Morphogenesis and specification of the muscle lineage

during Xenopus laevis embryo development

by

Armbien F. Sabillo

A dissertation submitted in partial satisfaction

of the requirements for the degree of

Doctor of Philosophy

in

Molecular & Cell Biology

in the

Graduate Division

of the

University of California, Berkeley

Committee in charge:

Professor Richard M. Harland, Chair

Professor John Gerhart

Professor Craig T. Miller

Professor Leslea Hlusko

Summer 2018

1

Abstract

Morphogenesis and specification of the muscle lineage

during Xenopus laevis embryo development

by

Armbien F. Sabillo

Doctor of Philosophy in Molecular & Cell Biology

University of California, Berkeley

Professor Richard M. Harland, Chair

Development consists of complex morphogenetic movements that shape individual tissues

as well as the embryo. Tissues must interact both physically and through signaling molecules

to coordinate the formation of various organs and the entire body plan. My thesis work

sought to characterize tissue interactions that govern muscle formation during vertebrate

embryo development using the African clawed frog Xenopus laevis as a model system.

Chapter 1 of this dissertation provides a general introduction to the morphogenetic events

and molecular regulation leading to muscle formation. Chapter 2 presents two previously-

undiscovered morphogenetic phenomena involving the muscle tissue as well as individual

muscle fibers. My experiments confirmed the function and mechanisms of muscle tissue

unfolding as well as the cell rearrangements that ultimately place muscle fibers into

organized arrays for proper functioning. Chapter 3 summarizes my work to further analyze

a previously-unreported reciprocal relationship between the neural tissue and the

prospective muscle tissue. My work shows that the neural plate influences the size of the

underlying muscle. Enlarging the neural plate results in a corresponding increase in the size

of the muscle tissue. Here, I dissect the source and fate of ectopic muscle tissue and

determine the effect of tissue size on muscle morphogenesis. As a whole, this work

summarizes my research to characterize the molecular and morphological events necessary

for proper muscle formation during embryo development.

i

Dedication

This dissertation is dedicated to the embryos that gave their lives for the sake of knowledge.

ii

Acknowledgments

I thank members of the Harland Lab and my thesis committee for their helpful insights.

Special thanks go to Edivinia Pangilinan for years of support and for ensuring that I never

felt too homesick for the Philippines, and to Marie Sabillo for assistance in image processing.

Thank you also to James Evans for his expertise and help with animal husbandry and to

Carmen Domingo for sincere encouragement and excellent mentorship. Two remarkable

students contributed to this work under my mentorship: Heng-Yi Liu and Ola Egu. Thank you

for your hard work and enthusiasm; every day you reminded me of the pure wonder of

scientific research.

iii

Table of Contents

Abstract…………………………………………………………………………..………………………….…………………1

Dedication………………………………………………………………………………………………………………….….I

Acknowledgments………………………………………………………………...…………………………………...…II

Table of Contents……………………………………………………………….………………………………………..III

List of Figures……………………………………………………………………………………………………………...IV

Chapter 1: Introduction

1.1 Cell movements underlying somite formation……………………………………………………………..1

1.2 Role of the extracellular matrix on somite formation…………………….……………………………...3

1.3 Early embryonic position determines presomitic cell behaviors………………………………….,,3

1.4 Formation of the dermomyotome………………………………………………………………..……………...4

1.5 Signals that influence somite patterning……………………………………………………………………...4

1.6 Myogenic waves and compartmentalization of the somite………………….………………………...5

1.7 Morphogens that pattern the somite…………………………………………………………………………...5

1.8 Transcriptional regulation of X. laevis myogenesis………………………………..……………………...8

1.9 Muscle Satellite Cells and Regeneration…………………………………………………………….………12

Chapter 2: Tissue morphogenesis and cell rearrangements leading to somite

formation

2.1 Abstract………………………………………………………………………………………………….………………14

2.2 Introduction……………………………………………………………………………………………………………14

2.3 Results……………………………………………………………………………………………………………………15

Marginal zone cells involute and form the presomitic mesoderm

Myotome forms in specific domains within the somite based on their initial position

Pre-somitic mesoderm tissue unfolds dorsally and ventrally as it forms somites

iv

Tissue unfolding positions PSM cells to the dorsal and ventral regions of the somite

Cells within the PSM undergo dramatic rearrangements after tissue unfolding

PSM cell rearrangements are accompanied by transient ECM anchor points

PSM tissue unfolding and cell rearrangements occur without the influence of adjacent

tissues

Autonomous PSM tissue unfolding and cell rearrangements are not mediated by the

Wnt/PCP pathway

2.4 Discussion………………………………………………………………………………………………………………21

PSM unfolding drives positioning of myotome in the dorsal-ventral axis

Cell rearrangements after PSM unfolding are accompanied by extracellular matrix

protein anchor points

PSM unfolding and cell rearrangements occur autonomously and do not require the

Wnt/PCP pathway

2.5 Methods………………………………………………………………………………….………………………………23

Chapter 3: The role of the neural plate on muscle fate specification

3.1 Abstract…………………………………………………………………………………………….……………………26

3.2 Introduction……………………………………………………………………………………………………………26

3.3 Results……………………………………………………………………………………………………………………27

Expansion of the neural plate results in a corresponding increase in presomitic

mesoderm size

Reducing neural plate size results in reduction of presomitic mesoderm size

Effect of neural plate size on presomitic mesoderm specification is non-cell

autonomous

Ectopic presomitic mesoderm is transient and results in disorganized somites

Ectopic presomitic mesoderm does not undergo apoptosis

Ectopic presomitic mesoderm is specified at the expense of adjacent tissues and is

later re-specified to the source tissue

Ectopic neural tissue is transient and is dorsal and posterior in identity

v

Expression of early and late myogenic regulatory factors is perturbed

Ectopic presomitic mesoderm results in abnormal somite morphogenesis

Ectopic presomitic mesoderm results in disrupted myotome alignment

Disrupted somites have abnormal extracellular matrix protein deposition

Sonic Hedgehog does not rescue MRF expression or somite morphogenesis

3.4 Discussion………………………………………………………………………………………………………………33

Neural plate size and presomitic mesoderm size are correlated

Foxd1 activity in the neural plate and paraxial mesoderm

Ectopic presomitic mesoderm tissue disrupts somite morphogenesis

3.5 Methods………………………………………………………………………………………….………………………36

Future Perspectives……………………………………………………………………………………………………38

Figures………………………………………………………………………………………………………...………………40

References……………………………………………………………………………………………………..……………78

vi

List of Figures

Chapter 1

Fig 1: Schematic of myotome rotation and extracellular matrix deposition

Fig 2: Fate map of presomitic mesoderm morphogenesis

Fig 3: Diagram of signaling between adjacent tissues during somitogenesis

Fig 4: Visual summary of myogenic regulatory factor expression

Chapter 2

Fig 1: Pre-somitic cells from different regions of the gastrula take distinct routes to reach the

dorsal part of the embryo.

Fig 2: Pre-somitic cells form myotome in specific domains along the anterior-posterior and

dorsal-ventral axes.

Fig 3: The pre-somitic mesoderm unfolds dorsally and ventrally as it matures into somites.

Fig 4: Tissue unfolding positions cells from the lateral edge to the dorsal and ventral

domains of the somite.

Fig 5: Cells undergo dramatic rearrangements after PSM unfolding.

Fig 6: Cell rearrangements following somite unfolding are accompanied by transient

depositions of ECM proteins in key locations.

Fig 7: PSM unfolding and cell rearrangements occur autonomously.

Fig 8: Somite unfolding and cell rearrangements are not mediated by non-canonical

Wnt/PCP pathway

Fig 9: Model for myotome positioning through autonomous somite unfolding and cell

rearrangements.

Supp Fig 1: Marginal zone cells in the LAT region undergo behaviors similar to cells in both

the ULL and LL regions

Supp Fig 2: β-dystroglycan also contributes to extracellular matrix hinges.

Supp Fig 3: Canonical Wnt and noncanonical Wnt/PCP mutants exhibit secondary defects,

but somites still unfold

vii

Chapter 3

Fig 1: foxD1 mRNA expands the neural plate and the paraxial mesoderm in a dose-

dependent manner.

Fig 2: Sox2 MO reduces the size of the neural plate and paraxial mesoderm.

Fig 3: Foxd1 mRNA-injected animal caps induce ectopic paraxial mesoderm non-cell

autonomously.

Fig 4: Ectopic paraxial mesoderm disappears after late tailbud stage.

Fig 5: Ectopic tissue does not undergo apoptosis

Fig 6: Ectopic paraxial mesoderm is specified at the expense of intermediate and lateral

plate mesoderm, and is re-specified to the source tissue by the late tailbud stage.

Fig 7: Ectopic neural tissue is dorsal and posterior in identity, but disappears by the late

tailbud stage.

Fig 8: Late MRF expression is abnormal.

Fig 9: Somite morphogenesis is disrupted.

Fig 10: Muscle fibers in somites are disorganized.

Fig 11: Extracellular matrix proteins are disrupted in foxd1-injected embryos.

Fig 12: Sonic hedgehog does not rescue somite morphology in foxd1-injected embryos.

Fig 13: Model for foxd1-mediated interaction of the neural plate and paraxial mesoderm.

Supp Fig 1: Somite morphogenesis is characterized by distinct cell behaviors

Supp Fig 2: Somite morphogenesis depends on proper ECM deposition in the intersomitic

boundaries

1

Chapter 1: Introduction

The modulation of segment shape, size, and number during evolution contributes to

the diversity of organisms. In vertebrates, segmentation involves the partitioning of the

presomitic mesoderm (PSM) into numerous metameric segments called somites, which will

subsequently form the axial skeleton, dermis, and skeletal muscle. Understanding the

molecular and cellular underpinnings of this process is of major importance as the somites

establish the segmented body plan of all vertebrates. Moreover, somites provide important

guidance cues during neural crest migration and nerve projections from the central nervous

system.

Given that Xenopus laevis undergoes indirect development (i.e. embryo forms a larva

before becoming an adult), the embryonic musculature varies in significant ways in

comparison to most vertebrate models, which are direct developers [1]. In particular, the

traditional somite of direct-developing vertebrates gives rise to the dermomyotome,

myotome, and sclerotome. However, the X. laevis somite is primarily comprised of myotome

fibers, which will go on to form the temporary muscle system of the tadpole. Given that the

tadpole’s musculature will be remodeled during metamorphosis, X. laevis provides a unique

opportunity to dissect the mechanisms behind embryonic versus adult myogenesis. This

mode of development allows researchers to determine which aspects of myogenesis are

evolutionarily conserved across vertebrates.

In this chapter we discuss key insights gained by investigating myogenesis during

tadpole formation in X. laevis. Given the excellent reviews that focus on mesoderm induction

in X. laevis [2,3], we emphasize patterning events that occur after the mesoderm has been

established. First, we describe the cell movements and morphogenetic events that lead to the

formation of muscle fibers in discrete locations in the tadpole. This is followed by a

discussion of the molecular signals necessary for the establishment and differentiation of the

muscle lineages. Finally, we examine the formation of muscle satellite cells and regeneration

in the tadpole. Together, this chapter aims to highlight the unique contributions of work in X.

laevis that has led to a better understanding of the molecular and cellular mechanisms

underlying embryonic muscle formation.

2. Cell movements underlying somite formation

Somite formation proceeds in an anterior to posterior direction during development,

with bilaterally symmetric somite pairs forming at constant intervals. This periodicity has

led to a model in which an intrinsic oscillator [4,5] functions to instruct presomitic cells to

form segments. The most prevalent of these models is the “clock and wavefront”, which is

based on observations of somitogenesis in X. laevis [4]. In this model, groups of presomitic

cells oscillate (the clock) between two states, “permissive” or “not permissive” for

2

segmentation, while the "wavefront" represents an anterior to posterior progression of

embryonic development. Thus, when a group of cells are in the right phase of the oscillator

cycle and then encounter the wavefront, they undergo changes in cell behavior that lead to

the formation of a somite. For details associated with the molecular signals regulating this

process please refer to a recent review [6]. In brief, studies have shown that Wnt, Fgf, and

Notch pathways regulate the clock. These signals are expressed in posterior to anterior

waves with the timing of their expression correlating with somite formation. Interestingly,

the rhythmic activation of these signaling pathways appears to be a conserved property

among vertebrates. However, in comparing the cyclic gene networks between the mouse,

chick and zebrafish, it appears that the specific cyclic genes within each of these pathways

vary considerably between these species [7]. The molecular signals that underlie the

wavefront appear to consist of antagonistic gradients of retinoic acid at the anterior end of

the embryo and Fgf and Wnt at the posterior end. The intersection of the anterior and

posterior gradients define the determination front where the next prospective somite will

form [8]. In X. laevis, retinoic acid directly activates the expression of thylacine1 (thy1), a

mesp2 homolog. Thy1 is expressed in dynamic stripes corresponding to prospective

segmental boundaries and acts as an indirect inhibitor of the Fgf signaling pathway [9]. In X.

laevis the coordination of these cycling genes along with the wavefront signals will lead to

the formation of bilaterally symmetric somite pairs every 50 minutes [8].

In X. laevis, a unique series of cell behaviors leads to the formation of somites

primarily comprised of aligned myotome fibers [8,9]. A particularly interesting aspect of

somitogenesis in X. laevis is the 90o rotation that occurs soon after a segment bisects a group

of cells from the PSM. Interestingly, X. laevis is not the only species to undergo somite cell

rotation. It has also been reported in two additional pipid frog species, Xenopus tropicalis and

Hymenochirus boettgeri [12]. Since somite cell rotation has not been observed in other

anuran species, it is thought that this process may represent a shared derived character

(synapomorphy) for Pipidae. Interestingly, Hollway and colleagues [13] reported whole-

somite rotation in zebrafish. They showed that the signaling pathway, stromal derived factor

1α (sdf-1α) along with its receptor cxcr4, were required for somite rotation. This signaling

pathway was also shown to play a role in regulating somite cell rotation in X. laevis. However,

the morphogenetic processes underlying somite rotation are quite distinct between X. laevis

and zebrafish [14].

In 2000, Keller provided a comprehensive review of somite morphogenesis among

several different amphibian species [15]. Using observations from time-lapse images of X.

laevis explants [16] and scanning electron micrographs [11], Keller described the 90o

rotation that cells within the somite undergo to form elongated myotome fibers. To extend

these observations, Domingo and colleagues (2006) used a membrane targeted GFP to

visualize cell shape changes associated with somitogenesis in intact embryos at various

different stages of development [17]. This study led to a four-step model in which the first

3

step consists of the progressive elongation along the mediolateral axis of PSM cells as they

reach the determination front (Fig. 1, Step 1). This is followed by the formation of the

intersomitic boundary, which begins at the lateral edge of the anterior PSM and proceeds

medially towards the notochord [16]. The progressive formation of the intersomitic

boundary corresponds with an increase in the protrusive behavior of cells within the

forming somite (Fig. 1, Step 2). Next, these cells will largely maintain their elongated cell

shape as they rotate 90o (Fig. 1, Step 3) to make stable attachments to the intersomitic

boundaries flanking the somite. In this fashion the individual cells within the somite form a

parallel array of elongated myotome fibers (Fig. 1, Step 4). These coordinated steps are

repeated every 50 minutes until the tadpole has formed 45 somite pairs [18]. Although the

cell behaviors that underlie this seemingly simple 90˚ rotation are well described, the

molecular signals that coordinate this process are quite intricate and not fully understood.

