1 Genetic Analysis of Skeletal Muscle Cell Fusion in Zebrafish Dissertation

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Genetic analysis of skeletal muscle cell fusion in zebrafish

Dissertation

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of The Ohio State University

Kimberly J. Hromowyk, B.S.

Molecular, Cellular, and Developmental Biology

The Ohio State University

2017

Dissertation Committee

Sharon L. Amacher Ph.D., Advisor

Denis Guttridge, Ph.D.

Jamie Jontes, Ph.D.

Beth Lee, Ph.D.

Copyrighted by

Kimberly Jane Hromowyk

2017

Abstract

Skeletal muscle fusion is a highly conserved process regulating muscle development, growth, and regeneration. Even as a well-studied process, the exact mechanisms and regulation of skeletal muscle fusion are still unknown. Utilizing the genetic and embryological advantages of zebrafish, I investigated skeletal muscle fusion function and regulation from development to adulthood. Using live imaging techniques I show zebrafish myoblast fusion utilizes a membrane dispersal mechanism similar to mechanisms used during Drosophila myogenesis. To genetically investigate skeletal muscle fusion, I generated CRISPR-Cas9-induced frameshifting alleles for zebrafish putative fusion regulators myomaker, kirrel3l, iqsec1b, and ckip-1. Only myomaker mutants disrupted embryonic myoblast fusion. Utilizing the myomaker mutant and an additional zebrafish fusion mutant, jam2a, I investigated the role for skeletal muscle fusion in muscle growth and maintenance. Here I show zebrafish muscle hypertrophic growth requires myomaker-dependent muscle fusion, but is independent of jam2a function. In addition, myomaker adults are significantly weaker than wild-type and jam2a individuals. Individual myofiber isolations reveal fusion recovery in jam2a fast and slow mutant fibers whereas myomaker mutant fibers remain mostly mononucleate. Because slow muscle fibers are mononucleate in embryos and multinucleate as adults, they must fuse post-embryonically. Unlike myoblast fusion, slow muscles do not appear to express ii myomaker during post-embryonic fusion, but myomaker expression in neighboring cells suggests that fusion may be mediated by an asymmetrical fusion event at this stage.

Finally, I examined cell and tissue morphology in wild-type and myomaker mutant adults.

In addition to being mononucleate, myomaker mutant adult muscle fibers have decreased

Laminin expression; additionally, cell proliferation and Pax7 expression are dramatically upregulated in myomaker mutant adult tissue. These phenotypes resemble muscle disease states where chronic muscle damage continuously activates the muscle regeneration program. This study reveals regulatory differences between myomaker-mediated embryonic and post-embryonic zebrafish muscle fusion and gives insight into the requirement for cell fusion in muscle growth and homeostasis.

iii

Dedication

Dedicated to my children and loving husband whose patience, support, and love made

this possible.

Acknowledgements

I would like to thank my advisor Sharon Amacher for giving me the opportunity to rotate and join her lab. Her positive attitude, encouragement, and support over the years made my graduate career such a pleasant and rewarding experience, and I could not have been happier. She always challenged me to think scientifically and helped to shape me into the scientist that I am today. I would like to thank Jared Talbot for all the training, guidance, and advice throughout my graduate career. He was always willing and happy to help with whatever I needed in the lab, and was a valuable resource for me when it came to performing new lab techniques, discussing muscle development, and analyzing data. I would also like to acknowledge Jared for generating the six1b:lyn-GFP transgenic that I used for my thesis work. I would like to acknowledge Michael Berberoglu for all the training and guidance on tissue sectioning, antibody staining on sections, and performing and analyzing needle- stick injuries. Lastly, I would like to thank the rest of the Amacher lab members that I have been lab mates with over the years, Tom, Kiel, Pooja, and Zach, for all the helpful discussions, advice, and friendship. I would like to acknowledge Brit Martin for performing the sarcomere strength test presented in Figure 2.4 D and E. The Introduction Section 1.2 and Figure 1.1 were modified from the sections that I wrote for the following publication: Li M, Hromowyk KJ, Amacher SL, Currie PD (2016) Muscular dystrophy modeling in zebrafish. Methods Cell Biol. 138, 347-380. doi: 10.1016/bs.mcb.2016.11.004.

Vita

2008-2010 ……………………………....The University of Northern Iowa

2010-2012……………………………….Bachelor of Science in Biochemistry, Biology

Minor, Winona State University

2012-Present…………………………….Graduate Research Associate, Department of

Molecular, Cellular, and Developmental

Biology, The Ohio State University

Publications

Berberoglu MA, Gallagher TL, Morrow ZT, Talbot JC, Hromowyk KJ, Tenente IM, Langenau DM, Amacher SL (2017) Satellite-like cells contribute to pax7-dependent skeletal muscle repair in adult zebrafish. Dev Biol. 424, 162-180. doi: 10.1016/j.ydbio.2017.03.004.

Li M, Hromowyk KJ, Amacher SL, Currie PD (2016) Muscular dystrophy modeling in zebrafish. Methods Cell Biol. 138, 347-380. doi: 10.1016/bs.mcb.2016.11.004.

Fields of Study

Major Field: Molecular, Cellular, and Developmental Biology

Table of Contents

Abstract ...... ii Dedication ...... iv Acknowledgements ...... v Vita ...... vi Table of Contents ...... vii List of Figures ...... viii Chapter 1. Introduction ...... 1 1.1 Zebrafish as a model organism...... 1 1.2 Zebrafish Skeletal Muscle Development ...... 2 1.3 Overview skeletal muscle cell fusion from muscle development, growth, and regeneration...... 9 Chapter 2. Myomaker-mediated muscle fusion differentially regulates fast and slow muscle fusion in zebrafish embryos and adults ...... 30 2.1 Introduction ...... 30 2.2 Methods ...... 34 2.3 Results ...... 39 2.4 Discussion ...... 48 Chapter 3. CRISPR/Cas9 mutagenesis of additional candidate zebrafish fusion regulators………………………………………………………………………………..76 3.1 Brief introduction to CRISPR/Cas9 mutagenesis ...... 76 3.2 Methods ...... 78 3.3 Generation and analysis of kirrel3l nonsense mutations ...... 79 3.4 Generation and analysis of iqsec1b and ckip-1 nonsense mutations ...... 81 3.5 Discussion ...... 84 Chapter 4. Conclusions & Future Directions ...... 94 Bibliography ...... 100

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List of Figures

Figure 1.1 Anterior to posterior development of the zebrafish myotome...... 24

Figure 1.2 Fusogenic synapse formation during FCM podosome-like invasion of FCs .. 26

Figure 1.3 Summary of vertebrate skeletal muscle fusion regulators acting at the membrane ...... 28

Figure 2.1 in vivo skeletal muscle cell fusion visualization suggests fusion occurs by small vesicle dispersal of the plasma membrane ...... 56

Figure 2.2 myomaker is expressed in embryonic zebrafish fast muscle specific cells ..... 58

Figure 2.3 Embryonic skeletal muscle fusion requires myomaker ...... 60

Figure 2.4 Muscle performance is severely compromised in myomaker mutant adults ... 62

Figure 2.5 myomaker, but not jam2a, is required for fast muscle hypertrophic growth ... 64

Figure 2.6 myomaker mutant adult skeletal muscle displays a variety of fiber sizes and mixing of fiber types ...... 66

Figure 2.7 In zebrafish adults, myomaker-dependent, jam2a-independent multinucleation occurs in fast and slow muscle fibers, although multinucleation in slow fibers is partially myomaker-dependent ...... 68

Figure 2.8 Slow muscle fibers become multinucleated 2-4 weeks post fertilization independently of myomaker expression ...... 70

Figure 2.9 myomaker mutants display weak Laminin staining correlating with increased cell proliferation ...... 72

Figure 2.10 myomakeroz17 adults have putative proliferating myoblasts and significantly increased number of Pax7-positive satellite-like cells...... 74 viii

Figure 3.1 CRISPR locus and necessary targeting components ...... 88

Figure 3.2 Identified kirrel3l mutants lack noticeable fusion defects ...... 90

Figure 3.3 CRISPR lesions and phenotype associated with iqsec1b and ckip-1 ...... 92

Chapter 1. Introduction

1.1 Zebrafish as a model organism

My thesis work utilized the genetic model organism Danio rerio, known as zebrafish. Zebrafish were first chosen as a genetic model in the late 1960s by George

Streisinger whose goal was to study vertebrate embryological development via mutational analysis (Grunwald and Eisen, 2002). Due to their high fecundity, rapid external development, and transparent embryos, zebrafish were highly amenable to forward genetic screens (Ingham, 1997). Traditionally genetic screens were performed by inducing germline mutations in male zebrafish with the alkylating agent N-ethyl-N- nitrosourea (ENU) (Solnica-Krezel et al., 1994). ENU mutagenesis was extremely successful, but required a two-generation breeding scheme to observe any mutations followed by complicated mapping procedures to identify mutations (Lawson and Wolfe,

2011). More recently, site-directed mutational strategies such as CRISPR-Cas9 have facilitated and accelerated zebrafish reverse genetics enabling gene-targeted approaches

(Auer et al., 2014; Talbot and Amacher, 2014). A gene duplication event in the teleost lineage generated a particular genetic advantage to zebrafish genetic screens (Taylor et al., 2003). Mutating a single zebrafish gene copy could reveal multiple gene functions otherwise masked by the loss of a single gene in other systems (Amores et al., 1998).

Concomitant with genetic manipulations, embryological manipulations such as injection

1 and transplantation experiments, enable gene overexpression, knockdown, and mutant rescue experiments along with genetic mosaic analysis (Westerfield, 2000). All of which are valuable for dissecting gene function. Collectively, when assessing the genetic and embryological advantages of zebrafish, they make a great model organism for studying the genetic regulation of development. For those reasons, I chose zebrafish to investigate the skeletal muscle fusion process.

1.2 Zebrafish Skeletal Muscle Development

Zebrafish skeletal muscle development, or myogenesis, occurs rapidly and is complete by 26 hours post-fertilization (hpf) (Devoto et al., 1996; Snow et al., 2008).

Similar to other model systems, zebrafish myogenesis requires cellular specification, differentiation, and fusion to establish the myotome (Rossi and Messina 2014). Once myogenesis is complete, the myotome is organized into fast muscle fibers, fibers that are fast to contract and fast to fatigue, slow muscle fibers, fibers slow to contract and slow to fatigue, and undifferentiated muscle precursor cells. The maintained muscle precursor cells contribute to post-embryonic muscle growth, aiding the transition from embryonic to mature skeletal muscle, and eventually form muscle stems cells. A basic knowledge of muscle myogenesis and myogenic regulation is key for understanding how fusion plays a role during development and post-embryonically. Therefore, I will summarize the major events governing zebrafish skeletal muscle myogenesis beginning with somitogenesis.

The major myogenic events found during zebrafish muscle development can serve as a general vertebrate model, but I will highlight and compare developmental differences between zebrafish and mammals during the muscle fusion review in Chapter 1.3. 2

1.2.1 Segmentation and Myogenic Onset

Like other vertebrates, zebrafish skeletal muscle derives from paraxial mesoderm originating from the presomitic mesoderm (PSM) (Holley, 2007). The PSM becomes segmented from head to tail into reiterated blocks of mesodermal tissue called somites, which later differentiate to become the future myotomes. The segmentation process, called somitogenesis, is regulated by pulsatile gene expression that oscillates within PSM cells with the same periodicity as somite formation, and by global gradients across the embryo, a mechanism broadly termed the ‘Clock and Wavefront model’ (Cooke &

Zeeman, 1976; Yabe and Takada, 2016). Recent advances allowing real-time visualization of oscillating gene expression in live zebrafish embryos and have shed new light on clock regulatory dynamics (Delaune et al., 2012; Shih et al., 2015; Schröter et al.,

2012; Webb et al., 2014; Webb et al., 2016; Yabe and Takada 2016). During somitogenesis, the processes of somite polarization and myogenesis are also underway, evident by spatially distinct expression of anterior and posterior somite compartment markers, some of which are myogenic regulatory factors (MRFs) (Oates et al., 2005;

Sawada et al., 2000; Kawamura et al., 2005; Windner et al., 2012; Windner et al., 2015).

MRF genes myod and myf5 are the first MRFs to be expressed in the PSM,

(Weinberg et al., 1996; Coutelle et al., 2001). Like in the mouse, function of either myoD or myf5 is required for zebrafish trunk skeletal muscle formation; myf5;myoD double mutants, but not singles, lack all trunk muscle (Hinits et al., 2011; Rudnicki et al., 1993).

As alluded to above, MRF expression is polarized within the newly formed zebrafish somite. In the anterior somite compartment, mespb functions to maintain pax3

3 expression, which prevents MRF expression, whereas in the posterior somite compartment, ripply1 functions to inhibit tbx6, which permits MRF expression

(Kawamura et al., 2005; Windner et al., 2012; Windner et al., 2015). Establishment of distinct somite compartments during somitogenesis is important for determining the three major myogenic fates found in the developing zebrafish myotome: slow muscle, fast muscle, and the external cell layer (ECL).

1.2.2 Specification and Slow/Fast Morphogenesis

Distinct from many other vertebrates, slow and fast muscle fibers are spatially segregated in the zebrafish myotome (Fig 1.1). Elegant lineage tracing experiments showed that slow muscle cell (SMC) and fast muscle cell (FMC) precursors are morphologically distinguishable even prior to somite formation (Devoto et al., 1996).

SMCs are the first muscle cells to mature and elongate into myofibers, and are derived from a medial row of myod- and myf5-expressing PSM ‘adaxial’ cells, located adjacent to the midline (Thisse et al., 1993; Devoto et al., 1996; Coutelle et al., 2001). Sonic hedgehog (Shh) signaling from the midline is both necessary and sufficient to induce

SMC fate (Currie & Ingham, 1996; Blagden et al., 1997; Lewis et al., 1999; Du et al.,

1997; Ingham & Kim 2005). One Shh target is prdm1a, which encodes a SMC transcription factor that permits expression of SMC downstream genes in part by inhibiting activity of Sox6, a FMC transcription factor that represses the SMC program

(Baxendale et al., 2004; von Hofsten et al., 2008). The FMC gene expression program requires both Sox6 function and the combined action of Myod with Pbx2/4 homeodomain

4 transcription factors at FMC-specific target promoters to drive FMC maturation (Maves et al., 2007; von Hofsten et al., 2008; Jackson et al., 2015).

Once formed, SMCs undergo a dramatic lateral migration through the somite to form the most superficial muscle cell layer (Devoto et al., 1996), although a small subset of Engrailed-expressing SMCs, referred to as muscle pioneers (Hatta et al., 1991), remains behind and forms the horizontal myoseptum that separates the myotome into dorsal and ventral halves (Fig 1.1). As SMCs migrate laterally, FMCs elongate in their wake (Cortés et al., 2003; Henry & Amacher, 2004) and are dependent upon SMCs for their timely elongation and subsequent maturation (Henry & Amacher, 2004). Unlike embryonic SMCs, which are mononucleate, FMCs fuse to form multinucleated myofibers

(Moore et al., 2007; Roy et al., 2001; Srinivas et al., 2007). The vertebrate-specific jam2a, jam3b, and tmem8c (myomaker) genes, which are expressed in FMCs and encode transmembrane proteins, are required for FMC fusion (Powell & Wright, 2011; Zhang and Roy, 2017; Di Gioia et al., 2017). In addition, the vertebrate-specific membrane localizing molecule, Myomerger (Myomixer or Minion), is also specifically expressed in fast muscle cells and required for zebrafish myoblast fusion (Shi et al., 2017).

Knockdown studies have revealed that muscle fusion factors initially discovered in mouse and flies (Rochlin et al., 2010), Crk/Crkl and DOCK1/5 (Moore et al, 2007),

CKIP-1 (Baas et al., 2012), and kirrel3l (Srinivas et al., 2007) are also implicated in zebrafish FMC fusion. The multinucleated fast muscle fibers form the bulk of the zebrafish myotome and are located medially to the superficial slow muscle fibers (Fig

1.1). Although somitic SMCs and FMCs are largely post-mitotic, a third population of

5 myogenic cells, the anterior border cells and their ECL derivatives, continue to proliferate and contribute to continual muscle growth throughout larval stages and adulthood

(Hollway et al., 2007; Stellabotte et al., 2007a).

