ABSTRACT WOMBLE, MANDY ASHTON. Left-Right Asymmetrical

ABSTRACT

WOMBLE, MANDY ASHTON. Left-Right Asymmetrical Liver Development. (Under the direction of Dr. Nanette Nascone-Yoder).

The liver is one of the many internal organs of the body that is left-right asymmetric in both anatomical location and morphology. In humans, the liver is located on the right side of the body cavity. Additionally, the right side of the liver is larger and consists of more lobes than the left. As many as 1 in 10,000 humans are born with defects in left-right asymmetry that often involve severe anomalies in liver laterality including aberrant lobation, abnormal organ position, and hepatic duct and biliary tree malformations, yet the mechanisms underlying the development of left-right asymmetries in the liver are unknown. In other asymmetric organs like the heart, lungs, stomach, and gastrointestinal tract, left- sided expression of Pitx2c, a homeobox transcription factor, is required for left-right asymmetric organogenesis. In the stomach and gastrointestinal tract of the mouse, chick, and frog, Pitx2c influences cell shape and cell rearrangements. However, it is unknown whether Pitx2c is required for asymmetrical liver development. Using the experimentally amenable embryo of the frog, Xenopus laevis, we show that Pitx2c is necessary and sufficient for left- right asymmetries in cell shape and epithelial character in the liver diverticulum. The right side cells of the liver diverticulum become apically constricted and elongated on the apical to basal axis, as the epithelium expands in surface area and ultimately forms a larger, elongated liver lobe. Pitx2c prevents left side cells from changing shape, inhibiting diverticulum expansion, ultimately resulting in a smaller, rounded liver lobe. This study is one of the first to demonstrate cellular level differences in liver morphogenesis and the first to link these differences to Pitx2c. Interestingly, the cell behaviors we observed downstream of Pitx2c in the liver are different behaviors than those observed in other organ systems demonstrating that Pitx2c’s influence on cell behavior and left-right asymmetrical morphogenesis is organ independent and may depend on the geometric constraints of the tissue as well as other factors intrinsic to that organ.

by Mandy Ashton Womble

A dissertation submitted to the Graduate Faculty of North Carolina State University in partial fulfillment of the requirements for the Degree of Doctor of Philosophy

Comparative and Biomedical Sciences

Raleigh, North Carolina

2017

APPROVED BY:

______Nanette Nascone-Yoder Troy Ghashghaei Committee Chair

______Jody Gookin Ke Cheng ii

DEDICATION

To my grandmother, for being my guardian angel. To my mom and grandfather, for teaching me that the sky is the limit. To my brothers and fathers, for making me stubborn, hard- headed, and determined. To my friends, girlfriend, and life coach, for keeping me sane. To my animals, for their unconditional love and sloppy kisses.

iii

BIOGRAPHY

Mandy Ashton Womble was born March 6, 1987 to Jeffrey Womble and Lori Chappell. She was raised by her mother, father, stepfather, Gene Chappell, and stepmother, Julie Womble. Mandy grew up in Clemmons, NC alongside her brothers, Matt Womble and Will Chappell. Her dream to become a veterinarian began at an early age, as she was always a lover of animals, biology, and science. Mandy attended West Forsyth High School where she was a scholar and athlete, participating in basketball, volleyball, and softball programs. During her senior year of high school, she was offered an opportunity to participate in an externship program at Clemmons Veterinary Clinic, where her desire to become a vet was solidified. Mandy moved to Raleigh, NC in 2005 to attend North Carolina State University to pursue a Bachelor of Science degree in Zoology. As an undergraduate, she was accepted into the University Honors Program, College of Agricultural and Life Sciences Honors Program, and the honors fraternity, Phi Sigma Pi. During her time at NCSU, Mandy continued to gain experience with animals and veterinary medicine by working at multiple animal hospitals in the area, volunteering at the Carolina Tiger Rescue and NC Zoo, and traveling to South Africa to work at the Center for Animal Rehabilitation and Education where she cared for orphaned, baby baboons. In her junior and senior years, she completed an undergraduate research project in the lab of Dr. Nanette Nascone-Yoder, receiving first place at the NCSU undergraduate research symposium and was published as second author in her first scientific journal. During her time in the lab, she gained an appreciation and love for the scientific process, cell biology, and developmental embryology. After graduating Summa Cum Laude from NCSU in the fall of 2008, Mandy gained employment at Duke University as a research assistant for one year in the laboratory of Brigid Hogan in the Cell Biology department. The next year, she worked in the laboratory of Dr. Garnett Kelsoe in Duke’s Immunology department where she was promoted to research analyst in the first four months and published two more scientific papers. In 2011, Mandy was accepted into NCSU’s College of Veterinary Medicine. During the summer prior to vet school, she joined the vet school club, Turtle Rescue Team, a non-profit organization that rescues and treats injured wild amphibians and reptiles. She has remained an active member of this club, acting in a variety