1.2 Role of the extracellular matrix on somite formation

The deposition and assembly of the extracellular matrix (ECM) in the intersomitic

boundaries play a key role in X. laevis somite formation and myotome alignment. Fibronectin

is one of the first ECM proteins detected in the intersomitic boundaries [19]. The proper

scaffolding of fibronectin within the intersomitic boundary appears to be driven by integrin

α5 expression during somite rotation [20]. This is followed by the deposition of laminin,

whose assembly requires -dystroglycan expression [21] (Fig. 1). Knockdown of either

integrin α5 or -dystroglycan disrupts intersomitic boundary formation and somite rotation

[20]. Thus, the proper formation of intersomitic boundaries is necessary for somite cell

realignment. Recently, paraxis, a member of the bHLH type family of transcription factors,

was shown to play a role in regulating somite cell adhesion as knockdown of paraxis led to a

decrease in E-cadherin and EP-cadherin expression [22], resulting in the inability of somite

cells to adhere to the intersomitic boundaries [23]. Although it is unclear how the various

adhesion complexes are integrated, it is clear that formation and scaffolding of the ECM that

constitutes the intersomitic boundary is tightly coordinated with the formation of specific

adhesion complexes that drive somite cells to realign along the anteroposterior axis and

adopt an elongated cell shape.

1.3 Early embryonic position determines presomitic cell behaviors

In the X. laevis gastrula, cells lateral and ventral to the organizer will give rise to the

paraxial mesoderm [24–26]. Interestingly, depending on their position in the gastrula, the

prospective paraxial mesoderm cells will undergo different cell behaviors and trajectories to

give rise to myotome fibers in specific dorsal-ventral and anterior-posterior locations in the

tadpole [27]. For example, cells adjacent to the organizer undergo convergence and

4

extension movements, much like the prospective notochord cells, to form myotome fibers

along the entire anterior-posterior axis of the tadpole. These cells will eventually lie next to

the notochord and span the medial to lateral extent of the central domain of each somite (Fig.

2A). The cells positioned more lateral to the organizer will undergo dorsal convergence and

intercalate among other PSM cells to form myotome fibers that span the dorsal to ventral

extent of the trunk somites (Fig. 2B). Cells positioned in the ventral region of the blastopore

lip will undergo the most dramatic dorsal convergence towards the midline (Fig. 2C). These

cells will be found in the lateral upper and lower layers of the PSM (Fig. 2, stage 13). As the

PSM expands dorsally, these cells will eventually be located in the dorsal and ventral-most

regions of the somite (Fig. 2, stage 24). This fate map confirms an early description of this

unfolding event in X. laevis documented by Hamilton [10] and reviewed by Harland [28].

Together, these studies reveal that the position of mesoderm cells within the gastrula

predicate the formation of myotome fibers in predictable locations within segments along

the anterior-posterior axis.

1.4 Formation of the dermomyotome

The dermomyotome appears as the paraxial mesoderm begins to unfold, forming a

thin epithelial layer on the lateral surface of the PSM and somites (Fig. 2). The

dermomyotome likely originates from mesodermal tissue adjacent to the paraxial mesoderm

[10,29]. As the two-layered paraxial mesoderm unfolds, the adjacent dermomyotome

converges dorsally to reside on the surface of the PSM as a layer of unrotated cells (Fig. 2,

stages 19 and 22). Thus, unlike other vertebrates where the dermomyotome differentiates

from a subset of cells within the somite, in X. laevis, the dermomyotome originates from a

separate, adjacent mesodermal tissue that flanks the PSM. The molecular signals associated

with dermomyotome formation are discussed later in this chapter. The unique origins of the

X. laevis dermomyotome require further studies to fully understand how this tissue is

specifed and formed.

1.5 Signals that influence somite patterning

Myogenesis is an outcome of the initial patterning of somites. Morphogens secreted

by adjacent tissues subdivide each somite into discrete compartments with distinct lineages

[30] (Fig. 3). Although the molecular signals that pattern the somite are largely conserved

across vertebrates, key differences exist between organisms, particularly between direct and

indirect developers. X. laevis myogenesis is biphasic due to metamorphosis, during which

much of the larval myotome is replaced by adult muscle [1]. Nevertheless, X. laevis

myogenesis shares features in common with both zebrafish and amniotes, making it an ideal

5

system for comparing the mechanisms of somite formation between amniotes and

anamniotes.

1.6 Myogenic waves and compartmentalization of the somite

Della Gaspera and colleagues [31] proposed that X. laevis myogenesis occurs in three

successive waves (Fig. 4). The first wave arises from the PSM at the onset of somitogenesis,

with medial and lateral trunk cells expressing different combinations of muscle-specific

transcription factors and directly differentiating into mononucleated myotome cells [31]. In

amniotes, trunk and limb myotome originate from the dermomyotome, a thin layer of pax3

and pax7-positive cells in the lateral portion of the somite from which the epaxial (back

muscles) and hypaxial (body wall and limb muscles) lineages arise [32,33]. Unlike amniotes,

however, the X. laevis dermomyotome is specified during the second wave of myogenesis,

after the first wave has already established a somite comprised of differentiated myotome

[31]. This second wave gives rise to the epaxial and hypaxial lineages, and is characterized

by the expression of the full suite of muscle-specific transcription factors [31]. The third

wave consists of myf5-dependent formation of multinucleated myofibers that may originate

from pax7-positive progenitor cells [34,35]. This last wave may be the transitory stage that

defines the boundary between larval and adult myogenesis [1], and appear as puncta

localized to the anterior and posterior region of each somite.

Not much is known about X. laevis sclerotome, the somitic lineage that gives rise

primarily to bone and other associated tissues. Based on analysis of cell morphology, it may

be represented by a small population of cells in the ventromedial region of the somite by the

late tailbud stage [36]. This observation is strengthened by recent studies that identify cells

in this region expressing sclerotome markers [37]. Furthermore, sclerotome induction

appears to involve similar genes as those observed among amniotes [22]. The late

appearance of sclerotome markers (stage 35) suggests that in X. laevis, differentiation of the

vertebrae that encase the notochord and neural tube occurs later than in other vertebrates.

Together, these studies indicate that X. laevis somite formation and skeletal myogenesis are

distinct from both amniotes and zebrafish. The existence of several myogenic waves and the

delayed induction of both dermomyotome and sclerotome make X. laevis uniquely suited to

separately examine the signaling pathways associated with these processes.

1.7 Morphogens that pattern the somite

wingless-type (Wnt): Several members of the Wnt family, an evolutionarily conserved

group of growth factors, play important roles in somite patterning [38]. Wnt signaling

mediates myogenesis at multiple steps, including proliferation, differentiation, and

6

homeostasis. In mice, Wnt/β-catenin induces myoblast proliferation and fusion, and

myofiber attachment to the surrounding extracellular matrix [39,40]. Similarly, in X. laevis

β-catenin stabilization in presumptive muscle is required for proper development [41], and

attenuation of Wnt signaling results in reduced myotome formation [42]. Wnt is also directly

involved in myogenesis; an isoform of Wnt11b regulates somite formation [43]. Additionally,

dominant-negative Wnt8 blocks induction of myoD expression [44], and inhibiting Wnt8

signaling results in decreased skeletal muscle [45]. Lastly, lithium treatment, which mimics

zygotic Wnt signaling, induces an immediate-early and direct response of the myogenic gene

myf5 [46]. Consistently, factors that modulate Wnt signaling also affect myogenesis, such as

the Wnt antagonist Dkk [47] and the Wnt activator Rspondin [48]. Together, these data

support a critical role for Wnt signaling in myoblast proliferation and differentiation in X.

laevis, with different members expressed at distinct time points of myogenesis.

sonic hedgehog (shh): Shh is a morphogen secreted from the midline that is critical during

somite formation [49]. Shh signaling determines the balance between epaxial and hypaxial

muscle differentiation from the dermomyotome (Fig. 3). The epaxial myoblasts stay localized

to the somites and differentiate into the back muscles of the trunk. In contrast, hypaxial

myoblasts migrate to the limbs and ventral body wall before differentiating into hypaxial

myotome [50,51]. In X. laevis, exogenous Shh induces premature epaxial and hypaxial

muscle differentiation, whereas elimination of Shh signaling results in an expanded

dermomyotome and ectopic hypaxial muscle [52], similar to what is found in zebrafish [53]

and mouse [54].

Shh signaling highlights evolutionary innovations in vertebrate limb and trunk

myogenesis. In amniote myogenesis, Hh signaling appears to have ambiguous roles

depending on whether the environment is somitic or in the limb [54,55]. Fortunately, X.

laevis is uniquely suited to address this question since hypaxial myoblasts migrate to the

tadpole body wall and differentiate despite the absence of limbs [56]. Taking advantage of

this unique feature of X. laevis myogenesis, it became clear that the effects of Shh on amniote

myogenesis seems to be a consequence of the amniote limb environment rather than

differences in hypaxial myoblast populations, indicating evolutionary changes in the amniote

limb [52]. Another striking example of key adaptations in myogenesis is evident in the

tetrapod trunk. In zebrafish, Shh is responsible for specification of slow-twitch muscle fibers

[57] as well as the dermomyotome [53]. The same appears to be true in X. laevis, since Shh is

required for early superficial slow-twitch muscle fiber formation in the tail [58]. However, X.

laevis trunk somites do not exhibit this Shh-dependent slow-twitch myogenesis, and instead

rely on a second wave of Shh-independent myogenesis derived from the dermomyotome

[58]. Tail myogenesis (but not trunk myogenesis) in X. laevis is similar to zebrafish suggests

that the first wave of slow-twitch muscle specification is the ancestral state, and that trunk

7

somites underwent significant adaptations during tetrapod evolution [58]. Nevertheless,

Shh is crucial to the induction of myotome and dermomyotome, albeit with slightly modified

roles in different species. These and other studies highlight the value of X. laevis as a model

for understanding the evolution of developmental processes.

bone morphogenetic protein (bmp): Members of the Transforming growth factor beta

(TGF-β) superfamily of proteins such as Bone morphogenetic proteins (BMPs) pattern many

tissue types [59]. Early in X. laevis development, BMP4 acts as a ventralizing factor that must

be antagonized by secreted proteins from the organizer to establish the paraxial mesoderm

[2]. In general, BMPs have been shown to act as inhibitors of myogenesis. In amniotes,

Noggin from the dorsal neural tube and presumptive somite counteract the effects of BMP4

from the lateral plate to establish the boundary between the paraxial mesoderm and lateral

plate [60,61]. BMP4 beads implanted in the axolotl trunk inhibit myogenesis and

dermomyotome formation nearby, yet at a threshold distance, myogenesis and

dermomyotome formation are more robust; these effects are reversed by addition of Noggin

[62]. This observation is supported by experiments in X. laevis, where satellite cell induction

in the dermomyotome requires moderate levels of BMP [29]. Together, these studies support

a role for BMP wherein it prevents myogenesis while enhancing proliferation in the

dermomyotome in a gradient-dependent manner, an effect also found in zebrafish [63] and

amniotes [64].

fibroblast growth factor (fgf): Fgfs are involved in several aspects of myogenesis, including

mesoderm induction [2], paraxial mesoderm specification [65], and somitogenesis [6]. Fgfs

play a central role in the differentiation stages of myogenesis by facilitating the community

effect, a phenomenon in which differentiation progresses only when a threshold number

[66] of cells are in contact with one another. This is thought to increase the homogeneity of

cell types within a given region and to demarcate the borders between adjacent populations

of different cell types to a greater extent than what induction alone can accomplish [67]. The

community effect is readily reproducible in multipotent animal cap cells in Xenopus [68], and

is crucial for proper myogenesis [69,70]. Through dissociation experiments, it was shown

that eFGF is needed for the community effect throughout gastrulation [71]. Interestingly, the

requirement for eFgf-mediated community interaction is dependent on the developmental

age of muscle precursors: posterior presomitic mesoderm (PSM) cells require cell contact to

properly differentiate into myotome, while anterior PSM cells do not [72]. In short, Fgfs play

crucial roles in multiple steps of myogenesis, particularly in the differentiation stages by

mediating the community effect.

8

In summary, somite formation and differentiation involves a carefully orchestrated

set of morphogens. Although the signaling molecules that pattern the somitic compartments

are conserved, the timing of induction and morphogenesis is very different in X. laevis. This

provides a powerful tool to individually dissect the mechanisms behind each inductive event,

giving insights into both vertebrate development and evolution.

1.8 Transcriptional regulation of X. laevis myogenesis

Transcriptional regulation of myogenesis in vertebrates is orchestrated by the

myogenic regulatory factors (MRFs), a family of structurally-related basic-helix-loop-helix

(bHLH)-class of transcription factors that dimerize and bind E box motifs to activate

expression of muscle specific target genes [30,73].

MRFs consist of myoD, myf5, mrf4/herculin/myf6, and myogenin [74–77].

Combinations of these factors are capable of activating the myogenic fate upon ectopic

expression in fibroblasts [78]. Of these, myod and myf5 are expressed earlier and initiation

of myogenesis is therefore attributed to them, whereas mrf4 and myogenin are associated

with later aspects of differentiation (Fig. 4) [79]. Consistent with this, the combined ectopic

expression of myoD and myf5 are not sufficient to stably activate myogenesis in X. laevis,

indicating that other MRFs are required to execute the full myogenic program [80]. MRFs

self-regulate and cross-activate each other, but trans-activate different sets of myogenic

genes [81]. For example, X. laevis MyoD, Myf5, and Mrf4 each induce the expression of

different subunits of a neuromuscular junction receptor, whereas Myogenin activates a set

of muscle structural genes [82]. Such allocation of target genes among the MRFs indicates

distinct roles for each MRF during myogenesis.

Upon myogenic differentiation, MRFs are negatively regulated by the Id family of

helix-loop-helix proteins lacking the basic DNA-binding domain. Id proteins form

nonfunctional heterodimer complexes with MRFs, thus blocking activation of target genes

[83]. The conserved functions of the MRFs present benefits and challenges to dissecting

vertebrate myogenesis. Fortunately, ease of embryological and biochemical experiments in

the X. laevis system has allowed dissection of the mechanisms of myogenesis and application

of this knowledge to other species.

myogenic differerentiation (myoD): MyoD is expressed in response to muscle-inducing

factors in the PSM [84], with early transcriptional targets that include downstream

components of Notch signaling [85]. At the gastrula stage, it is expressed in the entire

marginal zone except for the prospective notochord and remains highly expressed in the

PSM and early somites throughout myogenesis (Fig. 4). By the second and third wave of

9

myogenesis, myoD expression further extends to the epaxial myotome in the dorsal somite

and the hypaxial myotome in the ventral somite and ventral body wall. X. laevis embryos

have maternally-inherited myoD, ubiquitous even in in non-mesodermal cells and

ventralized embryos [86]. X. laevis has two copies of the myoD gene [87]. Furthermore, the

protein appears to be under negative control until it is induced to translocate to the nucleus

in presumptive muscle by MAP kinase activity induced by BMP4 antagonism [87,88]. The

change in localization coincides with myoD transcript amplification in the prospective

paraxial mesoderm as a delayed response to mesoderm induction. This occurs because of

the low levels of myoD transcripts that are suddenly increased by a 100-fold, several hours

after the midblastula transition [89]. The negative regulation of MyoD appears to be

sequence and species-specific, since injection of mouse myoD mRNA into X. laevis embryos

results in a much more potent myogenic activation than does injection of X. laevis myoD [90].