1.2.3 Additional Zebrafish Skeletal Muscle Domains

The Dermomyotome-like external cell layer

As SMCs migrate laterally, a third population of somitic cells, the anterior border cells (ABCs; also referred to as Row 1 cells), undergo an Sdf-dependent anterior-to- lateral rotation within each somite to form the somitic external cell layer (ECL) (Hollway et al., 2007; Stellabotte et al., 2007a). ABCs are distinguished from other somitic cells by several criteria; they express markers traditionally associated with the amniote dermomyotome, such as pax3 and pax7 homologs and meox, they are proliferative, and they contribute to myogenic populations outside the myotome, including appendicular muscle, hypaxial muscle, dermal cells, and dorsal fin cells (Groves et al., 2005;

Hammond et al., 2007; Hollway et al. 2007; Stellabotte & Devoto, 2007b). In addition,

ABC’s contribute to both proliferative Pax7-positive cells in the ECL and quiescent

Pax7-positive cells in the myotome, suggesting that ABCs/ECL give rise to both muscle progenitors and satellite cells (muscle stem cells) (Hammond et al., 2007, Hollway et al.,

2007; Pipalia et al., 2016; Stellabotte et al., 2007a). Because zebrafish ABCs/ECL share many characteristics with amniote dermomyotome cells, they are also sometimes referred to as dermomyotome cells (Stellabotte et al., 2007b; Windner et al., 2012). Thus, although the terms ECL and dermomyotome are used within the literature to describe the

6 same population of cells, I use the former terminology since ECL was the name initially adopted for cells in this position (Waterman, 1969) and because is not yet known whether zebrafish dermomyotome-like cells fully mimic later characteristics of the amniote dermomyotome (Hollway et al., 2007; Stellabotte et al. 2007a; Stellabotte et al. 2007b).

Satellite-like Cells

Mammalian skeletal muscle contains quiescent pax7-positive stem cell population known as satellite cells (SCs) (Montarras et al., 2013). SCs arise from dermomyotome tissue where they contribute to the myotome during development (Bentzinger et al.,

2012), early post-natal growth (White et al., 2010; Siegel et al., 2013), and during muscle repair and regeneration (Lepper et al., 2011; Sambasivan et al., 2011). The ability to self- renew, differentiate into myoblasts and additional mesogenic tissues, and promote muscle regeneration are key characteristics of SCs (Asakura et al 2001; Dhawan and Rando

2005; Zammit et al., 2006). Zebrafish have cells characteristic of mammalian SCs, known as satellite-like cells (SLCs) (Devoto et al., 2006; Hollway et al., 2007; Stellabotte et al.,

2007a; Seger et al., 2011; Siegel et al., 2013; Berberoglu et al., 2017). Zebrafish SLCs are thought to arise from the dermomyotome-like external cell layer and contribute to the developing myotome during myogenesis (Devoto et al., 2006; Hollway et al., 2007;

Stellabotte et al., 2007a; Nguyen et al., 2017). pax7-positive SLCs function to regenerate both larval and adult muscle tissue post injury (Seger et al., 2011; Pipalia et al., 2016;

Gurevich et al., 2016; Berberoglu et al., 2017) and proliferate in response to injury (Seger et al., 2011; Berberoglu et al., 2017), suggesting SLCs function as myogenic stem cells.

However, additional SLC functional studies need to be performed to fully validate their equivalence to mammalian SCs (Siegel et al., 2013).

Hypaxial Muscle

Vertebrate hypaxial muscles derive from ventrally-located somitic cells (Burke &

Nowicki 2003; Hernandez, Patterson & Devoto, 2005). These cells migrate out of the ventral myotomes to form craniofacial, appendicular (limb/fin), and abdominal muscles

(Burke & Nowicki 2003; Hernandez, Patterson & Devoto, 2005). In zebrafish, hypaxial musculature contributing to the face (Schilling & Kimmel, 1997) and pectoral fin is specifically derived from anterior somites (Haines et al., 2004; Minchin et al., 2013; Neyt et al., 2000; Ochi & Westerfield, 2009). Jaw muscles, the sternohyoid muscle (SHM), and oesophageal striated muscle (OSM) are derived from migratory muscle precursors in somites 1-3, while pectoral fin muscles are derived from the ventral somitic region of somites 2-4 (Hollway et al., 2007; Minchin et al., 2013; Neyt et al., 2000). The posterior hypaxial muscle derived from somites 5-6 develops into a skeletal muscle that attaches to the cleithrum, a membranous bone primarily found in fish and primitive tetrapods

(Haines et al., 2004). Like mammalian hypaxial muscles, the zebrafish genes lbx2, mox2, pax3, as well as the receptor tyrosine kinase Met and its ligand Hepatocyte Growth

Factor have been linked to hypaxial muscle morphogenesis and migration (Haines et al.,

2004; Lou et al., 2012; Minchin et al., 2013; Neyt et al., 2000; Ochi et al., 2009).

Additionally, the non-canonical Wnt pathway components Wnt5, Disheveled, and RhoA are required downstream of lbx2 function (Lou et al., 2012).

1.3 Overview skeletal muscle cell fusion from muscle development, growth, and regeneration

1.3.1 General description of Skeletal Muscle Fusion

Fusion of skeletal muscle cells forms syncytial skeletal muscle tissue required for muscle development and facilitates muscle growth and maintenance. Embryonic skeletal muscle fusion involves fusion between two myoblasts, called primary fusion, or fusion between a myoblast and a mature, multinucleate myofiber or myotube, termed secondary fusion (White et al., 2010). Both processes follow the same basic fusion steps during development. The muscle fusion process begins with recognition and adhesion of fusing partners, followed by actin cytoskeleton remodeling to increase membrane proximity, and concluded with lipid destabilization and cytoplasmic merging (Aguilar et al., 2013; Kim et al., 2015). Each process requires key regulators enabling the transition to the next fusion phase. The fusion mechanism during growth and regeneration is less characterized, but still requires differentiated myoblast fusion into myotubes or with each other. To understand what is known about each of these fusion processes, I will summarize key findings from Drosophila, mouse, and zebrafish model systems that have contributed to the current knowledge of muscle fusion.

1.3.2 Specification and fusion of myoblast during muscle development

Specification of mesoderm-derived muscle precursor cells into myoblasts is required for muscle fusion during development. However, this process varies between invertebrates and vertebrates contributing to difference in myoblast fusion events. To

9 compare both myoblast fusion models, I will summarize myoblast specification and fusion events from the invertebrate model Drosophila and the vertebrate mouse model.

Specification of Drosophila Myoblasts

Drosophila skeletal muscle derives from the somatic mesoderm (Dobi et al.,

2015). Complex transcriptional networks governed by twist, in combination with Wnt,

TGF-β, receptor tyrosine kinase (RTK), and Notch signaling cascades, pattern the mesoderm into reiterative segments with distinct dorsal-ventral and anterior-posterior identities (Dobi et al., 2015). Following twist-mediated mesodermal patterning, RTK signaling localizes the transcription factor Lethal of scute (L’sc) to promuscle clusters known as equivalence groups (Dobi et al., 2015). Notch-regulated lateral inhibition restricts L’sc to a single myogenic progenitor within the group, generating a founder cell

(FC). FCs then undergo asymmetric cellular divisions to produce adult muscle precursors

(AMPs), which maintain twist expression delaying proliferation and differentiation until metamorphosis (Bate et al., 1991). The remaining cells from the equivalence group express the Gli-family transcription factor Lame duck (Lmd) and adopt the fusion- competent myoblast (FCM) fate (Taylor 2002; Dobi et al., 2015). Select transcriptional profiles established by lateral inhibition and asymmetric divisions distinguish the myoblast populations from one another. Each specified FC fuses with multiple FMCs forming 30 unique abdominal muscle fibers with distinct shapes, sizes, orientation, and myonuclei number (Rochlin et al., 2010; Abmayr and Pavlath 2012). Embryonic myoblast fusion, or primary fusion, then drives abdominal muscle formation (Rochlin et

10 al., 2010; Abmayr and Pavlath 2012). Because Drosophila specify two transcriptionally distinct myoblasts, their fusion mechanism differs from vertebrates who specify one pool of myoblasts from medoserm-derived cells.

Specification of mammalian myoblasts

Since zebrafish myoblast specification was discussed in Chapter 1.2.1, I will focus on mammalian myoblast specification here. Mammalian skeletal muscle derives from the paraxial mesoderm (PSM) and is organized through somitogenesis. Wnt-, Fgf-, and Notch-regulated oscillatory gene expression in combination with Wnt, Fgf and retinoic acid gradients, drive reiterative PSM budding, forming structures known as somites (Pourquié 2003; Buckingham 2006; Aulehla and Pourquié 2010; Bentzinger et al., 2012; Rossi and Messina 2014; Oates et al., 2012). As somitogenesis progresses, cells in older somites cease oscillatory gene expression as they exit zones of high FGF expression and enter zones of high retinoic acid concentration (Aulehla and Pourquié

2010; Oates et al., 2012; Yabe and Takada 2016). When presumptive somites exit oscillating gene expression, somite boundaries can be established (Sage 2012). Similar to the zebrafish mechanism, mammals use a combination of mesp2 and ripply2 transcription to regulate Tbx6 and Notch signaling to establish somite boundaries and polarity (Saga

2012; Winder et al., 2015). Ventrally derived somitic cells develop into the mesenchymal sclerotome domain forming cartilage and bone precursor cells. Dorsal somitic cells receive critical Wnt signals activating Sine oculus-related homeobox transcription factors 1 and 4 (Six1/4) (Buckingham 2006; Bentzinger et al., 2012; Rossi

11 and Messina 2014). Six1/4 expression generates Pax3 and Pax7-expressing dermomyotome cells, establishing the skeletal muscle fate (Buckingham 2006;

Bentzinger et al., 2012; Rossi and Messina 2014). Wnt and Sonic hedgehog (Shh) signaling activate dermomyotome-derived skeletal muscle precursors (SMPs) initiating expression of myogenic regulatory factors (MRFs) Myf5 and MyoD (Buckingham 2006;

Bentzinger et al., 2012; Rossi and Messina 2014). Muscle progenitor cells expressing the myogenic regulatory factor (MRF) myf5 then delaminate form the dermomyotome to differentiate and form skeletal muscle below the dermomyotome layer. Myogenic onset activates the MRF myoD facilitating the terminal differentiation of the muscle progenitor cells into myoblasts. The MRFs myogenin and MRF4 are expressed downstream of myf5 and myoD and ultimately drive expression of genes regulating myotube formation

(Buckingham 2006; Bentzinger et al., 2012; Rossi and Messina 2014). The expression of

Sonic hedgehog (Shh) drives primary, or embryonic, myoblast formation, which mostly take on the slow muscle fate, and FGF expression directs the formation of secondary, or fetal, myoblasts which primarily take on the fast muscle fate (Wigmore et al., 2002;

Biressi et al., 2007a; Biressi et al., 2007b; Messina et al., 2010; Rossi and Messina 2014).

In order to complete myotube formation, the slow and fast muscle cells need to undergo myoblast fusion.

It is important to understand the differences between invertebrate and vertebrate myoblast specification because the specification mechanism ultimately guides how myoblast fusion occurs. Since Drosophila specify two transcriptionally distinct myoblast populations, their fusion mechanism differs from vertebrates, where a single pool of

12 myoblasts are originally established. Additionally, although mammalian and zebrafish slow and fast muscle specification and fusion differ, the mechanisms governing vertebrate myoblast fusion are largely conserved.

Drosophila Embryonic Myoblast Fusion

Following myoblast specification FCs and FCMs are spatially organized in the somatic mesoderm. FCs reside in the most lateral myoblast layer while FCMs comprise the next several medial myoblast layers (Rochlin et al., 2010; Abmayr and Pavlath 2012).

Communication between the defined myoblast regions occurs through cellular adhesion molecules (CAMs), driving migration, recognition, and adhesion (Ruiz-Gómez et al.,

2000). FCs express the immunoglobulin (Ig) domain-containing transmembrane CAMs

Dumbfounded/Kirre (Duf) and Roughtest/IrreC (Rst) to attract the medially located

FMCs (Rochlin et al., 2010; Abmayr and Pavlath 2012; Kim et al., 2015). FMCs receive the attractant signal via the Ig domain-containing transmembrane CAMs Sticks and

Stones (Sns) and Hibris (Hbs). In both FCs and FCMs, the CAMs act redundantly, as genetic analyses revealed that loss-of-function of both CAMs is required to completely disrupt myoblast fusion (Rochlin et al., 2010; Abmayr and Pavlath 2012; Kim et al.,

2015). The most lateral FCMs fuse first followed by the medial FCMs. Duf and Rst ectopic expression sufficiently attracts FCMs to ectopic sites, but their overexpression fails to induce fusion in heterologous cells. X-ray crystallography of Duf and Sns homologues, SYG-1 and SYG-2, showed their protein interaction generating a 45 nm gap between cell membranes, but cells cannot fuse with a gap that large between membranes

(Ozkan et al., 2014; Kim et al., 2015). Therefore, the interaction between CAMs successfully drives recognition and adhesion between FCs and FCMS, but cell fusion requires additional molecules.

Following CAM-mediated cell-cell recognition and adhesion, the transmembrane receptors organize into a ring-like structure called the fusion-restricted myogenic- adhesive structure, or FuRMAS (Rochlin et al., 2010). The FuRMAS serves as a signaling center transmitting signals from adhesion molecules to intracellular proteins necessary for F-actin polymerization and organization. SH2-SH3 domain-containing proteins Dreadlocks (DOCK) and Crk, or the guanine nucleotide exchange factor (GEF)

Loner, interact with the CAMs differentially activating the nucleation-promoting factors

(NPFs) Scar/WAVE or WASP in FCs and FCMs (Rochlin et al., 2010; Kim et al., 2015).

DOCK and Crk directly activate both Scar/WAVE and WASP complexes in FCMs promoting actin polymerization through Arp2/3. Scar/WAVE and WASP Arp2/3 activation require the stabilizing factor Kette/Nap1 and the binding proteins WIP/Solitary

(Sltr) respectively (Rochlin et al., 2010; Abmayr and Pavlath 2012; Kim et al., 2015). FC actin polymerization requires only Scar/WAVE complex activation. Loner functions with the small GTPase Arf6 localizing Rac to the polymerization site (Rochlin et al., 2010;

Abmayr and Pavlath 2012; Kim et al., 2015). The GEF Mbc-Elmo activates Rac enhancing Scar/Wave activity, ultimately increasing actin polymerization through

Arp2/3. Actin polymerization generates a protrusive finger-like structure within the FCM that invades the FC. The podosome-like structure (PLS) requires the WASP-WIP complex and the FCM specific proteins Blown fuse (Blow), a WASP stabilizing protein,

14 and Drosophila group I p21-activated kinases (Dpacks) 1 and 3, which promote branched actin filaments (Rochlin et al., 2010; Abmayr and Pavlath 2012; Kim et al., 2015).

Myoblast fusion requires FuRMAS formation in combination with the PLS, known collectively as the fusogenic synapse (Fig. 1.2) (Kim et al., 2015).

The invasive PLC initiates a FC mechanosensory response. Force induced Myosin

II fusogenic synapse localization sensors PLS invasion (Kim et al., 2015). Duf-mediated

Rho1 and Rok activation recruits additional Myosin II strengthening the FC sensory response (Kim et al., 2015). Myosin II combination with myosin 18 and actin accumulation, form an actin sheath hypothesized to resist the PLS (Kim et al., 2015). The actin sheath resistance to the PLS potentially generates enough mechanical tension forcing membranes over an activation barrier enabling fusion pore formation and myoblast fusion (Kim et al., 2015). Additional research is required to validate the mechanisms in Drosophila driving plasma membrane destabilization. Following FC and

FCM fusion, FCM nuclei adopt the FC transcriptional program enabling participation in additional fusion rounds (Rochlin et al., 2010; Abmayr and Pavlath 2012; Kim et al.,

2015).