iv of officer roles including rehabilitation coordinator, captain, and is currently vice president. Participation in this club (and her trip to South Africa) sparked Mandy’s interest in wildlife medicine and launched a desire to work in a museum, aquarium, or wildlife center after graduation. Towards this goal, Mandy decided to apply to the dual DVM/PhD program in order to combine her love for veterinary medicine and research to gain an edge when applying for future museum positions. She was fortunate enough to rejoin her undergraduate research laboratory under the supervision of Dr. Nanette Nascone-Yoder to complete a dissertation project in developmental biology. In her free time, Mandy enjoys playing beach volleyball, backpacking, reading and watching anything of the science-fiction, fantasy, or horror genre. She also loves hanging out with her girlfriend, Casey Snyder, her friends and family, and her own personal zoo including her dogs, Liberty Bell and Kitsune, her cats, Turtle Dove, Skunk, and Spooky, and her geckos, Jerry and Tabitha, in addition to the visiting, rehabilitating turtles that often occupy her bathtub.

ACKNOWLEDGMENTS

“Each person holds so much power within themselves that needs to be let out. Sometimes they just need a little nudge, a little direction, a little support, and a little coaching, and the

greatest things can happen.”

- Pete Carroll

First, I would like to thank my advisor, Dr. Nanette Nascone-Yoder. I entered your lab bright-eyed and bushy-tailed at the age of 19. Somehow you turned that crazy teenager into a scientist. You have always supported my many interests and have guided my research and future career. I have always described you as my “mom” away from home and I truly think of you that way. You have been my shoulder to cry on, my therapist, and my constant support. Most importantly, you have taught me how to be a strong woman and a humble but confident scientist. Thank you for being an inspiring role model.

I also want to thank other members of the NCSU faculty, especially my committee members, Troy Ghashghaei, Jody Gookin, Phillip Sannes, and Ke Cheng for your advice, support, ideas, and mentorship. Dr. Lewbart, my vet school advisor, has been a constant springing board for ideas about my career and has always been available for solid advice.

When people ask me what I want to be when I grow up, I typically say “Dr. Lewbart”. Thank you for being an inspiring role model and friend. I also would like to think my TA supervisor, Colleen Grant, for inspiring my interest in teaching. Thank you to my undergraduate mentor, Dr. Roger Powell for guiding my graduate school decisions.

I also would like to thank the various mentors that have guided and supported me before entering vet school. These include former employers at Duke University, Dr. Brigid Hogan

vi and Dr. Garnett Kelsoe, for instilling in me a love for research. Veterinarians, Dr. Allison

Fox, Dr. Pete Gilyard, Dr. Harold Pierce, and Dr. Amy Neligon allowed me to gain experience in small animal practice.

I would like to thank my lab mates, friends, and family who have helped me remain sane and supported me over all of these years. Thank you to Jordan Ferguson, Melissa

Pickett, Martha Alonzo-Johnsen, Nirav Amin, Mike Dush, and Adam Davis for supporting my research and providing valuable advice towards my project. To my friends, Jennifer

Rodriguez, Denise Monkovich-Pell, Denise Hark, Adam Ward, Ryan Walters, and Chelsey

Vanetten for your love and support. I would also like to express my gratitude and love to my girlfriend, Casey Snyder, for her loyalty and encouragement in the last year of my Ph.D.