MyoD regulation can be achieved in several ways, including binding of the transcription

factor Six1 to the myoD Core Enhancer Region [91], binding of the bHLH transcription factor

Mist1 to MyoD or to the E box directly [92], and binding of Hes6 to Grg2 and Grg4 to relieve

Groucho-mediated repression of gene expression [93].

MyoD initiates the temporal cascade of gene activation for skeletal muscle formation

through direct and indirect means. For instance, differential DNA-binding affinity between

MyoD and other E box-binding proteins temporally regulates muscle gene expression. Zeb1

binds G/C-centered E boxes and prevents gene expression in myoblasts but not in myotubes

[94]. This stage-dependent repression has also been demonstrated in X. laevis embryos,

where Zeb1 blocks MyoD binding in myoblasts but not in differentiating myotubes, thus

allowing temporal control of differentiation [95]. The temporal regulation of MyoD

expression is crucial to the proper timing of myoblast proliferation and differentiation, as

evident in lateral [96] and limb [97] myogenesis. Many of the mechanisms underlying myoD

regulation are similar across vertebrates, but studies in X. laevis have uncovered unique

characteristics in this system.

myogenic factor 5 (myf5): In contrast to myoD, myf5 is expressed in the dorsolateral domain

of the marginal zone in X. laevis (Fig. 4). The initial expression of myf5 depends on organizer-

mediated BMP antagonism [98]. After gastrulation, it becomes restricted to the posterior

PSM, although in lower levels than that of myoD [80]. During the second wave of myogenesis,

myf5 becomes highly expressed in the differentiating epaxial and hypaxial myotome.

Interestingly, the third wave of myogenesis is defined by myf5-dependent multinucleate

myofibers originating from pax7-postive progenitor cells [31,34,35].

The expression pattern of myf5 in the gastrula is determined through a T-box binding

site in the myf5 promoter to induce expression in the dorsal region [99], while an HBX2

regulatory element that is bound by Vent-1 represses myf5 in the ventral domain [100].

10

Additionally, myf5 is repressed in the midline through a TCF-3 binding site [101], while Smad

binding elements expand myf5 expression into the ventral marginal zone [102]. Upon

differentiation, an interferon regulatory factor-like DNA binding element represses myf5

expression in mature somites [103].

Myf5 and MyoD appear to have many features in common, but several studies have

also discovered unique functions for each. In X. laevis, injection of either myoD or myf5 mRNA

into embryos results in precocious and ectopic expression of muscle actin and myosin [104].

However, lithium treatment (which mimics Wnt/β-catenin signaling) induces ectopic myf5

but not myoD in the presence of the translational inhibitor cycloheximide, strongly

suggesting an immediate-early and preferential response of myf5 expression to zygotic Wnt

[46]. In contrast to amniote myogenesis, X. laevis MyoD is required for myf5 expression in

the early mesoderm [85]. Additionally, p38 MAP kinase is required for myf5 induction

independent of MyoD, with knockdown resulting in apoptosis and defects in segmentation

and differentiation, indicating distinct functions between Myf5 and MyoD [105,106]. This is

supported by experiments in mice suggesting that Myf5 and MyoD regulate independent

myogenic compartments [107]. In fact, the successive myogenic waves during X. laevis

somitogenesis include myf5-dependent and independent myogenic waves, further

supporting the unique contributions of myf5 during larval [31] and adult [108] myogenesis.

As mentioned above, both MyoD and Myf5 are typically associated with the specification

stage of myogenesis and have many overlapping, but also distinct functions.

myogenic regulatory factor 4 (mrf4): After the initial specification step mediated by myoD

and myf5, myogenesis shifts to myotome differentiation mediated by two additional MRFs,

mrf4 and myogenin. Mrf4, also known as Herculin and Myf6, encodes a 27-kD protein with

the bHLH motif characteristic of the other MRFs [75]. Mrf4 is expressed in differentiating

myotome, with faint expression at the late gastrula stage (Fig. 4) likely marking the initial

differentiation of myotome at the medial edge of the somite [58]. As somitogenesis

progresses, mrf4 becomes highly expressed as stripes corresponding to differentiating

myotome in somites. This strong expression continues throughout the second and third

waves of myogenesis in both epaxial and hypaxial lineages.

Although the transcriptional activation domains of Mrf4 and MyoD are mostly

interchangeable, amino acid differences in these regions may account for their interactions

with different co-regulator proteins and the specificity of their functions [109]. In mammals,

the mrf4 gene has a complex cis-regulatory structure that includes proximal and distal

regulatory elements that pattern mrf4 gene expression in early and late myogenic cells

[110,111]. Functional comparisons with mammalian mrf4 indicates that X. laevis mrf4 has a

610bp proximal promoter that includes an enhancer; over 300bp of this region is conserved

11

in both X. laevis and X. tropicalis and contains a MEF2 binding site essential for expression of

the protein [112]. Mrf4 knockout in mice results in normal myogenesis and upregulation of

Myogenin, suggesting that Myogenin can compensate for Mrf4 loss. Furthermore, mrf4 and

myoD double knockouts in mice result in severe muscle deficiency, although single

knockouts of each are viable. This suggests that they may have overlapping functions that

cannot be compensated for if both factors are eliminated [113].

Although many Mrf4 experimental phenotypes are similar between mammalian

systems and X. laevis, there are crucial functional differences at the molecular level. For

instance, within the 610bp X. laevis mrf4 promoter region, only about 150bp is conserved in

mammals; this difference in transcriptional control elements indicates divergent evolution

of myogenic gene regulatory elements [112]. During the formation of neuromuscular

connections in X. laevis mrf4 expression may be induced by innervation; in fact, denervation

of adult muscle results in reduced mrf4 but not myoD RNA levels [114,115]. However, a

subsequent study shows that surgical removal of the brain prior to motor axon outgrowth

had no effect on mrf4 expression levels [116]. The upregulation of mrf4 expression during X.

laevis muscle regeneration has also been shown to be independent of innervation [114].

Studies on Mrf4 further highlight the complex relationship between the MRFs, yet there is a

clear trend of a more central role in myotome differentiation rather than specification,

especially considering the timing and localization of mrf4 expression in mature myotome.

myogenin (myog): Myogenin was first shown to induce muscle-specific markers in

mesenchymal cell lines [77] and in mouse embryos [117]. In X. laevis, it was first shown to

be expressed in forming myotubes during muscle regeneration [118]. Myogenin is faintly

expressed at the neurula stage, and localizes to the more mature myotome in anterior

somites (Fig. 4). This expression continues throughout the second and third waves of

myogenesis in the epaxial and hypaxial lineages, in a pattern that overlaps with mrf4

expression. The localization and timing of myogenin strongly suggest a role for the

differentiation stages of myogenesis.

Reduction of mrf4 expression induced by denervation results in increased myogenin

expression, suggesting a compensatory role [115] particularly during regeneration [119].

Myogenin also plays a central role during X. laevis metamorphosis [120] by mediating

myoblast fusion [121] and activating adult myogenic genes [122]. Of all the MRFs, Myogenin

seems to be the least ambiguous in function, with a clear role in latter stages of myogenesis.

As with the morphogens described in the previous section, the myogenic

transcription factors are largely conserved across vertebrates. However, X. laevis exhibits a

few distinct features and thus, has helped to elucidate the specific roles of the MRFs. For

instance, the different combinations of MRFs expressed during each wave of myogenesis,

12

particularly the myf5-dependent third wave, supports the idea that each MRF has a unique

function in muscle specification and differentiation [31]. This and other features of X. laevis

myogenesis greatly facilitates the study of vertebrate MRFs.

1.9 Muscle Satellite Cells and Regeneration

Muscle satellite cells (MSCs) differentiate into muscle upon injury, or self-renew to

maintain a pool of MSCs [123–129]. MSCs reside under the basal lamina of muscle fibers and

are marked by pax7 and pax3 expression, which are both required for MSC survival [130–

133]. In chick and mice, MSC progenitors originate from the dermomyotome and give rise to

trunk and limb muscles [130,131,134]. Recent experiments using continuous conditional

inactivation of Pax7 expression reported defective muscle regeneration due to a reduction

in the number of proliferating MSCs in mice [135,136]. In X. laevis, Pax7 has been shown to

be required for the survival and proliferation of MSCs in the tadpole. Researchers showed

that in the absence of Pax7 function, the number of MSCs decrease, which then leads to the

inability of muscle to regenerate in the amputated tail of the tadpole [30]. Fate mapping

experiments have shown that MSCs originate from the dorsal region of mesoderm tissue that

lies lateral to the paraxial mesoderm and fated to become dermomyotome [24]. MSC

specification appears to begin at the neurula stage through a ventral to dorsal gradient of

BMP signals [24]. Thus, MSC formation and maintenance in X. laevis are quite similar to those

described in amniotes.

Amphibians have emerged as a leading animal model to investigate the cellular and

molecular mechanisms involved in regeneration of tails and limbs in the adult [137–139].

The mechanisms that govern regeneration are evolutionarily diverse. For example,

salamanders regenerate limbs via a progenitor pool called the blastema [140]. During

blastema formation, urodele amphibians replace lost limbs through dedifferentiation, a

mechanism wherein a terminally differentiated cell reverts back to a less differentiated state.

These dedifferentiated cells will then subsequently redifferentiate to replace lost tissue in a

lineage specific manner [141]. In the eastern newt, Notophthalmus viridescens, muscle

dedifferentiation makes a significant contribution to muscle regeneration. However in the

axolotl, Ambystoma mexicanum, Pax7-expressing MSCs are the main contributor to limb

muscle regeneration [142]. Unlike urodeles, adult anurans lose their ability to regenerate

lost limbs. In post-metamorphic stages, amputated limbs will form a muscle-less shaft

lacking segmented digits (i.e. a spike) [143]. Interestingly, X. laevis tadpoles retain the ability

to regenerate, but this ability is progressively lost as they reach metamorphosis [144–147].

Moreover, X. laevis tadpoles rely on pax7 positive MSCs to regenerate [35,148].

Studies in X. laevis have helped us to understand the signaling pathways involved in

regeneration. It was previously established that BMP, Wnt, Fgf and Shh are expressed during

tadpole limb regeneration [144–147]. Thus, maintenance of the expression of key signaling

13

molecules appears essential in maintaining the ability to regenerate [134,135]. Lin and

colleagues [149] showed that larval limb progenitor cells overexpressing Wnt/β-catenin

grafted to the amputated limb in post-metamorphic hosts are able to regenerate the limb,

suggesting that the activation of the Wnt pathway is essential for maintaining the ability to

regenerate the limb. These grafted progenitor cells also required the presence of exogenous

Shh, Fgf10, and thymosin beta 4 (Tb4) to form functional limb regenerates consisting of

innervated muscle and bone. This study indicates that a myriad of molecular signals are

necessary for proper limb regeneration in adult X. laevis. Another study in X. laevis

demonstrated that tadpole tail regeneration

requires the sustained production of reactive oxygen species (ROS) [150]. High levels of ROS

were shown to be required for maintaining an active Wnt/β-catenin signal, which in turn

activate downstream targets, such as fgf20, an important regulator of tissue regeneration

[150]. Together, these studies highlight X. laevis as a unique model to study the process of

regeneration, particularly given the wide range of microsurgical, transgenic, and

transcriptomic approaches available in this organism.

14

CHAPTER 2: Tissue morphogenesis and cell rearrangements leading to somite formation

2.1 Abstract

Cell movements and tissue morphogenesis shape both individual organs and the embryo as

a whole. After gastrulation in X. laevis, mesoderm cells of the marginal zone give rise to

myotome located in specific regions of somites along the anterior-posterior and dorsal-

ventral axes. While the origin of myotome fibers in the tadpole have been fate-mapped in the

gastrula, the morphogenetic movements that determine their final positions are unknown.

Here, we show the cell movements that marginal zone cells undergo as they form the pre-

somitic mesoderm (PSM) and myotome of the somites. We describe the unfolding of PSM

tissue, which positions muscle cells from the lateral edge of the PSM to the dorsal and ventral

domains of somites. This unfolding is followed by rearrangements of clusters of elongated

cells to be perpendicular to the notochord along the dorsal-ventral plane. We observed that

rearrangement of these clusters of cells is accompanied by the deposition of extracellular

matrix proteins including fibronectin and laminin in key locations within the tissue. Finally,

our data indicate that both PSM unfolding and cell rearrangements occur largely

independently of the non-canonical Wnt/PCP pathway. Together, we reveal two fascinating

morphogenetic phenomena distinct from amniote somitogenesis that highlight the crucial

roles that cell movements and tissue morphogenesis play during embryo development.

2.2 Introduction

Somite morphogenesis in Xenopus laevis consists of cell behaviors different from amniotes

and other vertebrates. First described by Hamilton (1969), a dual layer of pre-somitic

mesoderm (PSM) tissue with a narrow myocoel forms as marginal zone cells involute during

gastrulation [10]. This PSM tissue then unfolds dorsally and ventrally, eliminating the

transient myocoel. After unfolding, individual myotome cells undergo a 90˚ rotation to

become aligned parallel to the notochord [17, 28]. This pattern of somite cell rotation is also

found in two other pipid species, Xenopus tropicalis and Hymenochirus boettgeri and not in

other anuran species, suggesting that this pattern may be a shared derived character for

Pipidae [12]. The absence of a myocoel in mature X. laevis somites is quite different from

amniote somite morphogenesis, wherein the somite forms a rosette with a clear myocoel [6].

In addition, somite cell rotation has also been described in zebrafish and serves to establish

lineage-restricted subsomitic compartments [13].

Myotome rotation requires deposition of extracellular matrix proteins in the intersomitic

and notochord-somite boundaries to allow proper attachment and alignment of myotome

during rotation [21, 158, 159]. After rotation, the positions of myotomes in X. laevis are

15

finalized and the somite is compartmentalized into different lineages according to the signals

somitic cells receive from adjacent tissues [52, 58, 158]. Similar to zebrafish, X. laevis

myotome positioning is mediated by the Sdf family of secreted cytokines [13, 14].

Interestingly, the final positions of myotome within the anterior-posterior axis of the tadpole

and the dorsal-ventral axis of individual somites have been fate-mapped and shown to be an

outcome of their initial positions along the blastopore lip at the onset of gastrulation [27].