Vesicle trafficking associated with Drosophila embryonic fusion

Electron microscopy (EM) and live image analysis of Drosophila myoblast fusion reveal vesicles associated with the fusogenic synapse. Wild-type embryo EM analysis reveal the presence of vesicles, or pre-fusion complexes, along both fusing membranes associated with the fusogenic synapse (Rochlin et al., 2010; Abmayr and Pavlath 2012;

Kim et al., 2015. However, wip/sltr mutants, that disrupt actin polymerization, induce vesicle mislocalization suggesting vesicle localization and/or transports requires actin polymerization. Similarly, singles bar (sing) mutants are defective in myoblast fusion and display increased levels of both pre-fusion vesicles at the membrane and additional cytoplasmic vesicles (Rochlin et al., 2010; Abmayr and Pavlath 2012; Kim et al., 2015.

Sing contains a MARVEL domain found to function in vesicle trafficking, which supports the idea that myoblast fusion requires Sing-mediated vesicular trafficking for either vesicle localization or fusion to the fusogenic synapse in both FCs and FCMs.

Both Duf and Sns associate with either exocytic or endocytic vesicles within FCs and FMCs, respectively. Antisocial/Rolling pebbles (Ant/Rols), an ankryin repeat- containing protein, utilizes exocytic vesicles recycling Duf to FC membranes during myoblast fusion (Rochlin et al., 2010; Abmayr and Pavlath 2012; Kim et al., 2015.

Ant/Rols aids Myosin II recruitment establishing the PLS resistance and replenishes Duf protein enabling FCs to undergo multiple myoblast fusion rounds. Sns associates with endocytic vesicles during myoblast fusion. Live imaging analysis performed during fusion in the Drosophila dorsal pharyngeal musculature (DPM) reveal Rab5-containing endosomes transport Sns to lysosomes for degradation (Haralalka et al., 2014). It is hypothesized that Sns degradation helps maintain asymmetry between fusing FCs and

FMCs which is supported by Duf maintenance and stabilization in FCs.

The mechanism governing Drosophila myoblast fusion has been thoroughly investigated enabling construction of a detailed fusion mechanism. Due to the large genetic conservation of many Drosophila fusion molecules with vertebrates, the

16 identified Drosophila fusion regulators serve as a pool of potential vertebrate fusion- regulating candidates. Taking advantage of the detailed Drosophila fusion mechanism, I decided to mutate several conserved Drosophila fusion regulators and determine their function in zebrafish fusion, which is discussed in Chapter 3.

Vertebrate Embryonic Myoblast Fusion regulators

Currently there is no established vertebrate myoblast fusion mechanism as in

Drosophila. For that reason, I will be discussing molecules necessary for vertebrate muscle fusion and their putative function during the fusion process. As in Drosophila, vertebrate muscle fusion relies on transmembrane proteins guiding myoblast recognition and adhesion. The zebrafish Duf homologue, kirrel3l (Srinivas et al., 2007), and the vertebrate Sns homologue, nephrin (Sohn et al., 2009), have both been implicated in regulating vertebrate myoblast fusion, but Kirrel3l and Nephrin interaction is not required for fusion (Srinivas et al., 2007; Sohn et al., 2009). Myoblast fusion in zebrafish requires heterophilic interaction between the Ig domain-containing transmembrane proteins Jam2a and Jam3b, proposed to drive recognition between myoblasts (Powell and Wright 2011).

Similar to Drosophila CAMs, Jam2a and Jam3b are not sufficient to drive myoblast fusion (Powell and Wright 2011). However, Jam2a and Jam3b are not required for myoblast fusion in mammals (Powell and Wright 2011), suggesting additional transmembrane proteins are necessary for vertebrate myoblast fusion. Myomaker, a multi-pass transmembrane protein, is necessary and sufficient to drive myoblast fusion in mouse (Millay et al., 2012), chick (Luo et al., 2015), and zebrafish (Landemaine et al.,

2014; Zhang & Roy 2017; Di Gioia et al., 2017). Myomaker can drive fusion between myoblast and fibroblasts, but cannot induce fusion between two myomaker-expressing fibroblasts (Millay et al., 2012). Therefore, Myomaker functions in recognition and activation of downstream fusion machinery in myoblast cells, but requires additional factors to drive fusion in non-fusing cells. Myomerger (also known as Minion and

Myomixer) is a micropeptide specifically expressed in myoblasts during fusion and is necessary for myoblast fusion in mouse (Quinn et al., 2017, Zhang et al., 2017, Bi et al.,

2017) and zebrafish (Shi et al., 2017). Myomerger functions synergistically and in trans with Myomaker to drive fusion between two fibroblasts (Quinn et al., 2017, Zhang et al.,

2017, Bi et al., 2017), suggesting Myomerger is an essential Myomaker fusion partner.

Therefore, Myomaker and Myomerger together are necessary and sufficient to drive cellular fusion in cells that do not normally fuse. Myomaker and Myomerger are the first skeletal muscle fusogens discovered in mammals (Fig. 1.3).

Furthermore, mechanisms normally associated with apoptosis also promote myoblast fusion in mammals (van den Eijnde et al., 2001; Jeong and Conboy 2011;

Hochreiter-Hufford et al., 2013; Hamoud et al., 2014; Park et al., 2016).

Phosphatidylserine (PS) is a phospholipid exposed on the outer membrane during apoptosis helping to tag cells for destruction (Segawa and Nagata, 2015). The PS receptors brain-specific angiogenesis inhibitor 1 (BAI1) (Hochreiter-Hufford et al., 2013) and Stabilin-2 (Park et al., 2016) promote myoblast fusion in vivo. BAI1 and Stabilin-2 mutants have decreased muscle size and defective regeneration after injury (Hochreiter-

Hufford et al., 2013; Park et al., 2016). BAI1 PS reception recruits ELMO/DOCK

18 proteins which subsequently interact with brain-specific angiogenesis inhibitor 3 (BAI3) to promote myoblast fusion (Hochreiter-Hufford et al., 2013; Hamoud et al., 2014).

Similar to the BAI1 mutant phenotype, lack of BAI3 causes embryonic fusion defects, resulting in mononucleated myotubes in vivo (Hamoud et al., 2014) (Fig. 1.3).

In addition to known actin regulatory proteins, Casein kinase 2-interacting protein-1 (CKIP-1) (Baas et al., 2012) and GTPase Regulator Associated with Focal adhesion kinase-1 (GRAF1) (Lenhart et al., 2014) are two putative proteins in vertebrates that function to drive myoblast fusion downstream of recognition and adhesion molecules

(Kim et al., 2015). CKIP-1 localizes to the plasma membrane and interacts with actin capping proteins to help regulate actin polymerization (Baas et al., 2012). CKIP-1 knockdown in C2C12 myoblasts and zebrafish interferes with fusion between myoblasts, suggesting that CKIP-1 is necessary for actin dynamics required for vertebrate myoblast fusion (Baas et al., 2012). GRAF1 contains a BAR domain that functions in lipid-binding suggesting GRAF1 functions at the plasma membrane (Lenhart et al., 2014; Kim et al.,

2015). The plasma membrane-interacting BAR domain supports that GRAF1 functions in regulating membrane shape or structure during fusion, but the mechanisms by which it functions is still under investigation. Additionally, zebrafish knockdown of Dock1/5,

Crk/CrkL (Moore et al., 2007), or Rac1 (Srinivas et al., 2007) inhibits fast muscle elongation and fusion, which further implicate conserved actin cytoskeleton remodeling in vertebrate myoblast fusion. Recent discrepancies have surfaced between zebrafish knockdown and mutant studies, which potentially result from biological and/or mechanistic reasons (Stainier et al., 2017), but the discrepancies further motivate

19 mutational analysis to truly validate gene function. Therefore motivating the mutant screen of putative fusion regulators I discuss in Chapter 3. Thus, although several molecules have been implicated in vertebrate myoblast fusion, the complete fusion mechanism has yet to be fully elucidated (Fig. 1.3).

Although the myoblast specification and fusion mechanisms differ between invertebrates and vertebrates, there are genetically and functionally conserved molecules between each system discussed. Thus motivating the comparison and investigation of myoblast fusion in both invertebrate and vertebrate systems. Additionally, in Chapter 4, I discuss how post-embryonic fusion events in zebrafish may resemble Drosophila embryonic fusion, emphasizing the importance in understanding all types of myoblast fusion events.

1.3.3 Skeletal muscle fusion facilitating vertebrate post-embryonic muscle growth

Both mouse (Kitiyakara et al., 1963; White et al., 2010; Kostallari et al., 2015; Gu et al., 2016) and zebrafish (Patterson et al., 2008; Gurevich et al., 2015; Roy et al., 2017;

Nguyen et al., 2017) undergo post-embryonic muscle growth transitioning embryonic muscle to mature adult muscle fibers. During embryogenesis, muscle growth occurs by hyperplasia (fiber addition) and hypertrophy (fiber enlargement) (Bruusgaard et al., 2010;

White et al., 2010; Gurevich et al., 2015), which continues during zebrafish post- embryonic growth (Patterson et al., 2008; Gurevich et al., 2015; Roy et al., 2017; Nguyen et al., 2017). However, in mouse, hyperplasia and hypertrophy occur during distinct growth phases (White et al., 2010). In the extensor digitorum longus (EDL) muscle total

20 fiber number is set during post-natal week 1, while during post-natal weeks 2-17, the

EDL muscle weight increases 3.5 fold (Ontel et al, 1984; White et al., 2010), relying heavily on hypertrophy for muscle growth. Hypertrophic growth occurs in two forms, fusion-dependent growth, initiated by myoblast fusion, or fusion-independent growth, initiated by endogenous nuclei (Bruusgaard et al., 2010; White et al., 2010; Gurevich et al., 2015). In mouse, fusion-dependent hypertrophy specifically occurs during post-natal weeks 2-3 (White et al., 2010). Both mouse and zebrafish fusion-dependent, post- embryonic hypertrophic growth utilizes myoblasts derived from activated SCs (White et al., 2010; Kostallari et al., 2015; Gu et al., 2016) or SLCs (Gurevich et al., 2015; Nguyen et al., 2017), respectively . In mouse, fusion-dependent growth occurs via myoblast fusion at the ends of post-embryonic fibers (Kitiyakara et al., 1963, Gu et al., 2016). NF-

κB and Ephrin-A5 regulated NG2+ interstitial cells guide myoblasts to fiber ends, regulating post-embryonic growth (Kostallari et al., 2015; Gu et al., 2016). Stromal interaction molecule 1 (STIM1) has also been implicated in regulating post-embryonic, fusion-dependent hypertrophic growth in mouse (Li et al., 2012). However, the mechanism governing zebrafish post-embryonic muscle fusion is currently unknown, but

I discuss potential post-embryonic fusion mechanisms in Chapter 2.

1.3.4. Skeletal muscle fusion during muscle repair and regeneration

Depending on the degree of muscle damage, skeletal muscle either executes muscle repair or muscle regeneration (Hawke and Garry, 2001). Skeletal muscle repair involves myoblast fusion to the existing damaged fiber while skeletal muscle

21 regeneration involves nascent myoblast fusion to replace the damaged fibers (Hawke and

Garry 2001; Wang and Rudnicki 2012). Pax7-positive SCs or SLCs in mouse (Lepper et al., 2011; Sambasivan et al., 2011) and zebrafish (Seger et al., 2011; Gurevich et al.,

2016; Pipalia et al., 2016; Berberoglu et al., 2017), respectively, are required for muscle repair and regeneration, but the fusion mechanism by which Pax7-postive cells utilize is currently unknown. Mammalian Pax7-positive cell-mediated muscle regeneration requires Myomaker (Millay et al., 2014), and molecules regulating embryonic myoblast fusion such as BAI1 (Hochreiter-Hufford et al., 2013) and Stabilin-2 (Park et al., 2016) have been implicated in regulating skeletal muscle repair and regeneration, but the cell fusion mechanism in which these molecules function has yet to be resolved. Evidence obtain during my thesis work supports post-embryonic fusion events differ from embryonic mechanisms in zebrafish, suggesting fusion events during skeletal muscle repair and regeneration may utilize a unique fusion mechanism compared to embryonic myoblast fusion.

Skeletal muscle fusion is required for muscle development and contributes to muscle growth and maintenance. As a highly conserved process, both invertebrate and vertebrate fusion models have greatly contributed to the muscle fusion mechanism.

Utilizing the current muscle fusion knowledge, I developed a mutant screen to further understand how muscle fusion regulates muscle development, growth, and maintenance. I chose the zebrafish model organism for the embryological and genetic advantages beneficial for studying skeletal muscle fusion in a vertebrate system. Chapter 2 discusses my findings associated with mutating zebrafish Myomaker, while Chapter 3 highlights

22 my findings from mutating 3 additional putative zebrafish fusion regulators Kirrel3l,

Iqsec1b, and CKIP-1.

Figure 1.1 Anterior to posterior development of the zebrafish myotome

CONTINUED

Figure 1.1 CONTINUED (A) Lateral and (B) dorsal views of the anterior to posterior myogenic events occurring during zebrafish muscle development at 19 hpf. Each color represents a major morphogenetic or developmental event occurring in the zebrafish myotome. The yellow line traces the path adaxial cells (yellow squares) take as they mature into SMCs and migrate laterally during myogenesis. (C) Somite illustrations depicting each myogenic population SMCs (yellow), FMCs (blue), and ABCs (red) in new, intermediate, and old somites during myogenesis. Somite rotation (red arrow) occurs in new somites preceding SMC migration (green arrow) in intermediate somites.

(D) Cross-sectional illustration of a mature zebrafish myotome. FMCs (blue) comprise the medial myotome compartments while SMCs (yellow) and Pax7-positive SLCs (red) form the most superficial muscle layers. Specialized SMCs that do not migrate, muscle pioneer cells, form the horizontal myoseptum separating the myotome into dorsal and ventral halves. hpf (hours post fertilization); SMCs (slow muscle cells); FMCs (fast muscle cells); ABCs (anterior border cells); SLCs (satellite-like cells). Modified from Li et al., 2017.

Figure 1.2 Fusogenic synapse formation during FCM podosome-like invasion of FCs

CONTINUED

Figure 1.2 CONTINUED Molecular organization of adhesion and actin organizing molecules during fusogenic synapse formation in Drosophila. CAM interaction drives activation of two NPFs driving actin (red dots) polymerization through Arp2/3 complex.

Actin polymerization in FCMs (green cells) generates a PLS for FCM invasion into FCs

(Black cell). PLS invasion generates a force-mediated Myosin II (pink dots) response resisting the PLS. PLS resistance is hypothesized to generate the energy needed for membrane destabilization and subsequent cell fusion. CAM (cell adhesion molecule),

NPFs (nuclear promoting factors), FCM (fusion competent myoblast), PLS (podosome- like structure), FC (founder cell). Modified from Kim et al., 2015.

Figure 1.3 Summary of vertebrate skeletal muscle fusion regulators acting at the membrane

CONTINUED

Figure 1.3 CONTINUED (A) Summary of the membrane molecules regulating vertebrate myoblast fusion in zebrafish, chick, and/or mouse. (B-D) Membrane molecules necessary to drive fusion between two myoblasts (B), a myoblast and a fibroblast (C), and two fibroblasts (D). (C) Exogenous expression of Myomaker can drive heterologous fusion between a myoblast and a fibroblast, whereas (D) exogenesis expression of both

Myomaker AND Myomerger can induce fusion between two fibroblasts.

