulio.cossu @uniroma1.it

Bradley B. Olwin 347 UCB, MCD-Biology, University of Colorado, Boulder. CO 80309 Phone: 303-492-6816 Fax: 303-492-1587 e-mail: Bradley.Olwin@ colorado.edu

Nadia Rosenthal Cardiovascular Research Center, Massachusetts General Hospital-East, Charlestown, MA 02129 USA e-mail: rosentha@ helix.mgh.harvard.edn

Michael A. Rudnicki Program in Molecular Genetics, Ottawa Hospital Research Institute, 501 Smyth Road, Ottawa, Ontario, Canada K1H 8L6 Phone: 613-739-6740 Fax: 613-739-8803 e-mail: mrudnicki @ottawahospital.on.ca 143

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Stefano Schiaffino Department of Biomedical Sciences, CNR Unit for Muscle Biology and Physiopathology, University of Padova, 35121 Padova, Italy Phone: 39.049.827-6034 Fax: 39.049.827-6040 e-mail: schiaffi @civ.bio.unipd.it

Mathew J. Thayer 3181 SW Sam Jackson Park Road, Division of Molecular Medicine NRC 311 Oregon Health Sciences University, Portland, OR 97201 USA e-mail: thayerrn@ ohsu.edu

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