Although myotome rotation and positioning have been described and investigated in detail,

not much is known about somite unfolding besides Hamilton’s brief description in 1969. In

this report, we describe in greater detail pre-somitic mesoderm (PSM) tissue unfolding,

which occurs immediately prior to myotome rotation. We show that at the onset of

gastrulation, cells from the marginal zone involute into the blastopore and form specific

regions of the pre-somitic mesoderm (PSM) tissue. Upon forming the PSM, cells initially in

the lateral edge of the tissue then become positioned to the dorsal and ventral domains of

the somite as the tissue unfolds, demonstrating the role that somite unfolding plays in

myotome positioning. Furthermore, we describe a complex series of cell rearrangements

distinct from myotome rotation accompanied by dynamic expression of extracellular matrix

anchor points. Finally, we show that both PSM unfolding and cell rearrangements occur

autonomously in the absence of adjacent tissues such as the notochord and neural tube. We

hypothesized that both PSM unfolding and cell rearrangements are mediated by the non-

canonical Wnt/PCP pathway, which is necessary for establishing cell polarity and

coordinating convergent extension movements [172]. Remarkably, PSM unfolding and cell

rearrangements are only mildly affected by targeted expression of dominant-negative

canonical Wnt and noncanonical Wnt/PCP components, suggesting that these phenomena

are regulated by a different pathway. Together, these data describe PSM unfolding and cell

rearrangements, and deepen our understanding of somite morphogenesis.

2.3 Results

Marginal zone cells involute and form the presomitic mesoderm

The initial positions of marginal zone cells of the gastrula determine their final position as

myotome within somites [27]. For instance, mesoderm cells immediately adjacent to the

prospective notochord lateral to the upper blastopore lip (ULL=upper lateral lip) form

myotome in the central domain of somites along the entire anterior-posterior axis. On the

other hand, mesoderm cells opposite the prospective notochord (LL=lower lip) form

myotome in the dorsal and ventral domains of somites. Although the final positions of ULL-

and LL-derived myotome have been fate-mapped, the morphogenetic trajectories that place

them there remain unknown.

16

To determine the paths that marginal zone cells take as they form myotome, we made

homotopic and homochronic grafts of RDA-labeled cells into the ULL (Fig 1A-1D) and the LL

(Fig 1E-1H) regions, then tracked the progress of grafted cells over time in fixed embryos via

confocal microscopy. Cells grafted to the ULL region (Fig 1A) of the gastrula involute through

the blastopore and remain adjacent to the notochord at the early tailbud stage. Transverse

sections reveal RDA-labeled cells in the central domain of the developing somite (Fig 1C),

while lateral (Fig 1B) and dorsal views (Fig 1D) show labeled cells adjacent to the notochord

undergoing previously-described cell rotation as they differentiate into myotome [17].

In contrast, RDA-labeled cells grafted to the LL region (Fig 1E) of the gastrula must take a

less direct path to the dorsal region of the embryo. Transverse sections (Fig 1G) reveal that

LL cells must travel along the lateral edge of the embryo (dashed circle is archenteron cavity)

after involution through the blastopore (yellow asterisk). Lateral (Fig 1F) views show that

LL cells must traverse from the ventral region of the blastopore (yellow asterisk) to the

dorsal region of the embryo. Dorsal views (Fig 1H) show that, upon reaching the dorsal

region, LL cells stop at the lateral edge of the pre-somitic mesoderm (PSM). Cells from the

LAT region, which are located between the ULL and LL regions of the gastrula, exhibit

behaviors similar to both ULL and LL cells (Fig 10). The movements of marginal zone cells to

the presomitic mesoderm is schematized in the posterior region of the tailbud embryo in

(Fig 9A).

Myotome forms in specific domains within the somite based on their initial position

Upon reaching the dorsal part of the embryo, both ULL and LL cells form myotome in distinct

domains of the somite, albeit at different time points. Cells grafted to the ULL region (Fig 2E)

of the gastrula form myotome in the central domain of somites along the entire anterior-

posterior axis by the late tailbud (Fig 2A) and tadpole (Fig 2C) stages. Transverse views of

late tailbud (Fig 2B) and tadpole (Fig 2D) stages reveal that grafted ULL cells remain adjacent

to the notochord and span the medial-lateral axis of the entire somite.

On the other hand, cells grafted to the LL region (Fig 2F) of the gastrula form myotome in the

dorsal and ventral domains of posterior-trunk and tail somites, but not until the tadpole

stage, as seen in lateral (Fig 2J) and transverse (Fig 2K, magnified in Fig 2L) views.

Interestingly, grafted LL cells appear to split into dorsal and ventral populations in the late

tailbud stage as seen from a lateral view (Fig 2H, magnified in Fig 2G), with some cells

appearing in the lateral edge of the somite as seen in the transverse section of the same

embryo (Fig 2I).

Pre-somitic mesoderm tissue unfolds dorsally and ventrally as it forms somites

17

Given the lateral movement of cells to reach the PSM (Fig 2L), we decided to examine the

pre-somitic mesoderm (PSM) tissue more closely. Since the 12/101 antibody used in (Figs 1

& 2) only detects fully-differentiated myotome in mature somites, we turned to phalloidin

staining to view the cell outlines of PSM tissue. Using sequential transverse sections, we

discovered that the PSM tissue undergoes extensive morphogenesis from the neurula to

tadpole stages. Based on our previous lineage tracing and on the repositioning of grafted

cells in (Fig 4), we deduced that PSM unfolding is characterized by the repositioning of lateral

PSM tissue to the dorsal and ventral domains of the somite (white asterisks in Fig 3). At the

early neurula stage, the PSM tissue appears as a solid, rectangular block of cells (Fig 3A)

ventral to the neural plate (blue arrowhead) and flanking the developing notochord (blue

dashed circle). As the neural plate folds (Fig 3B, blue arrow head), the underlying PSM tissue

changes into a more circular shape, and myocoels appear to divide the future dorsal and

ventral thirds of the somite (Fig 3B, white arrowheads). These myocoels eventually open to

the lateral sides as the tissue unfold (white arrowheads, Figs 3C-3D). When the PSM is almost

fully unfolded (Fig 3F), the individual myotomes then rotate 90˚ posteriorly (previously

described in [17]), as can be seen in the differences in apparent cell morphology between

(Fig 3F) and (Fig 3G). After rotation, concentrations of actin appear in the tissue (white

arrowheads in Figs 3G), presumably as cells change shape to bend the unfolding tissue. As

the somite fully unfolds, phalloidin fluorescence vastly increases in the mature myotomes

(Figs 3H-3I). The actin bands described in (Figs 3G) are reduced by the tadpole stage (white

arrowheads in Fig 3H) and eventually disappear in a mature somite (Fig 3I). The

morphogenetic movements of PSM tissue unfolding are schematized in (Fig 3J), with A’-I’

corresponding to Figures 3A-3I, respectively.

Tissue unfolding positions PSM cells to the dorsal and ventral regions of the somite

Based on the position of the transplanted cells in (Fig 2G) and the observed phenomenon of

PSM unfolding (Fig 3), we hypothesized that PSM unfolding positions cells from the LL region

to the dorsal and ventral domains of somites. To test this hypothesis, we repeated the

grafting experiment from (Figs 1E & 1F) and followed the trajectory of cells transplanted in

the LL region of the gastrula. This time, however, we looked not at the mature somites, but

at the unfolding PSM. We found that after gastrulation, LL cells populate the lateral edge of

the PSM (Fig 4A), as suggested by earlier tracking experiments (Figs 1G & 1H). As the PSM

unfolds, transplanted cells initially in the lateral edge of the tissue are separated to the dorsal

and ventral thirds of the PSM (Fig 4B). As the somite fully unfolds, transplanted cells can be

found in both the dorsal and ventral domains (Figs 4C & 4D), with actin concentrations in

the tissue previously observed in (Figs 3F-3H). Finally, by the tadpole stage, the somite is

fully mature, with cells initially grafted to the LL region of the gastrula giving rise to myotome

in the dorsal and ventral domains of the somite (Fig 4F).

18

Cells within the PSM undergo dramatic rearrangements after tissue unfolding

As the somite forms, elongated myotome rotate 90 ֯ along the medial-lateral plane to become

align parallel to the notochord [17]. However, we observed yet another series of complex cell

rearrangements that occur immediately following PSM unfolding, but before the myotome

rotation described by Afonin and colleagues [17]. While folded, PSM cells and their nuclei

are uniformly arranged in a dual layer (Fig 3A, depicted in Fig 5M-I). As the tissue unfolds,

PSM cells elongate along the medial-lateral axis and localize their nuclei to the periphery of

the tissue (Fig 5A-C, depicted in Fig 5M-II and Fig 5M-III). Once the tissue unfolds, PSM cells

then become arranged perpendicular to the notochord along the dorsal-ventral plane, and

are grouped into three clusters (Fig 5D-F, depicted in Fig 5M-IV), mirroring the eventual final

positions of LL cells in the dorsal and ventral domains of the somite (Fig 4), and of ULL cells

in the central domain of the somite (Fig 2D). The boundaries of these clusters are delineated

by actin concentrations (white arrowheads in Fig 5D); furthermore, the nuclei of cells within

each cluster become organized into distinct bands (white arrowheads in Fig 5E). After this

conformation, the elongated PSM cells then undergo the rotation described in [17] to become

aligned parallel to the notochord (Fig 5G-I). Soon after rotation, the PSM tissue retain the

actin concentrations (white arrowheads in Fig 5G), as well as the bands of nuclei (white

arrowheads in Fig 5H). In a fully-mature somite (Fig 5J-L, depicted in Fig M-V), however, the

actin concentrations are no longer present (Fig 5J), and the nuclei become uniformly

arranged throughout the tissue (Fig 5K), though aligned in the middle of each elongated cell

[17]. Together, these data add yet another step in the complex morphogenesis that the PSM

tissues undergo as they mature into somites.

PSM cell rearrangements are accompanied by transient ECM anchor points

Cell rotation during X. laevis somitogenesis requires step-wise deposition of extracellular

matrix (ECM) proteins such as fibronectin and laminin (reviewed in [158]). Therefore, we

wanted to investigate if the cell rearrangements following somite unfolding also use

depositions of ECM proteins. By staining serial sections of early and late tailbud embryos, we

show that Fibronectin (Figs 6A-6J), β-integrin (Figs 6P-6Q), Laminin (Figs 6S-6T), and β-

dystroglycan (Sup Fig 2) are all expressed dynamically throughout PSM unfolding. As the

PSM finishes unfolding, Fibronectin and Laminin are deposited to span the width of the

tissue (white arrowheads in Figs 6H-6I & Fig 6T). At this stage, PSM cells express both β-

integrin and β-dystroglycan (Fig 6Q & Sup Fig 2). During PSM cell rearrangements, these

depositions of Fibronectin and Laminin presumably serve as anchor points for cells to attach

as they become perpendicular to the notochord in the dorsal-ventral plane (Fig 5D). Lastly,

as the cells undergo the rotation described in [17], the ECM anchor points disappear (white

19

arrowhead in Fig 6J). The transient expression of ECM anchor points coincides with the cell

rearrangements and organization of nuclei described in (Fig 5). Co-localization of nuclei and

ECM hinges in the same transverse sections demonstrates this observation more clearly

(Figs 6C-6D).

PSM tissue unfolding and cell rearrangements occur without the influence of adjacent

tissues

Differentiation of somite derivatives require a carefully orchestrated release of signaling

molecules secreted from adjacent tissues (reviewed in [158]). Furthermore, morphogenesis

often requires the mechanical contributions of key tissues, such as the notochord for axis

elongation. We therefore wanted to determine which tissue(s) are necessary for proper PSM

unfolding. To remove the physical and signaling influence of the notochord, we UV-irradiated

zygotes prior to cortical rotation, thus inhibiting the formation of Spemann’s Organizer and

the notochord (Fig 7A). By the late tailbud stage, irradiated embryos with a Dorsoanterior

Index (DAI) number of 2 were selected for analysis [160-163]. These embryos are

microcephalic, have truncated axes, and no notochord, but retain their neural tube and

somites (Fig 7E). Remarkably, UV-irradiated embryos undergo PSM unfolding (white

asterisks in Figs 7B-7D), but are fused at the midline due to the absence of the notochord

(white arrowhead in Fig 7B points to neural plate). The PSM cell rearrangements described

in (Figs 5 and 6) still occur, as evident in the presence of Fibronectin through the tissue

(white arrowheads in Figs 7F & 7G), and its eventual dissipation from the middle of the tissue

once the somite is fully unfolded and mature (Fig 7H).

Next, we wanted to determine if physical forces exerted by axis elongation causes PSM

unfolding. Although the PSM itself undergoes extensive convergent extension, adjacent

dorsal tissues such as the notochord and the neural tube also contribute extensively to axis

elongation, particularly from the late gastrula to the neurula stages, when anterior somites

are beginning to form [15]. Therefore, we eliminated the physical influence of both the

notochord and neural tube by surgically removing the progenitors of both tissues at the

gastrula stage [27]. By the late tailbud stage, dissected embryos exhibit dorsal flexion and

truncated axes (Fig 7M). However, the anterior somites of dissected embryos still undergo

unfolding (asterisks in Figs 7J-7L). The ECM anchor points necessary for PSM cell

rearrangements also remain unperturbed (Figs 7N-7P), as evident in the organized

deposition of Fibronectin in the PSM (arrowheads in Figs 7N & 7O), and their disappearance

upon completion of somite unfolding (Fig 7P).

Another significant driver of axis elongation, particularly from the neurula to the tailbud

stages, is the autonomous convergent extension of ventral mesoderm and epidermal tissues

[164]. To remove the physical influence of ventral tissues, we removed the ventral tissue of

20

embryos at the early neurula stage and cultured the dorsal explants until late tailbud stage

(Fig 7Q). Both dorsal and ventral explants elongated autonomously (Fig 7U, ventral tissues

at the bottom row). However, dorsal explants did not elongate as much as control embryos,

indicating that removal of ventral tissues eliminated the influence of ventral tissues from

posterior somites [164]. Therefore, this allowed us to examine posterior somites

independent of the convergent extension of ventral tissues. Interestingly, posterior somites

in dorsal explants still undergo somite unfolding (asterisks in Figs 7R-7T), evident in the

clefts in the PSM (arrowheads in Fig 7R) that separate the future dorsal and ventral thirds of

the tissue, as previously described (white arrowheads in Figs 3B-3D). Dorsal explants also

successfully completed myotome rotation to form a mature somite (Figs 7S-7T). Fibronectin

hinges are also present in the unfolding somites of dorsal explants (white arrowheads in Figs

7V-7W), which disappear once the somite is fully unfolded and mature (Fig 7X). Together,

these data indicate that both PSM unfolding and cell rearrangements occur despite the

absence of adjacent tissue crucial for axis elongation.