Chapter 2. Myomaker-mediated muscle fusion differentially regulates fast and slow muscle fusion in zebrafish embryos and adults

2.1 Introduction

As a genetically programmed process, cell fusion employs three characteristic stages: 1) cellular differentiation and fusion competence, 2) cell recognition and adhesion enabling cellular commitment, 3) lipid destabilization and membrane merging permitting cell fusion (Aguilar et al., 2013; Kim et al., 2015). Each process uses key regulators to transition to the next fusion phase. Each characteristic stage is observed during myoblast fusion (Kim et al., 2015) generating the syncytial tissue required for skeletal muscle development, growth, and regeneration (White et al., 2010; Millay et al., 2013; Millay et al., 2014; Goh and Millay 2017). Myoblast fusion has been genetically characterized in

Drosophila where fusion between transcriptionally-distinct myoblasts, founder cells

(FCs) and fusion competent myoblasts (FCMs), occurs. This asymmetric fusion requires cell adhesion molecule (CAM)-mediate recognition and adhesion, actin cytoskeleton- directed FCM protrusions into FCs, and membrane destabilization to allow cell merging

(Aguilar et al., 2013; Kim et al., 2015). Endocytic recycling of membrane proteins following cell fusion maintains FC asymmetry, enabling multiple fusion events

(Haralalka et al., 2014).

Expression and mutational analysis of vertebrate fusion regulators suggest that bona fide FCs are not present in vertebrates (Powell and Wright 2012), supporting the idea that some myoblast fusion mechanisms are vertebrate-specific. However, as in

Drosophila, myoblast-specific transmembrane proteins have been identified that are

30 involved in vertebrate myoblast fusion. Heterophilic interaction between vertebrate- specific junction adhesion molecules Jam2a and Jam3a is required for zebrafish myoblast fusion (Powell and Wright 2011). However, the mammalian orthologs, Jam-B and Jam-

C, function predominantly in cell migration, mobilization, and polarization, and do not appear to be required for muscle fusion (Powell and Wright 2011, Arcangeli et al., 2014;

Gliki et al., 2004). Myomaker, a muscle-specific multi-pass transmembrane protein, is required for skeletal muscle fusion in mouse, chick, and zebrafish (Millay et al., 2013;

Landemaine et al., 2014; Luo et al., 2015; Zhang & Roy 2017; Di Gioia et al., 2017). If expressed in non-muscle cell like a fibroblast, Myomaker can facilitate fusion with a myoblast (Millay et al, 2013). Additionally, Myomaker overexpression induces hyperfusion, suggesting Myomaker is sufficient to induce skeletal muscle fusion (Millay et al, 2013; Zhang & Roy 2017). Mutational analysis of the Myomaker protein determined that Myomaker fusogenic activity requires the C-terminus (Millay et al.,

2016), but the mechanism in which Myomaker drives fusion is still unknown. The discovery of the micropeptide, Myomerger, suggests Myomaker functions synergistically in trans- with Myomerger to drive fusion, but further investigation is required to understand how the fusogens work together (Quinn et al., 2017, Zhang et al., 2017, Bi et al., 2017). Myomaker in combination with Myomerger drive cell fusion in non-fusing cells, suggesting both proteins are sufficient for recognition, adhesion, and activating downstream fusion machinery (Quinn et al., 2017, Zhang et al., 2017, Bi et al., 2017)

Beyond muscle development, skeletal muscle fusion is required for muscle growth and regeneration (White et al., 2010; Millay et al., 2014; Goh and Millay 2017).

Muscle growth is generally categorized as either hyperplasia, the addition of new fibers, or hypertrophy, the growth of existing fibers (Bruusgaard et al., 2010; White et al., 2010;

Gurevich et al., 2015). During myogenesis, both hyperplasia and hypertrophy contribute to muscle growth. However, during murine post-natal growth fiber number is established shortly after birth and fusion-dependent hypertrophy exclusively contributes to muscle growth during post-natal weeks 2-3 (White et al., 2010). Beyond 3 weeks fusion- independent hypertrophy grows murine muscle into adulthood (White et al., 2010). Only after muscle injury or intense exercise will satellite cell-mediated muscle fusion occur, driving muscle repair or hypertrophy, respectively (Bruusgaar et al., 2010; Millay et al.,

2014, Goh and Millay, 2017). Transient myomaker expression in satellite cells is necessary both for regeneration after muscle injury (Millay et al., 2014) and hypertrophic growth after intense exercise (Goh and Millay 2017). Because myoblasts and satellite cells require myomaker for skeletal muscle fusion, myomaker can be used as a model to assess how fusion affects skeletal muscle development, growth, and regeneration.

The rapid, external development of the zebrafish embryo, continuous hyperplastic and hypertrophic muscle growth (Patterson et al., 2008; Gurevich et al., 2015; Roy et al.,

2017; Nguyen et al., 2017), and the use of satellite-like cells for some types of skeletal muscle regeneration in embryos and adults (Seger et al., 2011; Gurevich et al., 2016;

Pipalia et al., 2016; Berberoglu et al., 2017; Nguyen et al., 2017) make the zebrafish a valuable model to study skeletal muscle fusion. Additionally, multinucleate fast muscle fibers and mononucleate slow muscle fibers are spatially segregated during embryonic development (Devoto et al., 1996; Roy et al., 2001), facilitating fiber-type specific fusion

32 studies in zebrafish. Slow muscle precursors are specified medially and undergo a lateral migration forming the most superficial muscle layer (Devoto et al., 1996; Roy et al.,

2001; Talbot and Maves, 2016; Li et al., 2017). Upon their lateral migration slow muscle cells pass fast muscle precursors (Cortes et al., 2003; Henry et al., 2004); slow muscle cell migration is closely coordinated with fast muscle cell elongation, maturation, and fusion (Henry et al., 2004; Von Hofsten et al., 2008; Jackson and Ingham, 2013; Jackson et al., 2015; Li et al., 2017). Slow muscle cells remain mononucleate during myogenesis by inhibiting the fast muscle transcriptional program (Roy et al., 2001; Li et al., 2017).

Studies of zebrafish jam2a, jam3b (Powell and Wright 2011), and myomaker mutants

(Zhang & Roy 2017; Di Gioia et al., 2017) show that muscle specification, differentiation, and elongation do not require fusion. Studies have also indicated that zebrafish undergo continuous fusion-dependent hypertrophic growth into adulthood

(Patterson et al., 2008; Gurevich et al., 2015; Roy et al., 2017; Nguyen et al., 2017) and depend on Pax7-postive satellite-like cells for muscle regeneration (Seger et al., 2011;

Gurevich et al., 2016; Pipalia et al., 2016; Berberoglu et al., 2017; Nguyen et al., 2017).

In this study, I investigate how skeletal muscle fusion impacts zebrafish fast and slow muscle development by characterizing fusion mutants. I visualized myoblast fusion in vivo and show that zebrafish myoblast fusion occurs rapidly and simultaneously with myoblast elongation. By isolating and characterizing individual myofibers from wild-type and fusion mutant zebrafish at different developmental stages, I discovered a previously undescribed myomaker-dependent, jam2a-independent, post-embryonic fusion event that generates multinucleate slow muscle fibers beginning 2-4 weeks post fertilization (wpf). I

33 show fast muscle hypertrophic growth is dependent on myomaker but independent of jam2a function. Lastly, I show myomaker mutant adult muscle phenotypically resembles dystrophic and cachectic muscle. My findings reveal cellular dynamics of zebrafish myoblast fusion in vivo, show muscle fusion in embryos is genetically distinct from that in juveniles and adults, with myomaker being critical for both, and reveal that fusion- deficient zebrafish muscle has similar features to chronic muscle disease states. These data enable future studies investigating how fusion regulation differs between embryonic and post-embryonic fusion mechanisms and how the absence of fusion resembles a disease-like muscle state.

2.2 Methods

Animal Stocks and husbandry

Wild-type (AB strain), mutant, and transgenic zebrafish were raised and housed at

28.5C on a 14 hour light and 10 hour dark cycle. Embryos were collected from natural spawning of adult fish and were staged according to Kimmel et al. 1995. Zebrafish lines were maintained according to The Ohio State University Institutional Animal Care and

Use Committee (IACUC). Individual transgenic fish Tg(myog:H2B-mRFP) (Tang et al.

2016), Tg(smyhc1:EGFP)i104 (Elworthy et al. 2008), and Tg(mylpfa:lyn-Cyan) (Ignatius et al. 2012) were intercrossed to form the 3 Muscle Glow (3MG) transgenic line containing all three transgenes. The Tg(-10kb smyhc1:lyn-tdTomato) line was generated from a plasmid gifted by the Ingham Lab that had been used to generate the Tg(-10kb smyhc1:lyn-tdTomato)i261 transgenic line (Wang et al., 2001); our new derivative 34 transgenic line that was used in this work is designated Tg(-10kb smyhc1:lyn- tdTomato)oz29. Established mutant lines used in this study were jam2ahu3319 (generously provided by Gareth Powell and Gavin Wright), and the translucent Casper mutant, which is a homozygous double mutant (roya9; mitfaw2). Two new myomaker alleles, myomakeroz17 and myomakeroz25, were generated in this study and described in more detail below.

Live imaging of skeletal muscle fusion

To label all nuclei of slow and fast muscle labeled transgenic zebrafish, CFP-H2B microinjections were performed at the one-cell stage of Tg(six1b:lyn-GFP, smyhc1:lyn- tdTomato) embryos at a 5 ng/ul dose diluted into 0.2 M KCl with 0.05% Phenol Red. 19-

20 hpf embryos were mounted in 0.1% agarose with 5% Tricaine for anesthetizing the embryos during imaging. Imaging was complete in a 28.5ºC heated chamber on an Andor

Revolution WD spinning disk confocal microscope system (Inverted Nikon TiE microscope) over 4-5 hours. Images were captured at intervals of 5 or 2.5 minutes with a

40X or 60X objective, respectively, using an iXon Ultra EMCCD camera. Image and video processing were completed using MetaMorph software (Molecular Devices).

CRISPR Mutagenesis

CRISPR mutagenesis performed as previously described (Talbot and Amacher,

2014). The myomaker CRISPR target site used was 5’-GGCATTTACTCCGGCCCCAT-

3’. A 10 bp deletion allele that creates a premature stop codon at amino acid 125 of 220 was recovered and designated oz17. A 5 bp deletion allele that creates a premature stop codon at amino acid 170 of 220 was also recovered and designated oz25. Genotyping

35 oz17 and oz25 alleles was performed by High Resolution Melt Analysis (HRMA) analysis of the lesion site with the following primers, (Forward) 5’-

CGCAGCTGTGAGGATCTACC-3’ and (Reverse) 5’-

GACGTGTCTCAAACTCACCCA-3’, amplifying a 109 bp product. oz25 can also be identified by digesting the109 bp PCR product with HaeIII and only the WT amplicon will be cut into 51 and 58 bp fragments, while the mutant fragment is not cut. Mutants are also readily identified by after 24 hpf using the Tg(myog:H2B-mRFP) transgene.

In situ hybridization and Fluorescent in situ Hybridization (FISH)

In situ hybridization was performed as previously described (Jowett, 1999). The myomaker in situ probe was synthesized by PCR amplification of a 617 bp region of the

3’ UTR. The primers used were (Forward) 5’-TGGCAGACTTCACAACCTCAGA-3’

(Reverse) 5’-CAAGGGAGCTAATAATTCAGGGGGGCTAAT-3’. The T7 promoter sequence (5’-TAATACGACTCACTATAGGG-3’) was added to the reverse primer for antisense probe synthesis. The T7 probe was PCR amplified and purified with the Qiagen

MiniElute PCR Purification Kit (Cat. No. 28006). T7 RNA polymerase was use transcribe the DIG-labeled antisense riboprobe (Roche Life Science).

The FISH protocol performed on 2-4 week old larva was performed as described by Yi-Lin Yan and retrieved from the Zebrafish Information Network (ZFIN), University of Oregon, Eugene, OR 97403-5274, in the Protocols Wiki on the in situ hybridization

Techniques page: https://wiki.zfin.org/display/prot/3+color+Fluorescent+in+situ+on+sections.

Myofiber Preps

The zebrafish adult myofiber preparation protocol was performed as previously described (Horstick et al. 2013), with modifications. Adult transgenic fish were anesthetized in Tricaine. Fish were sacrificed by decapitation followed by de-finning, de- skinning, and removal of internal organs. Muscle fillets were placed in 3.125mg/mL

Collagenase IV in L15 media for 2-2.5 hours and washed twice in L15 media prior to titration with a pasteur pipet. Single fibers were selected for immediate confocal imaging in L15 media using an Andor Revolution WD spinning disk confocal microscope system

(Inverted Nikon TiE microscope). Nuclei quantifications were performed manually in

Photoshop.

Sectioning and Immunohistochemistry

Embryos, larva, and adult fish ≥ 3 months of age were processed for cryostat sectioning and antibody stained as previously described (Berberoglu et al., 2017). The following primary antibodies were used at the noted dilutions: chicken anti-GFP

(ab13970, Abcam: 1:2000), mouse anti-Pax7 (DSHB; 1:20), rabbit anti-Laminin (L9393,

Sigma; 1:200), rabbit anti-Rbfox1l (1:1000), and anti-Pcna (GTX124496, Genetex;

1:500). Pcna staining was performed as previously described (Katz et al 2016). Fiber diameter was measured in ImageJ and ≥ 80 fibers were measured from 2-3 sections for 2-

3 fish for each time point.

Myotome Nuclei Counts

Myonuclei counts were performed using Velocity software (PerkinElmer) on 1-3 dpf old zebrafish larvae containing the myog:H2B-mRFP transgene. Somites 12 and 15 were imaged and nuclei were manually labeled and counted through the entire z-stack

(1.0 um stacks) for each somite imaged (n= 6 for 24 hpf, n=8 for 48 hpf and 72 hpf). The values reported are the average of the averages for combined total nuclei from both somites 12 and 15.

Swim Tunnel Assay

Adult zebrafish (4-5 months) were tested in a swim tunnel that was adapted from that described in Gilbert et al. (2013). The swim tunnel was a 2 cm by 30 cm long tube connected to an EHEIM aquarium pump with King Instrument flowmeter that was adjusted by the use of a water valve. Each fish was acclimated to the swim tunnel for 15 minutes at the lowest flow rate of 6 cm/s. After acclimation the flow was increased 6 cm/s every 10 minutes for the endurance assay and every 1 minute for the sprint assay until fish fatigue. Fish fatigue is defined as the inability of the fish to swim and is determined when the fish is forced to the back of the tunnel for more than 5 seconds.

After the initial endurance and sprint assays, another 45 minute rest period and second endurance assay were also performed on each fish. Ucrit and Umax values were calculated as described (Gilbert, 2013).

Muscle Contractile Analysis

Contractile analysis of 3 dpf larvae was performed as described previously

(Martin et al., 2015.)

RT-PCR

Twenty embryos at 4-8 cell, 6 somite, 20 somite, and 48 hpf stages were solubilized in TRIzol (ThermoFisher) for RNA extraction. At larval stages (5 dpf, 20 embryos and 2 wpf, 12 embryos), fish were anesthetized in Tricaine and decapitated and tails were immediately placed into Ringer’s solution before Trizol solubilization. Total

RNA (2.5 ug) was purified and reverse transcribed into cDNA using SuperScript VILO

MasterMix (ThermoFisher) following manufacturer’s instructions.

Confocal Imaging

All confocal imaging was performed using an Andor Revolution WD spinning disk confocal system supported by the Ohio State Neuroscience Center Core P30

NS045758 grant. Lasers used were 405 nm, 488 nm, 561 nm, and 640nm using 10x, 20x,

40x, and 60x objectives.

2.3 Results

Time-lapse imaging reveals membrane-labeled punctae disperse between myoblast during cell fusion in zebrafish

To capture skeletal muscle fusion events in vivo, I performed confocal time-lapse imaging at 20 hours post fertilization (hpf) of embryos carrying two transgenes, one marking the plasma membrane of fast muscle precursors (six1b:lyn-GFPoz5) and the other marking cytoplasm of slow muscle cell precursors (smyhc1:td-Tomatooz29). To mark all nuclei, embryos were injected at the 1-cell stage with H2B-CFP mRNA. As expected

39 from previous studies, I do not observe slow muscle cell fusion during early embryogenesis (Roy et al., 2001), but do observe elongation and simultaneous fusion of fast muscle cell precursors, beginning soon after slow muscle cells migrate past them

(Cortes et al 2003; Henry & Amacher 2004) (Fig. 2.1 A). Fusion initiates as myoblasts intimately interact with each other until a fusion partner is chosen (Fig. 2.1 B). Fusion commitment appears as membrane coalescence between fusion-competent cells (Fig. 2.1

C-D), making individual membranes indistinguishable. Lastly, the plasma membrane breaks down and disperses into small GFP punctae (Fig. 2.1 E-G), creating a binucleate fast muscle cell (Fig. 2.1H). From fusion commitment to fusion completion, skeletal muscle fusion takes approximately 1 hour (62.5 ± 3.15 min, n=4) and I find myoblast fusion occurs before fast muscle myoblasts span the full length of the myotome.