Autonomous PSM tissue unfolding and cell rearrangements are not mediated by the

Wnt/PCP pathway

Non-canonical Wnt signaling regulates the polarity of intercalating mesoderm cells as they

participate in convergent extension. Targeted expression of Xenopus Dishevelled (Xdsh) to

the prospective paraxial mesoderm results in a “short and stout” phenotype due to inhibition

of axis elongation [165]. Based on the stereotyped cell arrangements we observed during

PSM unfolding (Figs 5K-5O), we hypothesized that non-canonical Wnt might coordinate PSM

cell movements as the tissue unfolds. To test this hypothesis, we visualized somite unfolding

in Xdsh deletion mutants. To uncouple the signaling properties of the protein, we used the

Xdd1 mutants to disrupt both canonical and non-canonical signaling pathways, and Xdsh-D2

mutants to disrupt only non-canonical Wnt while leaving canonical Wnt fully functional

[166-168]. Injection of 1ng of either Xdd1 or Xdsh-D2 mRNA into both dorsal vegetal

blastomeres at the 8-cell stage (Figs 8A & 8I) results in a short and stout phenotype with

some dorsal flexure (Figs 8E & 8M), as previously published [165]. Interestingly, transverse

sections of both Xdd1 (Figs 8B-8D) and Xdsh-D2 (Figs 8J-8L) mutants reveal that somite

unfolding proceeds with only minor defects. In both cases, inhibiting the non-canonical Wnt

pathway elicited a phenotype in which one set of somites is separated from the midline due

to convergent extension defects (Figs 8C & 8K). Furthermore, Xdd1 mutants also exhibited

defects in somite differentiation, as to be expected when canonical Wnt is inhibited (Sup Fig

3). Nevertheless, PSM unfolding occurs in Xdsh deletion mutants, even when the Xdsh-D2

dosage was increased to the point of causing open neural tube defects (Sup Fig 3).

Fibronectin deposits were also present in both Xdd1 (arrowheads in Figs8F-8G) and Xdsh-

D2 (arrowheads in Figs 8N-8O) -injected embryos. In addition, alignment of the myotome

21

nuclei was unaffected in both cases (arrowheads in Figs 8H & 8P). Together, these data

suggest that both PSM unfolding and PSM cell do not require a functional PCP pathway.

2.4 Discussion

PSM unfolding drives positioning of myotome in the dorsal-ventral axis

Cells from the marginal zone of the gastrula eventually form myotome in specific domains

along the anterior-posterior axis of the tadpole and the dorsal-ventral axis of somites [27,

158]. In addition, the cytokine SDF-1α and its receptor CXCR4 have been shown to mediate

the cell behaviors required for marginal zone cells to reach the dorsal part of the embryo as

well as those that organize myotome fibers into parallel arrays within somites [14]. Here, we

describe the morphogenetic movements that position marginal zone cells of the gastrula into

specific domains of the somite as myotome (Fig 9).

During gastrulation, cells positioned in the Upper Lateral Lip (ULL), Lateral Lip (LAT), and

Lower Lip (LL) of the blastopore involute and begin forming the presomitic mesoderm

(PSM). Cells in the dorsal lip will form the future notochord; cells from the ULL region form

the medial portion of the PSM, cells from the LL region form the lateral portion of the PSM,

and cells from the LAT region become peppered throughout the PSM (Fig 1, Supp Fig 1).

While both ULL and LAT cells maintain their positions in the PSM as the somite matures, LL

cells located in the lateral edge of the PSM eventually form the dorsal-most and ventral-most

myotome of the somites (Fig 2). Closer inspection of the PSM reveals that the tissue

undergoes remarkable morphogenetic movements described here as “unfolding” (Fig 3). As

the PSM matures, the lateral portion of the tissue – where LL cells are located – unfolds

dorsally and ventrally, eventually placing LL cells to the dorsal and ventral domains of the

somite (Fig 4). This is followed by a series of cell rearrangements that eventually organize

cells into three clusters – dorsal, ventral, and central – comprised of elongated cells

perpendicular to the notochord along the dorsal-ventral plane. Interestingly, the clusters are

organized such that cells from the ULL, LAT, and LL positions maintain their positional

identities despite extensive cell rearrangements. Notably, both PSM unfolding and cell

rearrangements occur regardless of the absence of adjacent tissues. Together, these data

provide a deeper understanding of the morphogenetic events that place cells from their

initial positions as marginal zone cells in the gastrula and into their final positions as

myotome in somites, as summarized in (Fig 9).

22

Cell rearrangements after PSM unfolding are accompanied by extracellular matrix

protein anchor points

Cell rotation during somitogenesis has been previously reported, including whole-somite

rotation observed in zebrafish [13] and individual myotome rotation described in X. laevis

[17]. Orchestrating X. laevis myotome rotation requires carefully-timed signaling cues as

well as step-wise expression of key extracellular matrix proteins, such as Fibronectin [14,

21, 158]. Deposition of ECM proteins in the intersomitic and notochord-somite boundaries

is particularly important to provide crucial attachment points for adjacent tissues as well as

for rotating myotome [159]. Here, we characterize another series of cell rearrangements

distinct from the myotome rotation described by [17]. Like myotome rotation, PSM cell

rearrangements are accompanied by the deposition of extracellular matrix proteins at key

locations. During unfolding, PSM cell nuclei rearrange themselves from a uniform

distribution (Fig 6K) to being localized to the periphery of the tissue (Fig 6L), and finally

become organized into distinct bands (Figs 6M-6N). The positions of nuclei in between the

ECM depositions (Figs 6C-6D) suggest that PSM cells use them as focal adhesion points to

orient and leverage themselves during rearrangement. This behavior is comparable to that

of myotome rotation, in which cells anchor to intersomitic ECM to align themselves parallel

to the notochord. However, the difference is that ECM anchor points, which are localized

across the PSM tissue, are much more transient than the ECM found in the intersomitic and

notochord-somite boundaries. Perisomitic ECM remains for as long as somites are present,

presumably to keep myotome anchored in place. On the other hand, ECM anchor points

appear to be transient, and disappear once the PSM has fully unfolded and rotated. Such

differences highlight the complex and dynamic nature of somite morphogenesis in X. laevis.

PSM unfolding and cell rearrangements occur autonomously and do not require the

Wnt/PCP pathway

Cell intercalation and convergent extension is a well-characterized behavior of mesoderm

tissue in X. laevis [169-171]. Convergent extension of mesoderm tissue is autonomous, and

proceeds even when very little other tissue is present, as when two dorsal marginal zone

explants in a Keller Sandwich and elongates [169]. To test the autonomy of somite unfolding,

we performed a series of manipulations that systematically removed the signaling and

biomechanical influence of tissues adjacent to the PSM. Initially, we hypothesized that the

chief physical influence of adjacent tissues on somite unfolding is an inward “squeezing”

force generated by the notochord when axis elongation changes the shape of the embryo

from a sphere into a narrow tube. However, we found that removal of the notochord (Figs

6A-6H), as well as removal of both the notochord and neural tube (Figs 6I-6P) had no effect

on somite unfolding. Removal of ventral epidermis and gut (Figs 6Q-6X) also had no effect.

Additionally, targeted inhibition of either the canonical Wnt or the noncanonical Wnt/PCP

23

pathways had no effect on somite unfolding aside from minor differentiation and

morphological defects (Fig 7), an interesting finding considering the role of the Wnt/PCP

pathway in coordinating convergence and extension [172]. Together, these data further

demonstrate the morphogenetic autonomy of mesoderm tissue.

2.5 Methods

Embryo Culture and Microinjections

X. laevis females were injected with 0.5ml human chorionic gonadotropin (HCG) hormone to

induce ovulation. Eggs were collected 15 to 20 hours post injection and in vitro fertilized

with X. laevis male sperm in 1/3X MR at 12°C or 16°C. Fertilized embryos were then dejellied

using 3% cysteine solution, then cultured in 1/3x MR. Donor embryos for transplantation

experiments were injected with a Drummond Nanoject II Automatic Oocyte Injector at the

1-cell stage with 9.2nl of 70,000 MW rhodamine dextran amine (RDA) in 2.5% Ficoll. Injected

embryos were then transferred to 1/3x MR and cultured at 12°C. For PSM unfolding

experiments, embryos were injected at the 8-cell stage using a Picospitzer II (General Valve

Corporation) and microinjection manipulator (Narishige Group). Constructs used were

Xdsh-ΔPDZ/D2 (Xdsh-D2) and Xdd1 [166-167].

Cell Transplantations

RDA-labeled donor and unlabeled host embryos were allowed to develop to gastrula stage.

The protective vitelline envelope was removed from both host and donor embryos using

forceps. Eyebrow and eyelash tools were used to remove approximately 30–50 RDA-labeled

cells from donor embryos. Cells were transplanted to host embryos on 2% agar coated Petri

dishes containing 1× MBS and then transferred to 2% agar coated Petri dishes containing

1/3× MBS for long-term culture. At various stages, embryos were fixed in 1× MEMFA

(100mM MOPS pH 7.4; 2mM EGTA, 1mM MgSO4; 3.7% formaldehyde) for one hour at room

temperature or overnight at 4°C. Fixed embryos were transferred to 100% methanol for

long-term storage.

Tissue manipulations

To remove the notochord, embryos at the one cell stage were irradiated with ultraviolet light

using an inverted Stratalinker® UV Crosslinker with the embryos placed on microscope

slides. The amount of energy emitted ranged from 10,000 KJ to 120,000KJ and lasted for 30-

90 seconds. Embryos were kept on a cold bench with ice to prevent cortical rotation while

24

performing the experiment. Embryos were then cultured in 1/3X MR at either 12°C or 16°C

to tailbud stages 25 -28. Embryos were fixed with PFA (10% 1X MEMFA salts, 10%

Formaldehyde, 80% H2O) for two hours at room temperature or overnight at 4°C. Embryos

were also fixed with DENTS fixative (80%MeOH, 20% DMSO) overnight at 4°C. Fixed

embryos were transferred to 1X PBS for storage.

For notochord and neural tube extirpations, the gastrula fate map [24] along with the

position of dorsal bottle cells was used to predict the position of the prospective notochord

at the onset of gastrulation at stage 10.5. The vitelline envelope was removed with sharpened

forceps. Micro knife tools were then used to excise the prospective notochord region from

stage 10.5 embryos. Extirpated embryos were then cultured to stagees 25-28 in 1/3X MR.

Embryos were fixed with PFA ( 10% 1X MEMFA salts, 10% Formaldehyde, 80% H2O) for

two hours at room temperature or overnight at 4°C. Fixed embryos were transferred to 1X

PBS for storage.

For dorsal and ventral tissue extirpations, the gastrula fate map was used to predict the

position of the prospective dorsal and ventral tissue at the neurula stage (15-16). The

vitelline envelope was removed with sharpened forceps. Eyelash and micro knife tools were

then used to follow the groove of the embryo, cutting through the blastopore, the lateral body

wall, and the prospective mouth region, ventral to the anterior neural folds [164]. Extirpated

embryos were cultured in 1/2X NAM (10mL 10XNAM, 2.7mL 100mM Phosphate Buffer, 1mL

100mM NaHCO3, 1.5mL 50mg/mL GENT, 185.5mL H2O). Isolated dorsal and ventral pieces

were then transferred to 1/3MR at 12°C or 16°C to collect tailbud stages 25 -28. Embryos

were fixed with PFA (10% 1X MEMFA salts, 10% Formaldehyde, 80% H2O) for two hours at

room temperature or overnight at 4°C. Fixed embryos were transferred to 1X PBS for

storage.

Immunohistochemistry

To visualize the presomitic mesoderm, embryos were sectioned into 200µm segments and

incubated overnight with 1:40 Phalloidin (Alexa Fluor): PBSt. To visualize the notochord and

mature somites, whole embryos were stained with 1:250 12/12101 antibody

(Developmental Studies Hybridoma Bank) and 1:5 Tor70 antibody (Harland Group),

respectively. For secondary antibodies, 1:250 Alexa Fluor 488 goat anti-mouse IgG and 1:250

Alexa Fluor 647 goat anti-mouse IgM were used.

To visualize extracellular matrix depositions, the following antibodies from the

Developmental Studies Hybridoma Bank were used on whole embryos: 1:50 8C8 (β-

integrin), 1:100 7DII (β-dystroglycan), and 1:100 MT4 (fibronectin). For secondary

antibodies, 1:250 Alexa Fluor 488 goat anti-mouse IgG or 1:250 Alexa Fluor 555 goat anti-

mouse IgG were used. 1:100 laminin antibody (Sigma Aldrich) with 1:250 Alexa Fluor 555

25

goat anti-rabbit IgM secondary antibody was also used. Antibodies were diluted with 1:10

cas-block (Life Technologies): PBSt.

Imaging

Whole embryos were cleared with 2:1 benzyl alcohol: benzyle benzoate, mounted onto

microscope slides, and imaged with a Zeiss LSM 700 confocal microscope. A Vibratome

(Vibratome series 1000, Pelco 101) was used to generate 200µm transverse sections from

agarose-mounted whole embryos. For phalloidin, embryos were sectioned prior to staining;

for ECM antibodies, embryos were stained prior to sectioning. Sections were mounted in

Vectashield mounting media and imaged with a Zeiss LSM 700 confocal microscope.

26

CHAPTER 3: The role of the neural plate on muscle fate specification

3.1 Abstract

Embryo development consists of a series of inductive events wherein neighboring tissues

communicate with another to specify the both tissue type and embryonic axis. During

Xenopus laevis development, the Spemann’s Organizer tissue (analogous to the zebrafish

shield and amniote node) has traditionally been thought to induce the formation of dorsal

tissues including the paraxial mesoderm. The paraxial mesoderm forms segments called

somites – transient structures that eventually give rise to many adult derivatives, including

the musculoskeletal system. Overlying the paraxial mesoderm tissue is the neural plate,

which eventually folds into the neural tube and forms the adult central nervous system. By

modulating the size of the neural plate, we show a corresponding change in the size of the

paraxial mesoderm. This suggests that Spemann’s Organizer is not solely responsible for

induction of dorsal tissues, and that in fact the neural plate also induces paraxial mesoderm

formation. In particular, increasing the size of the neural plate results in more paraxial

mesoderm tissue at the expense of surrounding tissues. In addition, modulating paraxial

mesoderm size has allowed us to dissect the biomechanics of somite morphogenesis. We

show that the paraxial (pre-somitic) mesoderm must unfold dorsally and ventrally as the

embryonic axis elongates. Indeed, disruption of this unfolding by increasing paraxial

mesoderm size results in defective somites with disorganized extracellular matrix and

misaligned muscle fibers. Together, these data show a novel role for the neural plate and

uncovers an important relationship between paraxial mesoderm specification and somite

morphogenesis.

3.2 Introduction

Embryo development involves the establishment of the body axis and the differentiation of

specialized tissues and organs. This is partly done through tissue interactions wherein

tissues influence each other’s development. One example of this is during somite formation.

Somites are derived from the paraxial mesoderm and eventually differentiate into adult

tissues including muscle, bone, and dermis. Somite patterning is crucial to the differentiation

of these lineages and is accomplished through morphogens secreted from adjacent tissues

[158].

Specific instances of morphogen-mediated tissue interactions during somitogenesis include

that of the notochord secreting Sonic Hedgehog (Shh) to regulate the ratio between epaxial

and hypaxial myotome within the somite [52]. Conversely, somites themselves secrete

27

morphogens that influence the development of adjacent tissues, such as specifying the

posterior fate in neural ectoderm through Wnt signaling [173].

A reciprocal relationship between the neural tissue and the underlying somitic mesoderm

was reported in X. laevis [174-175]. Enlarging the neural plate via foxd1 overexpression (as

well as other cell-autonomous neuralizing factors) also increased the size of the underlying

paraxial mesoderm as measured by myoD expression. In this study, we decided to dissect

this phenomenon in greater detail, particularly with regards to somite morphogenesis. We

found that the effect described by [175] between the neural plate and the paraxial mesoderm

is dose-dependent. Furthermore, we confirm that this effect is non-cell autonomous and that

reducing the size of the neural plate also reduces the size of the paraxial mesoderm. We

report the source and ultimate fate of ectopic paraxial mesoderm as well as neural plate

tissue, and dissect the activation of myogenic regulatory factors. Finally, we observe the

effects of ectopic tissue on somite morphogenesis and find that increasing the size of the

paraxial mesoderm results in misaligned somites with disorganized intersomitic boundaries,

an effect that cannot be rescued by exogenous Sonic Hedgehog, a signaling molecule crucial

to myotome differentiation. Together, our data help further illuminate a fascinating

interaction between the neural ectoderm and somitic mesoderm.

3.3 Results

Expansion of the neural plate results in a corresponding increase in presomitic

mesoderm size

Expansion of the neural plate through foxd1 mRNA overexpression results in a larger

presomitic mesoderm [175]. To more thoroughly dissect this phenomenon, we injected

foxd1 mRNA into X. laevis embryos to create a dose-response curve and assay its effects on

neural crest specification. Embryos were unilaterally-injected with 50pg (Fig 1Q-1T), 100pg

(Fig 1M-1P), 250pg (Fig 1I-1J), 500pg (Fig 1E-1H) foxd1 mRNA, fixed, then assayed for sox2,

snai2, pax3, and myoD expression via in situ hybridization. Compared to uninjected control

embryos (Fig 1A-1D), embryos showed expansion of sox2 and myoD expression in the

injected side (white arrowheads in Fig1, sox2 and myoD columns), indicating expansion of

the neural plate and the underlying presomitic mesoderm. Expression of neural crest

markers snai2 and pax3 confirmed expansion of the neural plate (white arrowheads in Fig 1,

snai2, and pax3 columns), as expression is shifted laterally in favor of neural plate fate. The

change in gene expression observed in injected embryos show a dose response, with 500pg

treatments (Fig 1E-1H) significantly more affected than 50pg treatments (Fig 1Q-1T).

Observed phenotypes were quantified using a binary metric of symmetrical or asymmetrical

expression, and are summarized in bar graphs (Fig 1U-1X). Together, these data confirm a

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dose-related correlation in expanding the neural plate at the expense of neural crest and a

corresponding increase in presomitic mesoderm size.

Reducing neural plate size results in reduction of presomitic mesoderm size

To complement the experiments described above, we sought to determine if reducing the

size of the neural plate would result in a reduction of presomitic mesoderm size. To reduce

neural plate size, we injected a morpholino targeting Sox2 transcripts, then assayed injected

embryos with relevant markers via in situ hybridization. We unilaterally-injected 80ng (Fig

2G-2I), 60ng (Fig 2J-2L), 40ng (Fig 2M-2O), 20ng (Fig 2P-2R), and 10ng (Fig 2S-2U) sox2 MO

and compared expression of pax3, myoD, and sox2 with uninjected control (Fig 2A-2C) and

standard MO-injected (Fig 2D-2F) embryos. Sox 2-injected embryos showed a reduction in

neural plate size, as seen in the medial shift of pax 3 expression (white arrowheads in Fig 2,

pax 3 column). Furthermore, myoD expression in injected embryos also showed a reduction

in size as compared to the contralateral, uninjected side (white arrowheads in Fig 2, myoD

column). Phenotypes followed a dose-specific trend wherein 80ng of sox2 MO elicited a

much more severe response than did 10ng of sox2 MO. Phenotypes were classified as either

symmetrical or asymmetrical and quantified (Fig 2V-2X). Together, these data indicate that

a reduction in neural plate size results in a corresponding reduction in presomitic mesoderm

size.

Effect of neural plate size on presomitic mesoderm specification is non-cell

autonomous

The correlation in the size of the neural plate and underlying presomitic mesoderm suggests

that the neural plate somehow influences that size of the presomitic mesoderm.

Alternatively, our foxd1 mRNA and sox2 MO injections could be directly inducing the

observed phenotypes in both the neural plate and presomitic mesoderm. To determine

which scenario is occurring, we carried out a transplantation experiment wherein we

transplanted the animal caps of foxd1-injected embryos to uninjected hosts (Fig 3A). As the

embryo develops from the blastula to the gastrula stage, prospective presomitic mesoderm

underlies transplanted animal cap (labeled with RDA) and is subject to its influence. At the

neurula stage, we assayed gene expression of grafted embryos via in situ hybridization. We

found that compared to control grafts (Fig 3B and 3E), caps with 500pg (Fig 3C and 3F) and

100pg (Fig 3D and 3G) foxd1 mRNA induced ectopic sox2 expression (white arrowheads in

Fig 3B-3D) and ectopic myoD expression (white arrowheads in Fig 3E-3G). These data

confirm that the neural plate influences presomitic mesoderm size non-autonomously.

29

Ectopic presomitic mesoderm is transient and results in disorganized somites

Next, we wanted to look more closely at the ectopic presomitic mesoderm induced by an

expanded neural plate. By analyzing myoD expression in embryos unilaterally injected with

foxd1 mRNA, we found that the ectopic presomitic mesoderm results in disorganized

somites. At the neurula and early tailbud stages, myoD expression is expanded in the injected

side (black arrowheads in Fig 4A-C). However, by the late tailbud stage, myoD expression no

longer appears expanded, and is instead restricted to the same surface area as the

contralateral, uninjected side. Interestingly, myoD expression in the injected side (black

arrowheads in Fig 4E and 4G) is disorganized, and does not display the same segmented

expression pattern as the contralateral side (Fig 4D and 4F). Together, this data shows that

the expanded presomitic mesoderm observed in neural plate-expanded embryos results in

perturbed somites and disorganized myoD expression by the late tailbud stages.

Ectopic presomitic mesoderm does not undergo apoptosis

The disappearance of ectopic presomitic mesoderm in the later stages of foxd1-injected

embryos prompted us to inquire on the ectopic tissue’s fate. We hypothesized that the

ectopic tissue was undergoing apoptosis and thereby returning to a similar size as the

uninjected control. However, our TUNEL labeling reveals that the PSM in foxd1-injected side

(Fig 5G-FL) of embryos undergoes the same low level of apoptosis as the uninjected side (Fig

5A-5F) at both the early tailbud (Fig 5A-5C; 5G-5I; and 5M-5O) and tadpole (Fig 5D-5F; 5J-

5L; and 5P-5R) stages, and is confirmed by transverse sections of the same embryos (Fig 5M-

5R). Together, this data indicates that the disappearance of ectopic presomitic mesoderm in

neural plate-expanded embryos is not due to apoptosis.

Ectopic presomitic mesoderm is specified at the expense of adjacent tissues and is

later re-specified to the source tissue

We hypothesized that the source of ectopic presomitic mesoderm tissue in foxd1-injected,

neural plate-expanded embryos is either overproliferation of PSM or re-specification of

adjacent tissues. To address the latter, we subjected embryos unilaterally injected with foxd1

mRNA to in situ hybridization probing for markers known to be expressed in tissues adjacent

to the presomitic mesoderm. Compared to the contralateral side (Fig 6A-6C), the injected

side of embryos had drastically reduced or absent expression of lim1, a marker of lateral

plate mesoderm (Fig 6-G), gata3, a marker of intermediate mesoderm (Fig 6H), and gata4, a

marker of cardiac mesoderm (Fig 6I) at the early tailbud stage. This suggests that ectopic

presomitic mesoderm is specified at the expense of intermediate and lateral plate

mesoderm.

30

Next, we wanted to address the fate of ectopic presomitic mesoderm tissue. Experiments

described above indicate that the ectopic tissue does not undergo apoptosis (Fig 5). Based

on the source of ectopic presomitic mesoderm (Figs 6A-6I), we hypothesized that rather than

undergoing apoptosis, ectopic presomitic mesoderm is instead being re-specified to its

original prospective fate. Indeed, in situ hybridization at later stages (Figs 6A-6I) supports

this hypothesis. Compared to the contralateral side (Fig 6D-6F), the injected side of embryos

showed a return of lim1, gata3, and gata4 expression at the late tailbud stage, albeit in a

slightly perturbed state (black arrowheads in Fig 6J-6L). This data suggests that ectopic

presomitic mesoderm expressing myod in foxd1-injected embryos originates from laterally-

adjacent tissues, but is then re-specified to the source tissue by the late tailbud stage.

Ectopic neural tissue is transient and is dorsal and posterior in identity

We also wanted to examine expanded neural plate in foxd1-injected embryos. First, we

probed unilaterally-injected embryos for three well-known neural markers: otx2, krox20,

and hoxb9 for forebrain, midbrain, and hindbrain, respectively. At the late neurula stage, our

data reveal an anterior shift in both otx2 and krox20 expression (black arrowheads in Fig

7A), and an extensive lateral expansion of hoxb9 (black arrowheads in Fig 7B-7C). By the late

tailbud and tadpole stages, however, the expansion of gene expression returns to near-

normal (black arrowheads in Fig7D-7I). This pattern mirrors that of the expression

dynamics for mesoderm markers used above, wherein the tissue is expanded until the early

tailbud stages, but then returns to near its original state by the late tailbud stage.

A similar expression pattern was observed in differentiated neural tissue, using n-tubulin as

an in situ hybridization probe. Early tailbud stage embryos showed a lateral expansion of n-

tubulin expression, visible from the dorsal view (black arrowheads in Fig 7J-7K), as well as

in transverse sections (black arrowhead in Fig 7L). However, by the late tailbud stage, the

expanded tissue begins to shrink (black arrowheads in Fig 7M-7O), and by the tadpole stage

has returned to almost normal (black arrowheads in Fig 7P-7R). Interestingly, although the

morphogenesis of the underlying mesoderm is perturbed, neural tube closure seems to

proceed relatively normally.

Expression of early and late myogenic regulatory factors is perturbed

Myogenic regulatory factors consist of four transcription factors expressed sequentially to

regulate muscle specification and differentiation [158]. For instance, myoD and myf5 are

expressed early, while mrf4 and myogenin are more active in the differentiation step of

myogenesis. Based on the expansion of myod expression in foxd1-injected embryos, and the

subsequent defects in somite morphogenesis, we suspected that not all of the myogenic

31

regulatory factors were being activated. Indeed, we found differences in the expression of

early compared to late myogenic regulatory factors. At the late gastrula stage, both myoD and

myf5 expression are expanded (black arrowheads in Fig 8B and Fig 8L) compared to the

uninjected control sides and to uninjected control embryos (Fig 8A and Fig 8K; quantified in

Fig 8BB). At the late neurula stage, myoD and myf5 expression is expanded (black

arrowheads in Fig 8D and Fig 8N), but mrf4 expression is reduced and disorganized (red

arrowheads in Fig 8P) compared to control embryos (Fig 8C, Fig 8M, and Fig 8O; quantified

in Fig 8CC). Expression of myogenin is not yet apparent in uninjected embryos at this stage

(Fig 8U). By the early tailbud stage, myod expression remains expanded (Fig 8F) compared

to the control side (Fig 8E), while mrf4 expression is disorganized (red arrowheads in Fig

8R) and does not form repeated segments like those found in the control side (Fig 8Q).

Expression of myogenin at this stage appears mildly disrupted in the injected side (Fig 8V)

compared to the uninjected side (Fig 8W; quantified in Fig 8DD). At the late tailbud and

tadpole stages, myoD expression becomes more similar in size between injected (Fig8H and

Fig 8J) and uninjected (Fig 8G and Fig 8I) sides. However, expression on the injected side is

perturbed and disorganized (black arrowheads in Fig 8G and Fig 8I), showing none of the

repeated segmental pattern of the uninjected side. Both mrf4 (red arrowheads in Fig 8T) and

myogenin (red arrowheads in Fig 8Y and Fig 8AA) expression are also disrupted at this stage,

compared to their uninjected control sides (Fig 8Q, Fig 8X, and Fig 8Z; quantified in Fig 8FF

and Fig 8GG). A summary of the different expression pattern disruptions by transcription

pattern and developmental stage can be found in (Figure 8HH). Together, this data indicates

that ectopic presomitic mesoderm induced by foxd1-mediated neural plate expansion

activates early but not late myogenic regulatory factors, thus resulting in an expanded

presomitic mesoderm tissue but ultimately disorganized somites.

Ectopic presomitic mesoderm results in abnormal somite morphogenesis

Failure to inactivate late myogenic regulatory factors led us to hypothesize that somite

morphogenesis must be unable to proceed to its final stages. Indeed, transverse sections of

embryos unilaterally-injected with foxd1 mRNA and fixed at various stages show that

presomitic mesoderm unfolding does not occur even by the early tailbud stage (Fig 9A-D).

Even by the late tailbud stage, the somite is not properly unfolded (Fig 9E-F). Interestingly,

embryos retain myoD expression throughout this process, suggesting that late myogenic

regulatory factors are needed for somite tissue morphogenesis.

Ectopic presomitic mesoderm results in disrupted myotome alignment

The disrupted expression of myogenic regulatory factors at the late tailbud stage prompted

us to investigate somite morphology via confocal microscopy. We unilaterally injected foxd1

32

mRNA along with RDA as a lineage tracer at the two-cell stage, then allowed injected

embryos to grow to the late tailbud stage. Using the 12101 antibody, we then visualized

somite morphology and myotome alignment and quantified phenotypes using a scoring

rubric we generated (Supp Fig 1). Uninjected control embryos at stages 21, 25, 28, and 38

(Fig 10A-D, respectively) were scored as a “5” on the rubric (Fig 10M). At St. 21 and 25,

embryos injected with 50pg or 100pg foxd1 mRNA had relatively mild phenotypes (Fig 10N

– 10O). However, by the late tailbud and tadpole stages, embryos injected with 50pg of foxd1

mRNA scored much lower at a “3” and “1” (Fig 10N). Somites had many morphological

defects, including non-elongated myotome, lack of intersomitic boundaries, and misaligned

myotome (Fig 10E – 10H). Phenotypes of embryos injected with 100pg foxd1 mRNA scored

much lower in both the early (Fig 10O) and late (Fig 10I-10L and Fig 10O) stages. At the late

tailbud and tadpole stages, embryos had non-elongated myotome, lack of intersomitic

boundaries, misaligned myotome, and lacked a compact overall somite morphology (Fig 10I-

10L). Together, these data show that a consequence of not activating the full suite of

myogenic regulatory factors observed in (Fig 8) is a severe disruption in somite morphology.

Disrupted somites have abnormal extracellular matrix protein deposition

Somite morphogenesis requires carefully-orchestrated cell behaviors that require cells to

anchor to extracellular matrix proteins deposited in the intersomitic boundaries [17, 158].