Skeletal muscle cell fusion is disrupted in myomakeroz17 mutant embryos

To genetically characterize skeletal muscle fusion in zebrafish, I used

CRISPR/Cas9 technology to generate mutations in myomaker, a gene encoding a muscle- specific transmembrane protein necessary and sufficient for skeletal muscle fusion in mammals (Millay et al., 2013) and chick (Luo et al., 2015) and, as recently discovered, in zebrafish (Landemaine et al., 2014; Zhang & Roy 2017; Di Gioia et al,. 2017). I generated two myomaker alleles: myomakeroz17, a 10 bp deletion predicted to encode a truncated protein lacking the C-terminal two-and-a-half transmembrane domains (TM 5-

7), and myomakeroz25, a 5 bp deletion predicted to encode a very similar protein with a longer stretch of aberrant amino acids (Fig. 2.3 A, B). Because the C-terminal domain is

40 essential for Myomaker function in mice (Millay et al., 2016), I expected that both zebrafish myomaker alleles would encode nonfunctional proteins. myomaker transcript is not detected at 48 hpf in myomakeroz17 mutants, suggesting that the mutant transcript is subject to nonsense-mediated decay and providing additional evidence that oz17 is a null mutation (Fig 2.2 G). Indeed, fast muscle fusion fails in myomaker mutant embryos, and by 48 hpf, the single myonucleus in each muscle cell characteristically aligns about halfway between fiber tips (Fig. 2.3 C-D’), similar to zebrafish jam2a and jam3b fusion mutants (Powell and Wright 2011). Concomitant with fusion failure, nuclear counts reveal the myomaker mutant myotome contains fewer myonuclei than WT siblings after

24 hpf (Fig. 2.3 E). These data support the absence of myoblast fusion into existing myofibers, or secondary fusion, in myomaker mutants (Hindi et al., 2013; Matsakas et al.,

2010; Richardson et al., 2008).

myomaker is necessary for normal adult muscle size and swimming

myomakeroz17 mutant adults are viable but have severe defects. At 3 months of age, the width of the myomaker mutant body is extremely thin compared to wild-type siblings (Fig. 2.4 A, B). myomaker mutant adults also have a severe jaw phenotype (Di

Gioia et al,. 2017), making them easily identifiable from wild-type siblings. Interestingly, even though myomaker and jam2a mutant embryos have extremely similar embryonic muscle fusion defects, myomaker and jam2a mutant adults appear strikingly different, with jam2a mutants indistinguishable from wild-type, suggesting that myofiber number and/or size differs in myomaker and jam2a mutant adults. Because skeletal muscle cell

41 size correlates with force and power (Russell et al., 2000), I tested skeletal muscle strength in both myomaker and jam2a mutants using a swim tunnel assay. I find in both endurance and sprint assays, myomaker mutants are extremely weak compared to both wild-type siblings and jam2a mutants (Fig 2.4 C). jam2a mutant strength matches their wild-type siblings suggesting adult myofiber strength is independent of jam2a. To investigate whether the same discrepancy is observed at the larval stage, I performed an in vivo contractile strength assay (Martin et al., 2015) on myomaker and jam2a mutants at

3 dpf. The assay records whole body contractile forces induced at increasing frequencies over time. At 3 dpf, raw contractile force at frequencies greater than 50Hz trends lower in both mutants compared to wild-type siblings (Fig. 3 D, E), but the difference is not statistically significant (difference calculated by ANOVA). These data suggest that larval, but not adult, mononucleate myofibers can maintain relatively normal muscle strength.

Fast muscle hypertrophic growth differentially requires zebrafish embryonic fusion regulators myomaker and jam2a

To characterize myofiber size and muscle growth, I sectioned muscle at 5 days, 2 weeks, 1 month, and 3 months post fertilization in wild-type (Fig 2.5 A-D) and myomaker mutants (Fig 2.5 A’-D’). Using the 3 Muscle Glow (3MG) transgenic combination, smyhc1:EGFP (Elworthy et al., 2008), mylfpa:lyn-cyn (Ignatius et al,.

2012), and myog:H2B-mRFP (Tang et al., 2016), I was able to simultaneously visualize slow muscle fibers, fast muscle fibers, and myonuclei, respectively. Consistent with the expectation that fusion failure would result in smaller diameter fibers, I observe that

42 myomaker mutant muscle fibers are significantly smaller in diameter than wild-type siblings at every time point examined (Fig 2.5 E). In fact, myomaker fast muscle fibers take 1-3 months to achieve a fiber diameter that wild-type fibers reach in only 5 days, suggesting myomaker muscle fibers undergo minimal growth (Fig 2.5 E). A lack of fusion is also predicted to result in more mononucleate fibers per myotomal area; indeed, there are more mutant fibers per muscle cross section in myomaker mutants than wild- type siblings (5 dpf: average of 203 fibers/WT myotome section [n=13] versus 459 fibers/mymk myotome; 2 wpf: average of 345 fibers/WT myotome [n= 5] versus 651 fibers/mymk myotome [n= 3]). The presence of small smyhc1- and mylfpa:lyn-cyan- labeled fibers suggests fast and slow fibers have recently differentiated into the adult myomaker mutant myotome (Fig 2.6 B’, B’’). Compared to wild-type, the distinct boundary between fast and slow muscle domains fails to be maintained in myomaker mutants, and we observe myomaker mutant fast muscle fibers within the slow muscle domain (Fig 2.6 B’’’) and vice versa (Fig 2.6 B). The myomaker mutant slow and fast muscle domains appear physically separated from each other (Fig 2.5 D’ and Fig 2.6 B), yet myog:H2B-mRFP labeled myonuclei are located between the two domains indicating that myofibers are/were present, even if they are no longer or not expressing either of these two fiber type-specific transgenes (Fig 2.5 D’). I also analyzed histological sections of jam2a mutant adult muscle and found no differences in muscle morphology or fiber diameter (Fig 2.5 F-H). These data suggest myomaker, but not jam2a, is required for many aspects of late larval and adult muscle fiber growth.

Adult slow muscle fibers are multinucleate, indicating that they become fusion-competent at post-embryonic stages

To characterize fusion capacity of the two major muscle cell types, I isolated individual myofibers from individuals carrying slow and fast muscle-specific transgenes and quantified myonuclei number per fiber. Because slow myofibers are strictly mononucleate in zebrafish embryos, I expected to observe significant differences between myonuclei number in slow and fast muscle types. Surprisingly, I find that slow myofibers are multinucleate by adulthood, achieving a similar number of myonuclei per fiber as fast myofibers (Fig 2.7 A, B, G). I also note that there is a dramatic increase in the number of myonuclei in both fiber types between embryonic and adult stages

(compare Fig 2.3 C, C’ to Fig 2.7 A, B, G).

Fusion is partially restored in jamb2a, but not myomaker, mutant adults

The number of muscle nuclei per fiber tends to correlate with fiber cross sectional area (Bruusgaard et al., 2010). To investigate whether the difference in myofiber diameter in jam2a and myomaker fusion mutants correlates with myonuclei number, I isolated individual myofibers from 3-month old wild-type, jam2a, and myomaker mutant adults and quantified the average number of nuclei per fiber for each genotype (Fig 2.7

A-F). I observe that myomaker mutant fast myofibers remain unvariably mononucleate into adulthood, consistent with a permanent block to fusion. Although, I do observe some multinucleate myomaker mutant slow myofibers, myonuclei number per fiber is extremely low compared to both wild-type and jam2a mutant adults (Fig. 2.7 C, D, G). In

44 contrast, both fast and slow myofibers in jam2a mutants are multinucleate, containing about half the number of myonuclei per fiber compared to wild-type sibling myofibers

(Fig 2.7 E-G). These data indicate that myomaker is required post-embryonically for muscle fusion and suggest that jam2a becomes largely dispensable for fusion at some point prior to adulthood. These findings also potentially explain the post-embryonic differences in muscle fiber diameter and strength of myomaker and jam2a mutant fish.

Slow muscle fibers do not express myomaker when slow muscle fusion initiates

I performed live imaging in pigmentless juvenile zebrafish (roya9; mitfw2 double mutants, also known as “Casper”) (White et al., 2008), and observe that multinucleate slow muscle fibers begin to appear between 2 and 4 wpf (data not shown), much later than when fast muscle begins to fuse (before the end of the first day of development).

Additionally, I isolated individual myofibers from 3 week old larvae carrying slow and fast muscle-specific transgenes (Fig 2.8 A-B’) and show that many slow muscle fibers are multinucleate by 3 wpf (Fig 2.8 B, B’). To investigate whether slow muscle cells express myomaker during the time of slow muscle fusion initiation, I performed fluorescent in situ hybridization at 2 and 4 wpf (Fig 2.8 C-D’’’). At both time points, I observe that although myomaker does not appear to be expressed in slow muscle fibers (Fig 2.8 C-

D’’’), it is expressed in nearby cells (Fig 2.8 C’-D’’’). These data suggest that slow myofiber multinucleation occurs as neighboring myomaker-expressing progenitor cells fuse into differentiated myomaker-negative slow muscle fibers. Thus, the addition of new

45 nuclei to both the slow and fast muscle fibers in juvenile zebrafish may be similar to what has been described in mammals.

myomaker mutant muscle has reduced basal lamina and increased proliferation

Since adult myomaker mutant muscle fiber fusion defects correlate with reduced muscle strength, I investigated whether myomaker mutant fibers showed signs of structural weakness or damage by assessing Laminin expression in the extracellular matrix, a commonly used assay for muscle integrity (Acharyya et al., 2005; He et al.,

2013). I find that Laminin expression is substantially decreased in both fast and slow muscle domains of myomaker mutant adult muscle compared to wild-type muscle (Fig

2.9 A-B’’). Because decreased Laminin is associated with damaged and regenerating tissue (Acharyya et al., 2005; He et al., 2013), I assessed cell proliferation using expression of PCNA, an established proliferation marker (Dray et al., 2015; Panza et al.,

2015; Katz et al., 2016), in myomaker mutant and wild-type adult muscle (Fig 2.9 C-E). I observe over one hundred-fold more proliferating cells in muscle sections of myomaker mutant adults than their wild-type siblings (Fig 2.9 E), though the identity of these proliferating cells is unknown. I postulate that the highly proliferative state in myomaker mutant muscle reflects activation of a muscle regenerative response.

The number of Pax7-positive cells is increased in myomaker mutant skeletal muscle

To assess whether muscle regeneration is activated in myomaker mutant adults, I quantified the number of Pax7-positive cells. Pax7 reliably labels satellite cells and is

46 required for satellite cell specification, maintenance, and muscle regeneration in both mammals and zebrafish (Seale et al., 2000; Seger et al., 2011; Gunther et al., 2013; von

Maltzahn et al., 2013; Gurevich et al., 2016; Pipalia et al., 2016; Berberoglu et al., 2017;

Nguyen et al., 2017). Our previous work demonstrated that Pax7-positive satellite-like cells are relatively rare in uninjured zebrafish adult muscle, though they are concentrated in the slow muscle domain, and that they increase dramatically upon muscle injury

(Berberoglu et al., 2017). I utilized two different methods to identify Pax7-positive satellite-like cells, a pax7a:GFP transgene (Seger et al., 2011) and the Pax7 antibody.

Because pax7a:GFP-driven GFP perdures longer than Pax7 protein (Berberoglu et al.,

2017), I infer that Pax7-positive, GFP-positive cells identify satellite-like cells, whereas

Pax7-negative, GFP-positive cells identify cells that have become activated muscle precursors and are no longer quiescent. These two pax7 markers were used in conjunction with two muscle differentiation markers: myog:H2B-mRFP (Tang et al., 2016;

Berberoglu et al., 2017) which identifies differentiated myonuclei, and Rbfox1l

(Gallagher et al., 2011; Berberoglu et al., 2017), which marks myoblast nuclei and differentiated myonuclei. In myomaker mutants, pax7a:GFP expression is dramatically increased compared to wild-type siblings (Fig. 2.10 A,B), and there is a 7-fold increase in the number of Pax7-positive cells at the horizontal myoseptum (a slow muscle domain) and in the fast muscle domain (Fig. 2.10 H). I also observe cells that co-express Rbfox1l and pax7a-GFP but only weakly express Pax7 and/or myog:H2B-mRFP (Fig 2.10 C’-

G’), suggestive of proliferating myoblasts (Berberoglu et al., 2017). The dramatic expression of the pax7a:GFP transgene implies widespread satellite-like cell activation in

47 myomaker mutants and the presence of pax7a:GFP-positive, Rbfox1l-positive cells suggests that these satellite-like cells can successfully become proliferating myoblasts.

2.4 Discussion

Zebrafish myoblast fusion facilitates fast muscle elongation following slow muscle migration

Previous studies have imaged zebrafish myoblast fusion mosaically during development (Snow et al., 2010) and during larval regeneration (Pipalia et al., 2016).

Myoblasts are thought to undergo 3 dynamic morphogenetic phases during development, defined by the presence of short precursor cells, intercalating and elongating myoblasts, and myotube formation with subsequent muscle fusion (Snow et al., 2010). However, when observing myoblast fusion live, I observed fusion prior to elongation and myotube formation. I find that during the intercalation and elongation phase, myoblasts identify fusion partners and undergo fusion likely mediating the elongation process. Because I cannot predict fusion partners there is the possibility that some fibers may not discover their fusion partners until after elongation. Further investigation into fusion commitment will help clarify why some mononucleate myotubes were previously observed (Snow et al., 2008). Fast muscle fusion is known to correlate with slow muscle migration (Devoto et al., 1996; Henry and Amacher 2004; Cortes et al., 2003), but the precise timing of fast muscle activation, elongation, and fusion relative to slow muscle migration was unknown. I verify fast muscle precursors begin to fuse after slow muscle cells migrate past them, in agreement with previous data showing a single transplanted slow muscle 48 fiber is sufficient to locally rescue myoblast elongation (and likely fusion) in slow muscle-deficient embryos (Henry and Amacher 2004). Even though the time to fusion commitment, when two cells begin to coalesce membranes, varies after slow muscle migration (unpublished observations), myoblasts rapidly fuse within approximately one hour which is one fourth the time it takes chick myoblasts to fuse (Sierio-Mosti et al.,

2014).

Of the three basic fusion steps, membrane merging is the least understood. Using electron microscopy (EM), studies have investigated membrane merging during myoblast fusion (Shamada 1971; Lipton and Koningsberg 1972; Rash and Fambrough 1973; van

Raamsdonk et al 1974; Fumagalli et al., 1981; Doberstein et al., 1997). In these studies, membranes were described to fuse either by fusion pore formation and subsequent pore widening (Lipton and Koningsberg 1972), or by plasma membrane breakdown and dispersal (Shamada 1971; Rash and Fambrough 1973; van Raamsdonk et al 1974;

Fumagalli et al., 1981; Doberstein et al., 1997). My in vivo imaging data are consistent with the latter idea. I see membrane-labeled fractions disperse at the site of membrane contact, suggesting that membranes between fusing cells are removed as the cytoplasm from both cells mix. A similar dispersal of membrane-labeled puncta between fusing cells is observed during Drosophila myoblast fusion in vivo (Haralalka et al., 2014).

Additionally, EM studies have observed multiple cytoplasmic bridges between fusing cells along with membrane fractions within the cytoplasm of myotubes (Rash and

Fambrough 1973; Fumagalli et al., 1981). I hypothesize myoblast fusion requires plasma

49 membrane removal between fusing cells rather than membrane merging through fusion pore formation.