The disruption in somite morphology observed in (Fig 10) included disorganization of

myotome and lack of intersomitic boundaries. Therefore, we sought to examine if foxd1-

injected embryos exhibit abnormal somite morphology as a result of – or resulting in –

disorganization of extracellular matrix deposition. Uninjected embryos were stained for

laminin, β-integrin, fibronectin, and β-dystroglycan, each of which are required for myotome

rotation and normal somite morphogenesis [158]. At stage 28 (Fig 11A-11D) and stage 38

(Fig 11E-11H), each of these ECM proteins are expressed primarily in the intersomitic

boundaries in regular, organized segments. However, in embryos unilaterally-injected with

foxd1 mRNA, ECM protein deposition was disorganized at both stage 28 ( white arrowheads

in Fig 11I-11L) and stage 38 (white arrowheads in Fig 11M-11P), exhibiting a range of

phenotypes from complete absence to asymmetrical outlining (quantified in Fig 11Q-11T;

scoring rubric in supp Fig 2). What remains unclear is whether abnormal ECM deposition is

a cause or effect of abnormal somite morphogenesis in foxd1-injected embryos.

Sonic Hedgehog does not rescue MRF expression or somite morphogenesis

Sonic Hedgehog (Shh) regulates hypaxial myotome differentiation in X. laevis [52]. Shh

signaling during somitogenesis originates from the midline, primarily the notochord [158].

Given that foxd1-injected embryos have disrupted somitogenesis stemming from a lateral

33

expansion of presomitic mesoderm, we wondered if the defects in somite differentiation are

due to the expanded lateral PSM receiving inadequate Shh signaling from the midline. To

address this, we unilaterally-injected Shh mRNA and foxd1 mRNA into 2-cell stage embryos

to see if somite morphogenesis would be rescued. At the neurula stage, uninjected control

embryos have symmetrical expression of pax3, myoD, and myf5 (Fig 12A-C). Injection of

100ng Shh and 50pg foxd1 in one of two cells still results in perturbed pax3 expression and

expanded myoD and myf5 expression (Fig 12D-F). Notably, injection of Shh alone does not

perturb expression of pax3, myoD, or myf5 at the neurula stage (Fig 12G-I). Phenotypes were

quantified and summarized in (Fig 12J-L).

Next, we examined gene expression patterns in St. 30 (Fig 12M-X) and St. 40 (Fig 12Y-JJ)

embryos. At St. 30, hypaxial myotome pax3 expression in shh-injected embryos is missing

(Fig 12V-X) as expected [52], compared to the uninjected control side (Fig 12S-U). However,

in shh + foxd1 mRNA-injected embryos, expression of myoD and mrf4 are still disrupted

(Fig12P-R) compared to the uninjected control side (Fig 12M-O). Similar phenotypes were

observed in St. 40 embryos (Fig 12Y-JJ), and phenotypes were quantified and summarized in

(Fig 12KK-PP).

Lastly, we used the 12101 antibody to visualize somite morphogenesis in foxd1 + shh-

injected embryos at St. 30 (Fig 12QQ-SS) and St. 40 (FigTT-VV). At both stages, foxd1+shh-

injected embryos still exhibit disrupted somite morphogenesis characterized by myotome

misalignment and disrupted intersomitic boundaries (Fig 12QQ and 12TT) compared to

uninjected control embryos (Fig 12SS and 12VV). As expected, embryos injected with shh

alone also exhibited minor morphogenetic defects attributed to disrupted hypaxial myotome

differentiation (Fig12RR and 12UU). Phenotypes were scored and quantified in (Fig 12WW-

YY). Together, this data shows that shh signaling does not rescue the disrupted somites in

foxd1-injected embryos.

3.4 Discussion

Neural plate size and presomitic mesoderm size are correlated

In this report, we describe the interesting phenomenon of a reciprocal relationship between

the neural plate and the paraxial mesoderm. It was previously shown that enlarging the

neural plate through overexpression of foxd1 mRNA caused myoD expression to expand

[175]. Here, we expanded on this observation by examining the source and fate of ectopic

paraxial mesoderm tissue, as well as by dissecting the expression patterns of the myogenic

regulatory factors involved in myotome differentiation. Overexpression of foxd1 mRNA

results in a dose-related increase in the expression of myoD and myf5, both early-activated

34

myogenic regulatory factors (Fig 1). We also found that reducing the size of the neural plate

via sox2 morpholino has a corresponding, dose-related effect in decreasing the size of the

paraxial mesoderm as measured by myoD expression (Fig 2). Tissue transplants

demonstrated that this effect is non-cell autonomous, and that ectopic paraxial mesoderm

can be induced by neutralized animal cap (Fig 3). Interestingly, ectopic paraxial mesoderm

results in somites with perturbed myoD expression (Fig 4), thus prompting us to inquire into

the fate of the ectopic tissue. Rather than undergoing apoptosis (Fig 5), we found that ectopic

paraxial mesoderm, which expanded laterally, was instead re-differentiating into the source

tissue – namely lateral plate mesoderm (Fig 6). Examining the neural plate over time also

yielded interesting results. Foxd1 expanded neural tissue that is dorsal and posterior in

origin, but interestingly, its lateral expansion is also transient (Fig 7). The disruption of myoD

expression in the somites of foxd1-injected embryos led us to examine the expression

patterns of the other myogenic regulatory factors. We found that myoD and myf5 – both early

myogenic regulatory factors – are expanded, whereas mrf4 and myogenin are instead

reduced and/or disrupted (Fig 8). This suggests that foxd1-mediated neural plate expansion

is causing increased presomitic mesoderm specification, yet is unable to activate a similar

increase in mrf4 and myogenin expression, thus preventing the ectopic tissue from fully

differentiating into myotome. Indeed, examining transverse sections of foxd1-injected

embryos reveals a phenotype in which the ectopic tissue indeed expresses myoD, but

remains in a folded configuration reminiscent of immature presomitic mesoderm, even

though the uninjected control side has already unfolded and formed a mature somite (Fig 9).

Foxd1 activity in the neural plate and paraxial mesoderm

Another possibility is that foxd1 is acting in both the neural plate and paraxial mesoderm

[176]. Here we present a possible model in the signaling dynamics between the neural

ectoderm, axial mesoderm, and paraxial mesoderm (Fig 13). In the axial mesoderm, Noggin

and other Bmp antagonists inhibit Bmp4 expression in both the neural ectoderm and

paraxial mesoderm, thus allowing for the expression of foxd1 in both tissues. In the paraxial

mesoderm, foxd1 acts to repress expression of lateral plate mesoderm markers such as lim1

and gata 3 and 4. In the neural ectoderm, foxd1 expression is dependent on both bmp4 and

wnt inhibition; wnt inhibition is carried out by dkk from the axial mesoderm. Neural plate

foxd1 then triggers the secretion of a factor that induces myoD and myf5 expression in the

underlying paraxial mesoderm. Reciprocally, the paraxial mesoderm and somite then

expresses wnt, which specifies a posterior fate in the overlying neural ectoderm. However,

whatever neural plate factor that is inducing myoD and myf5 expression is not sufficient to

activate mrf4 and myogenin expression, thus preventing full differentiation of any foxd1-

induced ectopic tissue to fully differentiate into myotome, resulting in disrupted somite

morphogenesis.

35

Ectopic presomitic mesoderm tissue disrupts somite morphogenesis

An interesting outcome of inducing ectopic paraxial mesoderm tissue is the resulting

disruption of somite morphogenesis. As discussed in the previous chapter, somite

morphogenesis consists of a series of complex morphogenetic movements at both the tissue

and cell level. The presomitic mesoderm (PSM) has been shown to unfold dorsally and

ventrally as it matures into a somite, effectively placing the lateral portion of the tissue to the

dorsal and ventral domains of the somite. Remarkably, this unfolding occurs autonomously,

and does not require the influence of adjacent tissues, nor is regulated by either the canonical

Wnt or noncanonical Wnt/PCP pathways. In foxd1-injected embryos, the PSM expands

laterally as early as the gastrula stage, but fails to unfold by the early tailbud stage even

though the uninjected control side has already unfolded. This seems to be a result of delayed

morphogenesis, since by the late tailbud stage, the presomitic mesoderm shows signs of

partially unfolding (Fig 9). Normally, PSM unfolding is followed by rearrangements of

clusters of cells, then ultimately rotation of individual myotome to organized them into

regular arrays parallel to the notochord. The disruption of somite morphogenesis in foxd1-

injected embryos suggests that these cell rearrangements and rotations are also perturbed,

possibly due to the tissue being unable to properly coordinate the movements of its ectopic

portions. Nevertheless, the addition of ectopic tissue raises interesting questions into its

effect on somite morphogenesis.

36

3.5 Methods

Embryo Culture and Microinjections

X. laevis females were injected with 0.5ml human chorionic gonadotropin (HCG) hormone to

induce ovulation. Eggs were collected 15 to 20 hours post injection and fertilized via in vitro

fertilization with X. laevis male sperm in 1/3X MR at 12°C or 16°C. Fertilized embryos were

then dejellied using 3% cysteine solution, then cultured in 1/3x MR. Donor embryos for

transplantation experiments were injected with a Picospitzer II (General Valve Corporation)

and microinjection manipulator (Narishige Group) at the 1-cell stage with 9.2nl of 70,000

MW rhodamine dextran amine (RDA) in 2.5% Ficoll. Injected embryos were then transferred

to 1/3x MR and placed in 12°C. Constructs used were XBF-2/Foxd1 [174] and Shh [52].

Animal cap transplants

RDA-labeled donor and unlabeled host embryos were allowed to develop to gastrula stage.

The protective vitelline envelope was removed from both host and donor embryos using

forceps. Eyebrow and eyelash tools were used to remove the animal cap from donor and host

embryos. Transplantations were performed in 2% agar coated Petri dishes containing 1×

MBS and then transferred to 2% agar coated Petri dishes containing 1/3× MBS for long-term

culture. At neurula stage, embryos were fixed in 1× MEMFA (100mM MOPS pH 7.4; 2mM

EGTA, 1mM MgSO4; 3.7% formaldehyde) for one hour at room temperature or overnight at

4°C. Fixed embryos were transferred to 100% methanol for long-term storage.

In situ hybridization and Immunohistochemistry

In situ hybridization was performed as previously described [163]. To visualize the

notochord and mature somites, whole embryos were stained with 1:250 12/101 antibody

(Developmental Studies Hybridoma Bank) and 1:5 Tor70 antibody (Harland Group),

respectively. For secondary antibodies, 1:250 Alexa Fluor 488 goat anti-mouse IgG and 1:250

Alexa Fluor 647 goat anti-mouse IgM were used.

To visualize extracellular matrix depositions, the following antibodies from the

Developmental Studies Hybridoma Bank were used on whole embryos: 1:50 8C8 (β-

integrin), 1:100 7DII (β-dystroglycan), and 1:100 MT4 (fibronectin). For secondary

antibodies, 1:250 Alexa Fluor 488 goat anti-mouse IgG or 1:250 Alexa Fluor 555 goat anti-

mouse IgG were used. 1:100 laminin antibody (Sigma Aldrich) with 1:250 Alexa Fluor 555

goat anti-rabbit IgM secondary antibody was also used. Antibodies were diluted with 1:10

cas-block [Life Technologies]: PBSt.

37

Imaging

Whole embryos were cleared with 2:1 benzyl alcohol: benzyl benzoate, mounted onto

microscope slides, and imaged with a Zeiss LSM 700 confocal microscope. A Vibratome

(Vibratome series 1000, Pelco 101) was used to generate 200µm transverse sections from

agarose-mounted whole embryos.

38

Future Perspectives

X. laevis provides an excellent context for dissecting the mechanisms of mesoderm

induction, morphogenesis and differentiation. The unique attributes of X. laevis development

allows the study of primary and adult myogenesis separately, and gives tantalizing insights

into the evolution of the tetrapod limb and trunk. However, many questions still remain

about the molecular and morphological mechanisms of myogenesis. Emerging studies on

gene regulation via alternative splicing have opened up many avenues of research to

challenge previously established roles of molecules during development [43]. At the genetic

level, there remains the problem of precisely identifying the specific gene targets of each

MRF in an effort to untangle which of their functions are redundant and which are not.

Although genetic studies have been historically difficult due to the pseudotetraploidy of X.

laevis, the use of dominant-negative proteins and morpholino technology have elucidated

the genetic bases of many developmental processes. Furthermore, the establishment of X.

tropicalis as a more genetically tractable counterpart to X. laevis has renewed the use of

molecular biology in this amphibian system [151,152]. In fact, comparative studies are

underway between the two species, particularly with regards to mesoderm patterning and

morphogenesis [153]. In addition, the availability of the X. tropicalis genome has facilitated

numerous genetic studies in this organism [154]. Indeed, the emergence of more efficient

gene-editing technologies, particularly the CRISPR/Cas9 system, has allowed for genetic

studies in both X. tropicalis [155] and X. laevis [156,157]. These, along with the many merits

of X. laevis as a model system, place this organism at the forefront of developmental biology

research, particularly in mesoderm patterning and differentiation.

39

40

Figure 1. A schematic representation of somite morphogenesis in X. laevis. A dorsal view of a step-wise process of somite morphogenesis that begins in the PSM (posterior end) and ends with the formation of aligned myotome fibers (anterior end). The process begins with cells in the PSM that become progressively more elongated in the mediolateral direction as they near the transition zone where somite formation will begin (Step 1). Somite formation begins with the formation of the intersomitic boundary which first appears from the lateral edge and moves medially toward the notochord. This early intersomitic boundary is comprised of fibronectin (red arrow). As this intersomitic boundary begins to form at the lateral edge of the paraxial mesoderm, somite cells increase their filopodial activity. These cells also express integrin β1 (Step 2). This is followed by a 90o rotation such that each elongated cell individually bends around the anterior to posterior axis (Step 3). At the same time, these cells begin to express β-dystroglycan, which plays an important role in crosslinking laminin (blue) to the intersomitic ECM. The last step involves the stable attachment of myotome fibers to the intersomitic boundary, thereby establishing an elongated and aligned morphology (Step 4). Adapted from [17] and [21].

41

42

Figure 2. Muscle fate maps of the gastrula and tadpole. Cells positioned in the upper lateral lip region (A; light green) will undergo a significant amount of convergence and extension to give rise to both the anterior-most somites as well as myotome fibers positioned in the central region of somites along the entire anterior to posterior axis. Cells positioned in the lateral circumblastoporal region (B; red) will intercalate with cells from region A to give rise to myotome fibers found throughout the dorsal and ventral extent of the trunk somites. Cells positioned in the lower lip region (C; blue) are found in the dorsal and ventral-most domain of somites located in the trunk and tail regions of the tadpole. Scanning electron micrographs of cross sections of embryos from stages 13 to 24 reveal the distribution of mesoderm cells from gastrula regions A, B, and C. Cells from region A are positioned medially adjacent to the notochord, while cells from region B will lie lateral to region A although a subset of B cells will intercalate among A cells and end up residing more medially. Cells from region C will lie lateral to region B (stage 13). As the embryo matures there is considerable dorsal convergence and expansion of the two-layered PSM (stages 16 to 24). These movements displace cells from the lateral region to the dorsal and ventral aspects of the somite. Cells from region B will undergo a significant amount of convergence and will mix with cells from regions A and C of the gastrula. Cells from region C of the gastrula will lie at the lateral edge of the PSM and will eventually contribute to the formation of the most dorsal and ventral cells of the somite. Around stage 19 we begin to see a thin epithelial layer consisting of cuboidal cells that lie on the surface of the paraxial mesoderm. This tissue is likely the dermomyotome (shown in white). Anterior is to the left for images of the tadpole musculature. Adapted from [27].

43

Figure 3. Signals that pattern the vertebrate somite. A simplified diagram of a cross-sectioned amniote (I) and X. laevis (II) embryo illustrating proteins secreted by adjacent tissues that pattern the somite. In the amniote somite, Wnts from the dorsal neural tube (tan) and the epidermal ectoderm (black), along with Bmp4 from the lateral plate (violet) maintain the dermomyotome (dark red) in an undifferentiated state. Hedgehog from the notochord and neural tube floor plate (dark blue) specify the sclerotome (light blue). Once the sclerotome segregates, the prospective epaxial (orange) and hypaxial (yellow) myotome differentiate from the dermomyotome. In X. laevis, Wnt1 from the dorsal neural tube (tan) and Wnt11 from the somite instruct myotome (green) proliferation and differentiation. Medium levels of Bmp4 from the lateral plate (violet) specify the dermomyotome (dark red) and muscle satellite cells. Hedgehog from the notochord and neural tube floorplate (dark blue) induces sclerotome (light blue) and slow-twitch myotome (green) in the tail. Hedgehog, along with Wnt7a from the epidermal ectoderm, regulates the differentiation of epaxial (orange) and hypaxial (yellow) myotome from the dermomyotome. Adapted from [30].

44

Figure 4. Expression patterns of Myogenic regulatory factors (MRFs) throughout the three myogenic waves. During the first wave, myoD and myf5 are expressed in the PSM. As somitogenesis progresses, myoD and myf5 become restricted to the caudal PSM and early somites, while mrf4 and myogenin become expressed in differentiating myotome. During the second wave, all MRFs are expressed in the epaxial (dorsal) region of the dermomyotome. By the third wave, myf5 is expressed as puncta in the anterior and posterior region of each somite, whereas mrf4 is expressed in the center of each myotome and myogenin is most highly expressed in the hypaxial myotome. Gastrulae are viewed from the vegetal perspective; neurulae are viewed from the dorsal perspective with anterior to the left; tadpoles are from the lateral perspective with anterior to the left. Dark green indicates high levels of expression, yellow indicates lower levels. Adapted from [31].

45

Fig 1: Pre-somitic cells from different regions of the gastrula take distinct routes to reach the

dorsal part of the embryo. Fluorescently-labeled cells grafted to the ULL region (A-D) and LL region

(E-H) of the gastrula involute into the blastopore during gastrulation and form somites as they reach

the dorsal part of the embryo. A-D: Cells in the ULL region involute adjacent to the developing

notochord and remain in the central domain of somites by the early tailbud stage. A: Schematic showing

graft location at the gastrula stage. B-D: Lateral (B), transverse (C), and dorsal (D) views of the same

embryo showing the location of grafted cells (red) adjacent to the notochord (blue) as they form somites

(green). Grafted cells in (B) and (D) are undergoing cell rotation as part of normal X. laevis

somitogenesis. E-H: Cells in the LL region involute opposite the prospective notochord and form the

lateral edge of the pre-somitic mesoderm. E: Schematic showing graft location at gastrula stage. F-H:

Lateral (F), transverse (G), and dorsal (H) views of the same embryo showing the location of grafted

cells (red) migrating from the blastopore (yellow asterisk) to the lateral edge of the pre-somitic

mesoderm. Autofluorescence was used to show the rest of the embryo. 12101: myotome, Tor70:

notochord, RDA (rhodamine dextran amine): grafted cells.

46

Fig 2: Pre-somitic cells form myotome in specific domains along the anterior-posterior and dorsal-

ventral axes. Fluorescently-labeled cells grafted to the ULL region (A-E) and the LL region (F-L) of the gastrula

form myotome in specific locations within the somite and along the anterior-posterior axis by the late tailbud

and tadpole stages. A-E: Cells in the ULL region remain adjacent to the notochord and form myotome across the

entire anteroposterior axis in the central domain of somites. E: Schematic showing graft location in the gastrula

and stage at which the grafted embryos were fixed. A-D: Lateral views of St. 28 (A) and St. 40 (B) grafted embryos

showing transplanted ULL cells (red) along the anterior-posterior axis in the central domain of somites. (B) and

(D) are transverse sections of (A) and (C), respectively, showing the location of transplanted ULL cells (red)

adjacent to the notochord (blue dashed line) in the central domain of the somite. F-L: Cells from the LL region

form myotome in the dorsal and ventral domains of posterior trunk and tail somites. F: Schematic showing graft

location in the gastrula and stage at which the grafted embryos were fixed. G-L: Lateral views of St. 28 (H) and

St. 40 (J) grafted embryos showing transplanted LL cells (red) in the dorsal and ventral domains of posterior

trunk and tail somites. (I) and (K) are transverse sections of (H) and (J), respectively. (L) is a magnified view of

(K) showing grafted cells in the ventral domain. (G) is a magnified view of the posterior region of (H) showing

grafted cells (red) splitting off into the dorsal and ventral domains of the developing somite. 12101: myotome,

Tor70: notochord, RDA (rhodamine dextran amine): grafted cells.

47

48

Fig 3: The pre-somitic mesoderm unfolds dorsally and ventrally as it matures into somites. Transverse

sections from the posterior region reveal the dynamic morphology of the unfolding pre-somitic mesoderm (PSM)

over time. A-C: Transverse sections of St. 17 embryos at the onset of neurulation. Blue arrowheads point to the

neural plate closing into the neural tube, and the blue dashed line is the notochord. White arrowheads in (B-D)

point to clefts that will open up and separate the dorsal and ventral thirds of the tissue at it unfolds. D-F:

Transverse sections of St. 23-25 embryos showing the dorsal and ventral thirds of the PSM continuing to unfold.

G-I: Transverse sections of St. 28-35 embryos showing unfolded somites. White arrowheads in (G) and (H) point

to actin concentrations at St. 28, which disappear by St. 35 (I). White dashed circle is the neural tube and the blue

dashed circle is the notochord. J: Schematic illustrating PSM unfolding; (A’-I’) are roughly equivalent to (A-I).

49

Fig 4: Tissue unfolding positions cells from the lateral edge to the dorsal and ventral domains of the

somite. A-F: Fluorescently-labeled cells were transplanted to the LL region of the gastrula; grafted embryos

were cultured to early (A-B) and late (C-F) tailbud stages and sectioned to determine the position of labeled

cells throughout somitogenesis. A: RDA-labeled cells are located in the lateral edge of the pre-somitic mesoderm

(PSM). B: As the PSM unfolds, labeled cells originally in the lateral edge become separated as they moved dorsally

or ventrally. C-D: As the PSM finishes unfolding and the myotome rotates, the labeled cells become positioned

to the dorsal and ventral domains of the somite. White arrowheads in (C) and (D) indicate concentrations of

actin marking the positions at which the PSM tissue unfolds. E-F: A fully-unfolded and mature somite, with

labeled cells located in the dorsal and ventral domains. Dashed blue circle : notochord, Ph: phalloidin, RDA:

rhodamine dextran amine.

50

Fig 5: Cells undergo dramatic rearrangements after PSM unfolding. Transverse sections of the

posterior regions of embryos show the progression of dynamic cell movements during and after PSM

unfolding. From a dual layer of uniformly-distributed cells (depicted in M-I), PSM cells elongate and bend,

localizing their nuclei to the periphery as the tissue unfolds (A-C, depicted in M-II and M-III). Once the

PSM unfolds, elongated cells become positioned perpendicular to the notochord in the dorsal-ventral

plane (D-F, depicted in M-IV). Clusters of cells become organized into three groups with actin

concentrations delineating the boundary between each cluster (white arrowheads in D). In this

conformation, nuclei become organized into distinct bands (white arrowheads in (E). Afterwards, all

three clusters rotate to become aligned parallel to the notochord (G-I); white arrowheads in (G) point to

localized actin concentrations, and white arrowheads in (H) point to the bands of nuclei. In a fully mature

somite (J-L, depicted in M-V), actin concentrations are no longer present (J), and the nuclei become

uniformly distributed throughout the tissue (K). Schematic in (M) summarizes the cell movements and

nuclei localizations during and after PSM unfolding.

51

Fig 6: Cell rearrangements following somite unfolding

are accompanied by transient depositions of ECM

proteins in key locations. Transverse sections of the

posterior regions of St. 25-28 embryos show the

localization of ECM depositions composed of Fibronectin

(A-J), Integrin (P-Q), and Laminin (S-T) during PSM

unfolding. ECM hinges are present in unfolding PSM (white

arrowheads in H-I, Q, and T), but are absent in folded PSM

(F-G) and fully-mature somites (J). Also shown are the cell

rearrangements during PSM unfolding, demonstrated by

dynamic localization of nuclei over time (K-O), culminating

in characteristic bands, as indicated by white arrowheads

in (M-N, R, and U).

52

53

Fig 7: PSM unfolding and cell rearrangements occur autonomously. Tissue removal experiments demonstrate

that PSM unfolding and cell rearrangements occur autonomously, independent of adjacent tissues. A-H: UV-

irradiation experiments remove the notochord without affecting PSM unfolding, cell rearrangements, and somite

differentiation. A: Schematic for UV-irradiation experiment. E: Brightfield image of representative embryos with

control embryo on top. B-D: PSM unfolding still occurs in UV-irradiated embryos, with PSM tissue fusing where the

notochord would be; white arrowhead points to the future neural tube and asterisks mark the dorsal and ventral

regions of the folded PSM. F-H: PSM cell rearrangements in UV-irradiated embryos are still accompanied by

Fibronectin anchor points (white arrowheads), which disappear by the time the somite is unfolded (H). I-P: Dorsal

tissue removal experiments show that PSM unfolding occurs even in the absence of the notochord and neural tube.

I: Schematic for dorsal tissue-removal experiment. M: Brightfield image of representative embryos with control

embryo on top. J-L: PSM unfolding still occurs in dorsal tissue-less embryos; asterisks mark the dorsal and ventral

regions of the folded PSM. N-P: PSM cell rearrangements in dorsal tissue-less embryos is still accompanied by

Fibronectin anchor points (white arrowheads), which disappear by the time the PSM is unfolded in (P). Q-X: Dorsal

explants remove the influence of ventral tissue without affecting PSM unfolding, cell rearrangements, and somite

differentiation. Q: Schematic for dorsal explants. U: Brightfield image of representative embryos with control

embryo on top, dorsal explants in the middle, and ventral tissue pieces at the bottom. R-T: PSM unfolding still occurs

in dorsal explants; asterisks mark the dorsal and ventral regions of the folded PSM. V-X: PSM cell rearrangements in

dorsal explants is still accompanied by Fibronectin anchor points (white arrowheads), which disappear by the time

the PSM is unfolded (X). White dashed circle is the neural tube and blue dashed circle is the notochord.

54

Fig 8: Somite unfolding and cell rearrangements are not mediated by non-canonical Wnt/PCP pathway.

8-cell stage embryos were injected in both dorsal vegetal blastomeres to target prospective presomitic

mesoderm with either Xdd1 (A-H) or Xdsh-D2 (I-P). Injected embryos exhibited a short and stout phenotype,

with a slight dorsal flexure for both Xdd1 (E) and Xdsh-D2 (M). Cross sections reveal that somite unfolding still

occurs in both Xdd1 (B-D) and Xdsh-D2 (J-L) injected embryos, as demonstrated in rotated, mature myotome

shown in (C) and (K); RDA (D) and (L) was used as a lineage tracer for both constructs. In addition, fibronectin

hinges were present in Xdd1 (F-H) and Xdsh-D2 (N-P) embryos, indicated by white arrowheads in (G) for Xdd1

and (O) for Xdsh-D2. Characteristic bands of nuclei were also observed in Xdd1 (H) and Xdsh-D2 (P) injected

embryos, indicated by white arrowheads. Blue dashed circles in (B-D, F-H, J-L, and N-P) outline the notochord,

while the white dashed circles outline the neural tube.

55

56

Fig 9: Model for myotome positioning through autonomous somite unfolding and cell

rearrangements. A: At the onset of gastrulation, marginal zone cells involute into the blastopore and form

the presomitic mesoderm; ULL cells (green) form the medial edge of the PSM and LL cells (red) form the

lateral edge (Fig 1, depicted in Fig 9B-I). This involution continues as the axis elongates, and is depicted

here in the posterior region of a late tailbud stage embryo. B: Upon reaching the dorsal region of the embryo,

PSM tissue begins to unfold (Fig 3, depicted in Fig 9B-II and 9B-III). PSM unfolding is followed by dynamic

cell rearrangements and transient depositions of ECM hinges (dark blue bands) that disappear once the

PSM is fully unfolded and rotated (Figs 4-6, depicted in Fig 9B-IV). PSM unfolding effectively positions LL

cells (red) from the lateral edge of the PSM and to the dorsal and ventral domains of the somite to form

myotome after rotation (Figs 2-5, depicted in Fig 9B-V). PSM unfolding and cell rearrangements are

autonomous, and do not require adjacent tissues (Fig 7), nor is mediated by canonical Wnt or noncanonical

Wnt/PCP pathways (Fig 8).

57

Fig 10: Marginal zone cells in the LAT region undergo behaviors similar to cells in both the ULL

and LL regions. Fluorescently-labeled cells grafted to the LAT region (A-C) of the gastrula involute into

the blastopore during gastrulation and form somites as they reach the dorsal part of the embryo. Cells

from the LAT region populate both the center and lateral edgeof the PSM (B). LAT cells form myotome

throughout the dorsal-ventral axis of somites (C).

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Fig 11: β-dystroglycan also contributes to extracellular matrix hinges. Transverse sections of the

posterior regions of St. 28 embryos show the localization of β-dystroglycan (white arrowheads in A

and B) to the ECM anchor points described in (Fig 6).

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Fig 12: Canonical Wnt and noncanonical Wnt/PCP mutants exhibit secondary defects, but somites still

unfold. A-D: 8-cell stage embryos were injected with Xdd1 mRNA in one dorsal vegetal blastomere, cultured to

st. 30, sectioned, then imaged. Some embryos had neural tubes that failed to close, and somite differentiation

was negatively affected on the injected side (white arrowheads in A-D). However, the somite still unfolds and

myotome nuclei remain organized similarly to the uninjected side (Injected side denoted by white arrowheads

in A-D; nuclei organization apparent in white arrowhead in D). E-G: 8-cell stage embryos were injected with

Xdsh-D2 mRNA in both dorsal blastomeres, cultured to st. 30, sectioned, then imaged. Some embryos had neural

tubes that failed to close, but somites till unfolded normally with aligned and rotated mytome (F).

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