Fusion mechanisms change during zebrafish development

Slow muscle precursors inhibit the fast muscle differentiation program early in myogenesis (Von Hofsten et al., 2008, Yao et al., 2013), preventing fusion-promoting gene expression in slow muscle cells (Powell and Wright 2011; Li et al., 2017) (Fig S1

F). Contrary to their mononucleate nature in embryos, slow muscle cells become multinucleate during juvenile and adult stages. My inability to detect myomaker expression in slow muscle fibers combined with slow muscle fusion independence of jam2a, suggests juvenile slow muscle fusion utilizes a fusion mechanism distinct from that used in embryos. Since myomaker is required for post-embryonic fusion, I propose that an asymmetric fusion mechanism that requires myomaker expression in only one of the fusing cells drives both slow and post-embryonic muscle fusion. Activated satellite cells transiently express myomaker during mammalian adult muscle regeneration and after exercise (Millay et al., 2014; Goh and Millay 2017). I hypothesize satellite-like cells express myomaker during juvenile growth, which is sufficient to drive asymmetric fusion with slow muscle cells. It is interesting that zebrafish slow muscle contains about 20-fold more satellite-like cells than fast muscle (Berberoglu et al., 2017), which may explain why adult slow myofibers have similar numbers of myonuclei as fast myofibers, even though fusion initiates much later, and why myomaker mutant slow fibers occasionally do have more than one nucleus.

It is known that many fish undergo continuous hypertrophic growth (Weatherley et al., 1988; Koumans and Akster 1995; Nguyen et al., 2017), suggesting proliferating satellite-like cells are present during the time of slow muscle fusion. However, juvenile slow muscle cells must undergo a genetic or developmental change to become fusion- competent, a process that does not occur in embryos. Interestingly, a transgenic Smad3 reporter revealed that expression of Smad3, which is activated by TGFb signaling to regulate skeletal myogenesis and growth (Ge at al., 2011), initiates in slow muscle cells around 1 month post-fertilization (Casari et al., 2014). Smad3-deficient mammalian myofibers display an atrophic phenotype accompanied by insufficient muscle fusion and decreased expression of fusion-regulating genes (Ge at al., 2011). Thus it is intriguing to speculate that the expression of Smad3 in slow myofibers may induce fusion-regulating genes that permit fusion competence.

If slow muscle fibers undergo muscle fusion during post-embryonic growth, why don’t they undergo fusion during embryonic development? During zebrafish myogenesis, there are 3 distinct muscle cell compartments: slow muscle, fast muscle, and anterior border cells (ABCs) (Hollway et al., 2007, Stellabotte et al., 2007). ABCs become the most superficial compartment and form the external cell layer (ECL). ECL cells express

Pax7 and are the source of satellite-like cells (Hollway et al., 2007, Stellabotte et al.,

2007). I hypothesize that non-fusogenic slow muscle cells that migrate laterally form a

“protective” layer between the ECL and fusogenic fast muscle cells, establishing a stem cell niche during post-natal muscle growth to provide a favorable environment for

51 satellite-like cell formation. Once the satellite-like cell population is established, I propose that slow muscle fibers can then become fusogenic.

Zebrafish hypertrophic growth requires myomaker-dependent muscle fusion, but is independent of jam2a function

Rat and mouse muscle fibers utilize different combinations of hyperplasia and hypertrophy during muscle development, but both rely solely on hypertrophy after fiber number is established (Kelly, 1978; White et al., 2010). Fish, however, are known to add new myofibers throughout their life (Weatherley et al., 1988; Koumans and Akster 1995;

Nguyen et al., 2017), but it is not known whether the fish myotome can properly grow in the absence of fusion-dependent hypertrophy. My data strongly suggests zebrafish muscle requires satellite-like cell-mediated muscle fusion for hypertrophic growth. myomaker functions in satellite cell-directed regeneration and muscle-overload induced hypertrophy in mice (Millay et al., 2014; Goh and Millay 2017), suggesting satellite-like cell-mediated repair and growth is also disrupted in zebrafish myomaker mutants.

Although jam2a mutants have disrupted muscle fusion during embryonic development, post-embryonic fusion appears relatively normal and correspondingly, there is no hypertrophic phenotype in jam2a mutant adults. Adult jam2a mutant myofibers contain only half the nuclei of wild-type fibers, suggesting half the normal nuclei number is sufficient to support muscle growth in zebrafish. However, myomaker mutant satellite- like cells cannot induce hypertrophic growth, generating the growth defect in mutant adults.

The present study suggests that skeletal muscle hypertrophic growth requires satellite cell contribution. In vertebrates, it has long been debated whether hypertrophic growth requires contribution from satellite cells (Bruusgaard et al., 2010; Blaauw and

Reggiani 2014). Since myomaker mutants are deficient in fusion, and fiber growth fails to occur in mutants, this suggests that there is a connection between fusion and growth.

Since satellite cells are thought to contribute new nuclei to fibers via fusion

(Pallafacchina et al., 2013; Blaauw and Reggiani 2014; Goh and Millay 2017), then my results support the model wherein hypertrophic growth requires satellite cell fusion. In agreement, Goh and Millay (2017) showed that hypertrophic growth is defective in adult mice with a conditional myomaker knockout in satellite cells.

There are conflicting results among published zebrafish myomaker mutant adult phenotypes. In one study, myomaker mutant adults exhibited size and jaw phenotypes that are consistent with those I report here (Di Gioia et al., 2017). However, in another study, the authors report no overt adult phenotype in a myomaker deletion allele, myomakersq36 (Zhang and Roy, 2017). In this latter study, there was also no detectable difference in swimming performance of juvenile (1 month old) myomaker mutants versus wild-type siblings. I note that the latter study is not entirely comparable to our study. At one month, we are only just able to phenotypically distinguish myomaker mutant juveniles from their wild-type siblings, so I performed swimming tunnel tests in adults

(>3 months) when the mutant phenotype is obvious. Although it is formally possible that partial truncation alleles have a more severe phenotype than the myomakersq36 deletion allele (transcript and protein null), I feel it is unlikely for two reasons. First, I do not

53 detect any phenotype in myomakeroz17 heterozygotes, suggesting that the putative truncation alleles do not have dominant, neomorphic, or dominant-negative functions.

Second, I do not detect any myomaker transcript in myomakeroz17 mutant embryos, suggesting that mutant transcript is rapidly degraded by nonsense-mediated decay and is thus a likely protein null allele. It is possible that phenotypes in myomakersq36 adults arise later in life, or that differences in genetic background among the mutant strains affect phenotypic severity.

myomaker mutant adult skeletal muscle displays features commonly observed in diseased muscle

Two hallmark characteristics of skeletal muscle disease and muscle wasting are muscle damage and satellite cell activation (Tisdale 2002; Wallace et al., 2009;

Tabebordbar et al., 2013; He et al., 2013). myomaker mutant adult skeletal muscle is

Laminin-deficient, and the number of both proliferative cells and satellite-like cells is significantly increased. Damaged muscle may result from the inability of weak myomaker mutant myofibers to maintain normal swimming, but the source of the damage signal is currently unknown.

The fact that I see 7-fold more Pax7-positive cells in myomaker mutant than wild- type adults suggests that the regeneration program is activated and upregulated in myomaker mutant adults. I detect cells that express both pax7a:GFP and Rbfox1l, but not

Pax7, suggesting that satellite-like cells are able to become proliferating myoblasts, but the lack of overlap with myog:H2B-mRFP may indicate a defect in the transition from

54 proliferative myoblasts to differentiated myoblasts. In muscle wasting models, myoblasts that fail to fuse into myotubes contribute to disease progression (He et al., 2013). Yet the presence of small, differentiated fibers in myomaker adult muscle suggest that new fibers are added. However, I cannot rule out the possibility that mutant fibers are undergoing atrophy. To distinguish between a muscle wasting versus dystrophic phenotype, further investigation is required. It is known that immune cells accumulate in dystrophic muscle

(Wallace et al., 2009; Tabebordbar et al., 2013), whereas immune cell infiltration is not characteristic in muscle wasting models (He et al., 2013). Therefore, I know the regeneration program is successfully activated in myomaker mutants, but I cannot distinguish between a muscle wasting versus dystrophic phenotype. Either way, in the absence of skeletal muscle fusion, adult skeletal muscle recapitulates features of diseased-like muscle.

Figure 2.1 in vivo skeletal muscle cell fusion visualization suggests fusion occurs by small vesicle dispersal of the plasma membrane

CONTINUED

Figure 2.1 CONTINUED (A-H) Frames from a time-lapse movie of a 20 hpf six1b:lyn-

GFP (green); smyhc1:td-tomato (blue) double transgenic embryo injected with mRNA encoding H2B-CFP (red). The movie which spans 92.5 minutes, was taken at a developmental time when fusion is actively occurring. Illustrations (A’-H’) depict a pair of fusing cells from the movie (A-H; white arrowheads). The time stamp of each frame is indicated.

Figure 2.2 myomaker is expressed in embryonic zebrafish fast muscle specific cells

CONTINUED

Figure 2.2 CONTINUED In situ hybridization using an antisense myomaker (tmem8c) probe at 10-somite (A, A’) and 12-somite (B, B’) stages reveals that myomaker expression in fast muscle myoblasts begins sometime between these two stages (around

14.5-15 hpf). (C-D’) Onset and fast muscle-specific myomaker expression appears the same in control DMSO-treated embryos (C, C’) and embryos treated with the

Smoothened inhibitor cyclopamine (CyA) between 5.5-19 hpf to prevent slow muscle formation (Barresi et al. 2001, Peterson and Henry 2009) (D, D’) (lateral views in C, D; dorsal views in C’ and D’). (E-F’’) At 19 hpf, myomaker expression is restricted to fast muscle cells (E-E’’), but in prdm1anrd mutant embryos (F-F”), myomaker expression is also detected in adaxial cells located adjacent to the midline (black arrow). Adaxial cells are normally fated to become slow muscle precursors (Devoto et al. 1996), but are transfated to fast muscle in prdm1anrd mutant embryos (lateral views in E and F; dorsal views in E’-F’’). (G) RT-PCR expression of myomaker and jam2a in WT and myomakeroz17 mutant individuals at 48 hpf. Lack of myomaker expression in the mutant sample verifies that the mutant transcript cannot be detected, suggesting rapid mRNA decay occurs. The boxed region in E’ and F’ is magnified in E’’ and F’’, respectively.

Scale bar in B’ (for A-B’), in D’ (for C-D’), in F’ (for E-F’’), and in F’’ (for E’’ and F’’) are 100 µm.

Figure 2.3 Embryonic skeletal muscle fusion requires myomaker

CONTINUED

Figure 2.3 CONTINUED (A) The DNA lesions for myomakeroz17 and myomakeroz25 (red text) induced at the CRISPR target sequence (bold text). (B) The frame-shifting mutations are predicted to introduce 4 and 30 aberrant amino acids (grey text) respectively, followed by a premature stop codon (*), disrupting the protein in transmembrane (TM) domain 5. The mutant Myomakeroz17 protein is predicted to lack half of TM domain 5, all of TM domains 6 and 7, while both proteins lose the critical C- terminal fusogenic elements (Millay et al. 2016). (C-D’) Wild-type (WT) embryos (C,

C’) contain multinucleated fast fibers at 48 hpf, while fast fibers in myomakeroz17 mutant embryos (D, D’) are mononucleate, with the single nucleus in each fiber located about midway between fiber tips. myog:H2B-mRFP (red) mark muscle nuclei, mylfpa:lyn-cyan

(green) labels fast muscle, and smyhc1:EGFP (blue) labels slow muscle cells. (E) The number of total myonuclei within 2 myotomes (12 and 15) is reduced in myomaker mutants compared to WT siblings ≥ 48 hpf. (Student’s t-test, p*** < .001). Scale bar in

D’ (for C-D’) is 100 µm.

Figure 2.4 Muscle performance is severely compromised in myomaker mutant adults

CONTINUED

Figure 2.4 CONTINUED (A,B) Dorsal view of WT (A) and myomaker mutant (B) fish displaying the significant difference in muscle mass between adults. jam2a individuals are indistinguishable from wild-type. (C) Swim tunnel assays to assess endurance and sprint capacity were performed on jam2ahu3319 mutant and wild-type sibling adults (n=6 each) and myomakeroz17 mutant and wild-type adults (n=5 each) (3-6 months) (D,E)

Assays of muscle-specific force in 3 dpf myomaker mutant and sibling control larvae (D)

(n=10-3 (WT) n= 10-3 (myomaker)) (E) and jam2a mutant and sibling control larvae

(n=4 (WT), n=6 (jam2a)) (D,E) indicate no significant force difference among comparable genotypes. Although muscle-specific force at higher frequencies appears lower in both mutants compared to their respective wild-type siblings, the trend is not statistically significant. (Student’s t-test, p*** < .001).

Figure 2.5 myomaker, but not jam2a, is required for fast muscle hypertrophic growth

CONTINUED

Figure 2.5 CONTINUED Transverse sections through the mid-trunk region of wild-type

(WT) (A-D) and myomakeroz17 (A’-D’) 3MG transgenic individuals at 5 dpf, 2 wpf, 1 month, and 3 months. myog:H2B-mRFP (red) labels myonuclei, mylfpa:lyn-cyan (green) marks fast muscle cells, and smyhc1:EGFP (blue) marks slow muscle cells. Insets show magnified images of regions indicated by the dotted box. (E) Average fiber diameter of

100+ fast muscle myofibers in WT and myomakeroz17 individuals at the same stages. (F-

G) Transverse sections through the mid-trunk region of WT (F) and jam2ahu3319 (G) 3- month old adult fish showing Rbfox1l immunoreactivity. (H) The average fiber diameter of 3-month old wild-type and jam2ahu3319 adult muscle fibers near the horizontal myoseptum is indistinguishable. Student’s t-test (p*** < .001) Scale bar in A’ (for A-A’) and B’ (for B-B’) is 50 µm; scale bar in D’ (for C-D’) and in G (for F,G) is 100 µm.

Figure 2.6 myomaker mutant adult skeletal muscle displays a variety of fiber sizes and mixing of fiber types

CONTINUED

Figure 2.6 CONTINUED (A-B) Transverse sections of adult wild-type (A) and myomaker mutant (B) 3MG skeletal muscle. myog:H2B-mRFP (red) labels myonuclei, mylfpa:lyn-cyan (green) marks fast muscle cell membranes, and smyhc1:EGFP (blue) marks slow muscle cells. White arrowhead in (A) indicates a smyhc1:EGFP positive cell.

(B’-B’’) zoomed views of variable fibers sizes in slow (B’) and fast (B’’) myofibers and fast muscle fibers deep in the slow muscle domain (B’’’). Scale bar in B (for A-B) 100

µm and the scale bar in B’’’ (for B’-B’’’) is 15 µm.

Figure 2.7 In zebrafish adults, myomaker-dependent, jam2a-independent multinucleation occurs in fast and slow muscle fibers, although multinucleation in slow fibers is partially myomaker-dependent

CONTINUED

Figure 2.7 CONTINUED (A-F) Individual myofibers were isolated from wild-type (A-

B), myomaker mutant (C-D), and jam2a mutant (E-F) adults. In A-F, fiber type and myonuclei are distinguished by transgenes (myog:H2B-mRFP [myonuclei, red], mylfpa:lyn-cyan [fast fibers, blue], and smyhc1:EGFP [slow fibers, green]). (G) Box plots indicating the range and average nuclei per fiber for fast (blue) and slow (green)

WT, jam2a, and myomaker adult myofibers. Outliers are defined as >1.5x the interquartile range (IQR). Scale bars in D (for A-D) and in F (for E, F) are 100 µm.

Figure 2.8 Slow muscle fibers become multinucleated 2-

4 weeks post fertilization independently of myomaker expression

CONTINUED

Figure 2.8 CONTINUED (A-B’) Individual fast muscle (A-A’) and slow muscle (B-B’) fibers were isolated from 3 week old WT larvae. Fiber types were distinguished by transgenes mylfpa:lyn-cyan (fast fiber membranes, blue) and smyhc1:EGFP (slow fibers, green). (A’-B’) Myonuclei labeled with myog:H2B-mRFP indicating the fusion status of each fiber. (C-D) Transverse sections of Fluorescent in situ Hybridizations (FISH) of 2 week (C) and 4 week (D) WT larvae labeling smyhc1 (green) and myomaker (red). 2 week merge (C’) and 4 week merge (D’) insets show lack of co-localization (white arrows) between smyhc1 (C’’ and D’’) and myomaker (C’’’ and D’’’).

Figure 2.9 myomaker mutants display weak Laminin staining correlating with increased cell proliferation

CONTINUED

Figure 2.9 CONTINUED (A-B”) Transverse sections of WT (A-A’’) and myomaker mutant (B-B’’) adults, near the horizontal myosteptum (HM), labeled with Laminin

(green). Magnified views of boxed regions in A and B show fast muscle (FM) (A’ and B’

[white box]) and slow muscle (SM) (A’’ and B’’ [yellow box]) regions of WT and myomaker individuals respectively (C, D) Transverse sections of myog:H2B-mRFP transgenic WT (C) and myomaker mutant (D) adult skeletal muscle, near the HM, antibody stained for PCNA. myog:H2B-mRFP (red) labels myonuclei while PCNA marks proliferating nuclei (green). (C’ and D’) Magnified images of boxed regions in C and D.

(E) The average number of PCNA-positive cells per nm2 near the HM in WT versus myomaker mutant adults. Student’s t-test (p*** < .001). Scale bar in B (for A and B) and in D (for C and D) is 100 µm, scale bar in B’’ (for A’-B’’) is 25 µm, and in D’ (for C’-

D’) is 15 µm.

Figure 2.10 myomakeroz17 adults have putative proliferating myoblasts and significantly increased number of Pax7-positive satellite-like cells.

CONTINUED

Figure 2.10 CONTINUED (A, B) Transverse sections of pax7:GFP (green); myog:H2B- mRFP (magenta) transgenic WT (A) and myomaker mutant (B) adult skeletal muscle co- labeled for Pax7 (red) and Rbfox1l (blue) nuclear antigens. pax7a:GFP (green) marks satellite-like cells (SLCs) as well as cells that have begun to differentiate; pax7a:GFP- positive SLCs can be unambigously identified by their Pax7-positive nuclei. myog:H2B- mRFP (magenta) marks differentiated myonuclei; expression of myog:H2B-mRFP and pax7:GFP transgenes rarely overlap in wild-type adult muscle (Berberoglu et al. 2017).

(C-G and C’-G’) Magnified views of boxed regions in A and B show the merge (C, C’) and individual laser channels Pax7 (D, D’), pax7a:GFP (E, E’), myog:H2B-mRFP (F, F’) and Rbfox1l (G, G’), in wild-type (C-G) and myomaker mutant (C’-G’) adult muscle. (H)

The average number of Pax7-positive cells per µm2 near the horizontal myoseptum (HM) and in dorsal fast muscle in wild-type versus myomaker mutant adults. Student’s t-test

(p*** < .001). Scale bar in B (for A, B) is 100 µm and in G’ (for C-G’) is 25 µm.

Chapter 3. CRISPR/Cas9 mutagenesis of additional candidate zebrafish fusion regulators

3.1 Brief introduction to CRISPR/Cas9 mutagenesis

Due to work in mammalian cell culture and zebrafish knockdown studies, many

Drosophila fusion regulators have been implicated to function in vertebrate muscle fusion (Rochlin et al., 2010). To test whether these fusion regulators identified in the invertebrate fly model also function in vertebrates, I performed a CRISPR-Cas mutagenesis in zebrafish of several candidate fusion regulators. The clustered, regularly interspaced, short palindromic repeats (CRISPR) and CRISPR-associated system (Cas) is utilized as a bacterial and archaea defense mechanism against invading nucleic acids, such as viruses or plasmids (Fineran and Charpentier, 2012; Barrangou, 2013; Barrangou and Marraffini, 2014; Charpentier, 2015). The CRISPR-Cas system originates from a

CRISPR loci that contains three essential components: 1) short palindromic DNA repeats

(CRISPR repeats) interspaced by 2) variable sequences derived from invading nucleic acids (CRISPR spacers) and 3) Cas-associated genes (Barrangou and Marraffini, 2014;

Ma et al., 2014) (Fig 3.1 A). From the CRISPR loci, transcription and RNA processing generates short CRISPR RNAs (crRNAs) that guide Cas proteins to invading nucleic acids for double-stranded cleavage and silencing (Fineran and Charpentier, 2012;

Barrangou, 2013; Barrangou and Marraffini, 2014) (Fig 3.1 B). The crRNA contains one

CRISPR repeat that folds into a hairpin and one CRISPR spacer sequence. The hairpin interacts with Cas while the spacer guides Cas to the target site (Fineran and Charpentier,

2012; Barrangou, 2013; Barrangou and Marraffini, 2014) (Fig. 3.1 C). Because the spacer

76 is derived from an exogenous nucleic acid, the crRNA will target the complementary spacer sequence, thus targeting exogenous nucleic acids (Fineran and Charpentier, 2012;

Barrangou and Marraffini, 2014). The sequence recognized by the crRNA is referred to as the protospacer and requires a highly conserved, adjacent 2-3 nucleotide sequence known as the protospacer adjacent motif (PAM) (Mojica et al., 2009; Barrangou and

Marraffini, 2014). The PAM is required for efficient crRNA targeting (Mojica et al.,

2009).

Due to the site-specific targeting of CRISPR-Cas, the anti-viral system was adapted for site-directed mutagenesis in model organisms (Ma et al., 2014; Charpentier,

2015) Three main variations of the CRISPR-Cas system exist, differing in the Cas protein machinery and Cas proteins used (Fineran and Charpentier, 2012; Barrangou, 2013;

Barrangou and Marraffini, 2014). The CRISPR-Cas system II requires minimal Cas9 machinery, enabling its optimization for efficient site-directed mutagenesis and genome editing in zebrafish and other model organisms (Barrangou and Marraffini, 2014; Ma et al., 2014; Charpentier, 2015). Due to the minimal Cas9 machinery, system II requires a trans-activating crRNA (tracrRNA) along with crRNA for proper targeting (Barrangou and Marraffini, 2014; Ma et al., 2014). Synthetic single guide RNAs (sgRNAs) were engineered to encompass both crRNA and tracrRNAs facilitating Cas9 targeting

(Barrangou and Marraffini, 2014; Ma et al., 2014; Charpentier, 2015). Generating sgRNAs complementary to protospacers within a gene of interest will induce CRISPR-

Cas9 double-strand breaks within the gene. Inefficient repair via nonhomologous end joining induces nucleotide insertions or deletions likely disrupting the targeted gene. The

CRISPR-Cas system is an efficient and effective way to induce mutations in target genes

(Ma et al., 2014). Using this system, I induced nonsense mutations in 3 putative fusion regulators kirrel3l, iqsec1b, and ckip-1.

3.2 Methods

CRISPR Mutagenesis

CRISPR mutagenesis performed as previously described (Talbot and Amacher,

2014). The kirrel3l CRISPR target site used was 5’-GGTGGTGAACGAGACCTTCCTG

-3’. The iqsec1b CRISPR target site used was 5’- GGGTTGAGTCGACAGATGAT-3’.

The ckip-1 CRISPR target site used was 5’- GGCTGGATCCGCAAGTTCTG-3’.

Genotyping for alleles was performed by High Resolution Melt Analysis (HRMA) analysis of the lesion site with the primers flanking the CRISPR target site generating products between 100 and 300 bps.

Immunohistochemistry

Antibody staining was performed as previously described (Berberoglu et al.,

2017). The following primary antibodies were used at the noted dilutions: rabbit anti-

Rbfox1l (1:1000), mouse anti-A4.125 (DSHB 1:1000), and mouse anti-F310 (DSHB

1:1000)

RT-PCR

78 tails were immediately placed into Ringer’s solution before Trizol solubilization. Total

RNA (2.5 ug) was purified and reverse transcribed into cDNA using SuperScript VILO

MasterMix (ThermoFisher) following manufacturer’s instructions.

Needle-stick Injury

Needle-stick injury was perform as previously described (Berberoglu et al., 2017).

Confocal Imaging

All confocal imaging was performed using an Andor Revolution WD spinning disk confocal system supported by the Ohio State Neuroscience Center Core P30

NS045758 grant. Lasers used were 405 nm, 488 nm, 561 nm, and 640nm using 10x, 20x,

40x, and 60x objectives.

3.3 Generation and analysis of kirrel3l nonsense mutations

Motivation for generating a kirrel3l mutant

The vertebrate Kirre protein family consists of Ig domain-containing transmembrane proteins homologous to Drosophila Dumfounded (Duf), also known as Kin-of-IrreC

(Kirre), and Roughest (Rst). Duf and Rst are transmembrane cell adhesion molecules

(CAMs) necessary for founder cell (FC) interaction with fusion competent myoblasts

(FCMs) during Drosophila cell fusion (Ruiz-Gómez et al., 2000). In the absence of both

Duf and Rst, FCs cannot communicate with FCMs and fusion fails (Ruiz-Gómez et al.,

2000; Rochlin et al., 2010; Abmayr and Pavlath 2012; Kim et al., 2015). Most vertebrate

Kirre family proteins regulate kidney function, but in zebrafish, kirrel3l is expressed specifically in fast muscle precursors starting at 6 somites (Srinivas et al., 2007), implicating a potential function in muscle fusion. Fast muscle specific expression shortly 79 before fusion begins, supporting the idea that kirrel3l may function in myoblast fusion.

Additionally, strict amino acid conservation between the five Ig/Ig-like domains between

Duf, Rst, and Kirrel3l suggests conserved protein function. Previous work by others showed that blocking kirrel3l function by injecting an antisense morpholino oligonucleotide to block kirrel3l mRNA translation resulted in severe myoblast elongation defects and disrupted myoblast fusion (Srinivas et al., 2007). Slow muscle appeared normal and fast muscle differentiated normally, suggesting kirrel3l knockdown did not disrupt other aspects of the muscle differentiation program, except fast muscle- specific fusion (Svinivas et al, 2007). Due to conserved protein structure and the precedent from knockdown studies, I generated zebrafish kirrel3l mutants to generate a genetic model to study kirrel3l-mediated skeletal muscle fusion.

kirrel3l CRISPR sgRNA design and new mutant alleles

Both full protein deletions and C-terminal truncations completely disrupt Duf protein function (Ruiz-Gómez et al., 2000; Bulchand et al., 2010). To generate a likely null allele, I designed my CRISPR sgRNA to target kirrel3l in exon 2 of 15 (Fig 3.2).

Mutating kirrel3l in exon 2 was predicted to disrupt 4 of the 5 extracellular Ig-domains and also completely eliminate the required intracellular domains (Bulchand et al., 2010), likely generating a null protein. I recovered two nonsense mutations, named oz11 and oz12, which are predicted to truncate Kirrel3l at amino acid 70 of 754 (Fig 3.2). Due to the severity of the predicted Kirrel3l protein truncations, I expected that both mutations would result in non-functional Kirrel3l protein.

Characterization of kirrel3l mutant phenotype

To assay skeletal muscle fusion defects in my recovered kirrel3l mutants, I generated homozygous mutants by intercrossing F2 heterozygous adults and characterized skeletal muscle morphology. By immunocytochemistry, I observed multinucleate fast muscle fibers in both kirrel3loz11 and kirrel3loz12 homozygous mutants at 24 hpf (Fig 3.2), implicating that zebrafish Kirrel3l is not required for embryonic fusion. However, because post-embryonic fusion is involved in muscle growth and regeneration, I also raised kirrel3l mutants to adulthood. kirrel3l mutants are indistinguishable from wild- type siblings, suggesting that muscle growth occurs normally. To determine if kirrel3l mutant adults undergo normal muscle repair, I performed needle-stick injury and assayed expression of regeneration markers Pax7 and Rbfoxl. Confocal imaging revealed kirrel3l mutants mount a normal repair response similar to wild-type siblings, suggesting muscle repair occurs normally in kirrel3l mutants (Fig 3.2). To date, I have not been able to discover any fusion defects in kirrel3l mutants.

3.4 Generation and analysis of iqsec1b and ckip-1 nonsense mutations

Motivation for generating iqsec1b and ckip-1 mutants

In addition to transmembrane proteins that regulate skeletal muscle fusion, downstream fusion regulators such as the intracellular proteins Loner (Pajcini et al.,

2008), CKIP-1 (Baas et al., 2012), Dock, and Crk/Crkl (Moore et al., 2007) have also been implicated in vertebrate myoblast fusion. Because these studies were performed either by antisense morpholino knockdown or in cell culture, I wanted to validate gene

81 function by generating zebrafish CRISPR mutants. At the time I began the project, mutagenesis of Dock and Crk/Crkl was already underway in our lab, so I decided to focus on the zebrafish homologues of Drosophila Loner and CKIP-1.

Drosophila loner was discovered in a genetic screen for genes regulating muscle development (Chen et al., 2003). Specifically, Loner is an ARF6 guanine nucleotide exchange factor (GEF) necessary for myoblast fusion (Chen et al., 2003, Donaldson

2003). It requires a functional Sec7 domain, GEF activity, and pleckstrin homology (PH) domain, and plasma membrane localization, for fusogenic activity (Chen et al., 2003,

Donaldson 2003). Loner co-localizes with Duf/Rst on the plasma membrane (Chen et al.,

2003; Bulchand et al., 2010) and physically interacts with the Duf intracellular domain

(Bulchand et al., 2010). Currently two models exist to explain Loner function during

Drosophila myoblast fusion (Chen et al., 2003; Dottermusch-Heidel et al., 2012). One model proposes ARF6 recruitment by Loner is necessary for cytoskeletal rearrangements required for cell alignment (Chen at al., 2003), while the other model suggests Loner functions to remove N-cadherin from the cell-cell contact site to facilitate myoblast fusion (Dottermusch-Heidel et al., 2012). In mammals, the Loner homologue Brag2 is also necessary for ARF6 activation and efficient myoblast fusion of cultured C2C12 cells

(Pajcini et al., 2008), suggesting a conserved role for ARF6 activation in mammalian myoblast fusion. To validate the requirement for Loner/Brag2 in a whole animal model, I generated an iqsec1b mutant allele, the zebrafish ortholog of Loner/Brag2.

Actin cytoskeleton regulation is necessary for both invertebrate and vertebrate myoblast fusion (reviewed in Rochlin et al., 2010). Specifically in zebrafish, live imaging

82 analysis of muscle morphogenesis associates major cell morphogenetic movements with myoblast fusion (Snow et al., 2008). Actin nucleation and termination via capping proteins is tightly coupled to regulate actin-base cell motility and morphology (Carlier and Pantaloni 1997; Carlier et al., 2015). CKIP-1 promotes actin nucleation and influences cell morphology by interacting with actin capping proteins, inhibiting their ability to terminate actin nucleation, (Canton et al., 2005; Canton et al., 2006) and recruiting the actin nucleation factor actin-related protein 2/3 (Arp2/3) (Baas et al., 2012).

The CKIP-1 PH domain is necessary for plasma membrane localization (Olsten et al.,

2004; Safi et al., 2004; Canton et al., 2006) and functions in C2C12 muscle differentiation (Safi et al., 2004). C2C12 and zebrafish knockdown and overexpression studies suggest CKIP-1 is necessary for myoblast fusion in both mammalian cell culture and zebrafish (Baas et al., 2012). Due to the association between dynamic actin cytoskeleton movements with zebrafish myoblast fusion (Snow et al., 2008) and the implication of CKIP-1 in vertebrate myoblast fusion (Baas et al., 2012), I designed

CRISPR sgRNAs to mutate zebrafish CKIP-1.

iqsec1b and ckip-1CRISPR sgRNA design, new mutant alleles, and phenotype characterization

As in Loner, the zebrafish homologue Iqsec1b contains a Sec7 and PH domain.

Because both the Sec7 and PH domain are necessary for Loner function, I designed a

CRISPR sgRNA predicted to induce mutations disrupting both domains (Fig 3.3).

Targeting the Sec7 domain, I recovered a 7 bp deletion that generated a premature stop

83 codon at amino acid 559 of 1122 (Fig 3.3). This deletion allele truncates the Iqsec1b protein early in the Sec7 domain and completely eliminates the PH domain, likely generating a nonfunctional protein hypothesized to disrupt myoblast fusion. Similarly, when designing the ckip-1 sgRNA, I targeted the necessary PH domain (Fig 3.3). I found a 5 bp deletion creating an in-frame stop codon at amino acid 60 of 403. The premature stop codon truncates CKIP-1 early in the PH domain implying protein function is lost.

To assay both iqsec1b and ckip-1 fusion phenotypes I analyzed myoblast fusion at 24 hpf in homozygous mutants. Using muscle specific markers Rbfox1l, a muscle nuclear marker, F310, fast muscle marker, and A4.1025,a myosin heavy-chain marker, I observed normal fast muscle fusion in both iqsec1b and ckip-1 mutants (Fig 3.3). These data suggest that both Iqsec1b and CKIP-1 are either compensated for or not necessary for zebrafish fast muscle fusion at 24 hpf. Alternatively, both ckip-1 (Baas et al., 2012) and brag2 (Fig 3.3) RNA is maternally provided potentially generating enough wild-type protein necessary for fast muscle fusion. Thus, the maternally provided RNA could be masking any potential embryonic fusion phenotype. To determine whether iqsec1b or ckip-1 are required for post-embryonic muscle growth, I raised homozygous mutants to adulthood. Both mutants were indistinguishable from wild-type siblings suggesting iqsec1b and ckip-1 are not required for zebrafish muscle growth.

3.5 Discussion

Morpholino-induced artifacts or genetic compensation may cause discrepancies between mutant and morphant phenotypes

Morpholinos (MOs) are synthetic antisense oligonucleotides designed to inhibit gene expression by blocking RNA translation or splicing (Summerton and Weller 1997;

Kok et al., 2015; Stainier et al., 2017). Before efficient gene-targeted mutagenesis strategies were developed for zebrafish, MOs were used to inhibit gene expression in a gene-specific manner and successfully recapitulated characterized mutant phenotypes

(Nasevicius and Ekker 2000; Draper et al., 2001). However, with recent advances in gene-targeted mutagenesis, discrepancies between mutant and MO-knockdown

(morphant) phenotypes have surfaced (Law and Sargent 2014; Kok et al., 2015;

Novodvorsky et al., 2015; Stainier et al., 2017). Because I observed similar discrepancies between morphant and mutant phenotypes for kirrel3l, iqsec1b, and ckip-1, potential

MO-related artifacts could have generated a false phenotype (Kok et al., 2015;

Novodvorsky et al., 2015). MO off-target affects or tissue-specific MO toxicity, such as in skeletal muscle, may have induced the severe fusion defects seen in kirrel3l, iqsec1b, and ckip-1 morphants that were not observed in mutants (Stainier et al., 2017).

Alternatively, genetic compensation within the single mutants may mask the morphant phenotype (Rossi et al., 2015; Stainier et al., 2017). Morpholinos block both maternal and zygotic RNA expression, preventing all gene expression, while mutants may have functional maternal RNA sufficient to promote myoblast fusion. Both iqsec1b and ckip-1 are maternally provided suggesting maternal RNA compensation may occur in zygotic mutants. Generating maternal-zygotic mutants for all three genes will determine if maternal compensation occurs within the zygotic mutants. Additionally, embryos may compensate for a defective gene by upregulating an alternative pathway (Rossi et al.,

2015). In zebrafish, all three gene function alongside other molecules performing similar functions, such as Jam2a, Jam3b, and Myomaker for Kirrel3l and Dock1/5 and Crk/Crkl for Iqsec1b and CKIP-1. Performing gene expression analyses on each mutant will help clarify whether or not alternative pathways are being upregulated to compensate for the mutated gene.

Genomic compensation may prevent CRISPR-induced mutant phenotypes

Alternative to MO-related artifacts or genetic compensation, a lack in mutant phenotype may occur due to mRNA processing. A recent study discovered 3 mRNA splicing strategies genes use to bypass induced genetic mutations, referred to as genomic compensation (Anderson et al., 2017). The three genomic compensation strategies were:

1) skipping exons with mutations at or near essential splice sites, 2) skipping exons with premature terminating codons (PTCs), and 3) utilizing cryptic splice sites induced by a mutation (Anderson et al., 2017). All strategies have the potential to produce functional proteins if the transcript remains in-frame and no essential domains are affected. Thus, critical mRNA analysis must be done to ensure induced mutations do not promote genomic compensation. If an induce mutation generates a PTC within 50 bp of an exon junction, generates a PTC within an exon that can be skipped without inducing a frameshift, or generates a PTC in a non-essential exon, genomic compensation could occur generating a functional protein (Anderson et al., 2017). mRNA analysis performed on induced CRISPR mutations eliminated genomic compensation for iqsec1b and ckip-1, but is left possible for kirrel3l. The kirrel3l mutation is induced near an essential splice

86 site and if exons 1-2 are skipped, the transcript remains in frame with 4 of 5 functional Ig domains. However, to fully verify the status of mutant mRNA, sequencing of the mutant mRNA should be performed. The presence of mutant mRNA could possibly account for a lack in mutant phenotype.

Figure 3.1 CRISPR locus and necessary targeting components

CONTINUED

Figure 3.1 CONTINUED (A) An illustration of a basic CRISPR loci containing the 3 essential segments: 1) the CRISPR repeats (black polygons) 2) CRISPR spacers (colored rounded rectangles) and 3) cas associated genes (green rectangle). The CRISPR repeat and spacer region is typically preceded by a leader sequence (yellow rounded rectangle).

(B) Transcription and RNA processing generates a specific crRNA (blue spacer) that will guide Cas protein to the complementary protospacer. (C) The hairpin generated from the

CRISPR repeat interacts with Cas protein while the spacer sequence guides Cas to the target site for cleavage. (Adapted from Barrangou and Marraffini 2014)

Figure 3.2 Identified kirrel3l mutants lack noticeable fusion defects

CONTINUED

Figure 3.2 CONTINUED (A) The DNA lesions for two isolated kirrel3l mutations

(CRISPR target sequence in bold and affected bases in red). kirreloz11 consists of a 4 bp deletion (red dashes) with a 2 bp change (red italics) while kirrel3loz12 consists of only the

4 bp deletion (red dashes). (B) Both frameshifting mutations truncate the protein at amino acid 70 of 754. The truncated protein is predicted to completely lack 4 of 5 extracellular

Ig domain and the entire intracellular domain. (C-D) 24 hpf antibody labeling of wild- type (C) and kirrel3loz12 mutants (D). Cell membranes are labeled with beta-catenin

(green), fast muscle fibers are marked by F310 (blue), and all nuclei are labeled with TO-

PRO-3 (red), revealing multinucleate fast muscle fibers at 24 hpf. (E-F) Transverse sections of adult wild-type (E) and kirrel3loz12 mutant (F) muscle following needle-stick injury. Muscle fibers are labeled with a myosin heavy-chain marker A4.1025 (red) and differentiated myonuclei are labeled with Rbfox1l (green). Rbfox1l cytoplasmic expression is associated with regenerating myofibers (Berberoglu et al., 2017), suggesting both wild-type and kirrel3loz12 mutants are actively regenerating post-needle stick injury.

Figure 3.3 CRISPR lesions and phenotype associated with iqsec1b and ckip-1

CONTINUED

Figure 3.3 CONTINUED Figure 3.2 CONTINUED (A) The DNA lesions for iqsec1b and ckip-1 CRISPR mutations (CRISPR target sequence in bold and affected bases in red). The iqsec1bmut lesion consists of a 7 bp deletion (red dashes) while ckip-1oz13 consists of a 5 bp deletion (red dashes). (B) Both frameshifting mutations truncate the protein in essential protein domains. (C-F) 24 hpf antibody labeling of wild-type (C, E) and iqsec1bmut mutants (D) or ckip-1oz13 mutants (F). Myosin heavy chain is labeled with

A4.1025 (red), fast muscle fibers are marked by F310 (blue), and all nuclei are labeled with Rbfox1l (green), revealing multinucleate fast muscle fibers at 24 hpf for both mutants. (G) RT-PCR expression for iqsec1b at various developmental stages. iqsec1b expression during the 4-8 cell stage indicates maternal deposition of iqsec1b RNA. Cell division cycle 5-like (cdc5l) was used as a positive control for the RT-PCR.

Chapter 4. Conclusions & Future Directions

I have shown zebrafish myoblast fusion is coincident with fast muscle elongation and correlated with slow muscle migration. This study also suggests that myoblast fusion requires the clearance of interacting membranes for successful myoblast merging.

Additionally, I revealed post-embryonic slow muscle fusion utilizes a previously undescribed myomaker-mediated fusion mechanism compared to embryos. I proposed muscle hypertrophic growth requires myomaker-mediate satellite cell fusion and found a

50% reduction in myonuclei is sufficient to maintain muscle growth and strength. I revealed that fusion-deficient muscle resembles a muscle diseased state with decreased

Laminin protein, increased cell proliferation, and increased satellite-like cell activation.

Lastly, I speculated why three putative zebrafish fusion mutants kirrel3l, iqsec1b, and ckip-1 display no obvious fusion defects. This study proposes zebrafish mononucleate slow fibers may provide an environment for satellite-like cell maturation, enables future investigation into embryonic versus adult fusion mechanisms, and promotes investigation of muscle regeneration in fusion-deficient skeletal muscle.

Post-embryonic slow muscle fusion suggests different fusion mechanisms regulate embryonic and post-embryonic muscle fusion

One of the main findings of this study was that slow muscle multinucleation occurs post-embryonically in zebrafish and is likely due to asymmetric fusion, a mechanism that differs from embryonic fusion events. This is interesting because it suggests that embryonic and adult fusion mechanisms are different and fusion regulation 94 differs at different developmental stages. I proposed mononucleate slow muscle cells may function to protect future SLCs from fusogenic fast muscle cells during myogenesis.

Thus, after SLCs are established post-embryonically, slow muscle fusion is permitted.

Testing this hypothesis requires Pax7-positive cell quantification in the presence of fusogenic slow muscle cells, which can be achieved with ectopic myomaker expression

(Zhang and Roy 2017). If mononucleate slow muscle cells serve as a protective layer, then inducing fusogenic activity in slow muscle would significantly reduce the number of

Pax7-positive cells.

This study revealed slow muscle fusion occurs post-embryonically, but it is currently unknown why slow muscle cells become multinucleate. During myogenesis, fast muscle specific genes are specifically inhibited in slow muscle preventing the expression of fusion-promoting genes (Baxendale et al., 2004; von Hofsten et al., 2008). myomaker expression analysis during slow muscle fusion suggests slow muscle fusion does not require Myomaker cell-autonomously, but Myomaker is required for slow muscle fusion. These data suggest Myomaker is only required on one cell for post- embryonic fusion. However, the regulatory switch from both cells requiring Myomaker, embryonic fusion, to one cell requiring Myomaker, post-embryonic fusion, is currently unknown. It would be interesting to analyze slow muscle transcription from embryonic to the fusogenic juvenile stage to determine if change in slow muscle transcription induces fusogenic promoting factors. If no transcriptional change occurs in slow muscle cells, then external factors, such as those potentially associated with a SLC niche, or rapid muscle growth may prime slow muscle cells for post-embryonic fusion. Alternatively,

95 jam2a mutants could also be used to identify regulatory changes between embryonic and post-embryonic fusion events, because both mononucleate fast and slow jam2a mutant fibers become multinucleate post-embryonically.

myomaker mutants provide a model to determine if hypertrophic growth can be induced without muscle fusion

This study found that zebrafish skeletal muscle requires Myomaker-mediated muscle fusion to undergo hypertrophic growth. I found that muscle strength and size correlated with levels of hypertrophic growth. Thus, myomaker mutants serve as a model to investigate pathways regulating fusion-independent hypertrophic growth. It would be interesting to determine if sufficient fusion-independent hypertrophy could be induce in myomaker mutant fibers to maintain wild-type size and strength. Uncovering the mechanisms required to increase muscle size and strength without muscle fusion may provide new treatments for muscle diseases that lose the ability to undergo myoblast fusion, such as in cancer cachexia (He et al., 2013).

Investigating the myomaker mutant adult muscle environment will determine whether mutants display a dystrophic or cachectic muscle phenotype

In this study I present evidence for muscle damage and increased cell proliferation which are characteristic in both muscular dystrophy and muscle wasting models (Tisdale

2002; Fanzani et al., 2013; He et al., 2013; Argilés et al., 2014; Madaro and Bouché

2014). A key difference between muscular dystrophy and cachectic muscle pathology is

96 the presence or absence of invading immune cells (He et al., 2013). Immune cells, such as neutrophils, macrophages, and lymphocytes, readily invade dystrophic muscle and contribute to the degenerative pathology (Choi et al., 2009; Pillon et al., 2013; Madaro and Bouché 2014), unlike cachectic muscle (Tisdale 2002; He et al., 2013). Cachectic muscle is defined by increased catabolic activity and decreased anabolic activity leading to muscle wasting (Tisdale 2002; He et al., 2013; Argilés et al., 2014). Key molecules involved in cachectic muscle are the cytokines tumour necrosis factor-α (TNF-α) and interleukin-1 (IL-1) (Argilés et al., 2014). TNF- α and IL-1 promote muscle wasting by activating the nuclear factor-κβ (NF- κβ) and p38 MAPK pathways which upregulate muscle RING finger-containing protein-1 (MURF-1) and muscle atrophy F-box protein

(MAFBX) (He et al., 2013; Argilés et al., 2014). MURF-1 and MAFBX are E3 ligases that inhibit protein synthesis and promote muscle protein breakdown (Clark et al., 2007;

Cohen et al., 2009; Glass 2010; Argilés et al., 2014). It would be interesting to investigate whether immune cells infiltrate myomaker mutant adult skeletal muscle, or whether the molecules associate which muscle wasting are upregulated. Further investigation into both dystrophic and cachectic mechanisms will help determine whether myomaker mutants display a dystrophic or muscle wasting phenotype. Myomaker mutants could then serve as a model for how fusion deficiency promotes the respective phenotype.

In addition to determining what type of muscle disease myomaker mutants resemble, it would be interesting to investigate how fusion deficiency alters disease progression. Various dystrophy mutants have been modeled and recapitulated in zebrafish (Berger and Currie 2012; Li et al., 2017). Generating double mutants with

97 myomaker and various dystrophic mutant alleles, such as sapje (Bassett et al., 2003), will give insight into how fusion deficiency either promotes or inhibits disease progression.

Similarly, inducing rhabdomyosarcomas in myomaker mutants via rag2:kRASG12D injection (Langenau et al., 2007), will give insight into how fusion deficiency alters progression of rhabdomyosarcoma tumors in zebrafish.

Zebrafish myoblast fusion may resemble Drosophila membrane fusion dynamics

This study found plasma membrane-labeled puncta, which may represent vesicle- like structures, disperse during zebrafish myoblast fusion. In Drosophila, endocytic vesicles function to maintain founder cell (FC) identity following asymmetric myoblast fusion. The vesicles remove fusion competent myoblast (FCM)-specific membrane components, maintaining FC membrane identity. Zebrafish myoblast fusion does not involve asymmetric fusion, but endocytic vesicles may provide a buffer mechanisms to prevent ectopic fusion between fast and slow muscle cells. Myomaker can induce fusion between two myoblasts when only expressed in one cell, albeit inefficiently (Millay et al.,

2012), therefore removing Myomaker from fast muscle membranes may prevent ectopic fusion and/or regulate the number of fusion events. Alternatively, post-embryonic zebrafish fusion does appear to be asymmetric. It would be interesting to investigate endocytic vesicles during post-embryonic fusion to determine if cell identity maintenance is required following post-embryonic fusion. Maintaining cell identities following post- embryonic fusion events could regulate the number of fusion events or prevent unnecessary hyperfusion in adult muscle. Future studies characterizing the type of vesicles formed and their fate may shed light on their function during fusion. The 98 vesicles may function simply in membrane dispersal and recycling to allow physical merging of the cells, or may have a deeper role maintaining cell identity by recycling fusion-promoting transmembrane proteins

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