Finally, I would like to thank my “Mama”, Lori Chappell. From an early age, you have taught me to have the “eye of the tiger”. You have supported and encouraged me throughout my whole life, no matter how unattainable my dreams may have seemed. You taught me the strength of women, the loyalty of a best friend, and most importantly the perseverance to accomplish anything I set my mind too. Thank you for being my number one fan and the best cheerleader anyone could ask for.

vii

TABLE OF CONTENTS

LIST OF TABLES ...... ix LIST OF FIGURES ...... x CHAPTER 1: Introduction ...... 1 1) The Establishment of Left-right Asymmetry in Development...... 4 1.1) The initial left-right symmetry breaking event at the “node” ...... 4 1.2) The transfer of left-right cues from the “node” to the lateral plate mesoderm (LPM) ...... 7 1.3) The establishment of the midline barrier ...... 9 1.4) The induction of left-right asymmetric organogenesis ...... 10 1.5) The evolutionary conservation of LR asymmetry ...... 11 1.6) The discovery and characterization of Pitx2 ...... 12 2) Left-right Asymmetric Organogenesis ...... 13 2.1) Heart looping ...... 14 2.2) Asymmetric regression of blood vessels closely associated with the heart ...... 20 2.3) Gut looping ...... 21 2.4) Vascular and Lymphatic asymmetries in the dorsal mesentery...... 29 2.5) Stomach Curvature ...... 31 2.6) Asymmetric regression of the spleen ...... 33 2.7) Lung Lobation ...... 33 2.8) Liver lobation ...... 34 2.9) Conclusions ...... 37 3. Rationale and Hypothesis ...... 37 References ...... 45 CHAPTER 2: The left-right asymmetry of liver lobation is generated by Pitx2c- mediated asymmetries in the hepatic diverticulum ...... 50 Introduction ...... 51 Materials and Methods ...... 52 Animals ...... 52 Immunohistochemistry ...... 53 RNA in situ hybridization ...... 53 X. laevis Loss- and Gain-of-function experiments ...... 54 Measurements and Statistics ...... 54 Results ...... 55 The liver diverticulum is left-right asymmetric ...... 55 Cellular left-right asymmetries in the early liver diverticulum ...... 56 Liver asymmetry is under the control of the left-right asymmetry pathway ...... 57 Pitx2c is required for left lobe morphogenesis ...... 57 Ectopic R-sided Pitx2c induces heterotaxy ...... 58 Pitx2c is sufficient to induce left-sided cell character ...... 59 Discussion ...... 60 Pitx2c is necessary and sufficient to induce left-sided epithelial characteristics ...... 60 Pitx2c’s influence on cell behavior is organ specific ...... 61

viii

Ramifications for human health ...... 63 References ...... 81 CHAPTER 3: Discussion ...... 86 1) Pitx2 induces left-right asymmetries in cell shape ...... 86 1.1) Pitx2’s influence on Apical-Basal Cell Elongation and Apical Constriction ...... 87 1.2) The role of cell adhesion in cell shape change ...... 89 2) Pitx2c directs left-right asymmetrical epithelial remodeling ...... 91 2.1) Cellular rearrangement ...... 92 2.2) Interkinetic nuclear migration ...... 94 3) Pitx2c expression in the mesoderm influences endodermal tissue architecture ...... 97 3.1) Endodermal and Mesodermal interactions mediated by signaling ligands/receptors ...... 98 3.2) Mesodermal and Endodermal interactions mediated by ECM components ...... 99 3.3) Differential expression of Pitx2 in organ tissues layers ...... 101 4) Downstream targets of Pitx2c during left-right liver development ...... 103 4.1) Components of the Wnt signaling pathway ...... 103 4.2) ECM components ...... 106 4.3) Soluble Growth Factors ...... 109 4.4) Chemokines ...... 110 4.5) Conclusions ...... 111 5) Ramifications for human health ...... 112 References ...... 115 CHAPTER 4: Conclusions and Future Directions ...... 121 References ...... 124 APPENDIX ...... 127 Appendix A: Budgett’s frog (Lepidobatrachus laevis): a new amphibian embryo for developmental biology ...... 128 Appendix B: Frogs as integrative models for understanding digestive organ development and evolution ...... 170 Appendix C: Developmental Constraints on Endoderm Morphogenesis Underlie the Evolution of Gut Length ...... 225 Appendix D: Foregut gene expression in X. laevis...... 239

LIST OF TABLES

Table 3- 1) Genes with Pitx2 binding sites in the promoter ...... 113 Table 3- 2) Direct protein-protein binding partners of Pitx2 ...... 113

Table B- 1) Anthropogenic toxicants found to disrupt gut development in Xenopus laevis ...... 196

LIST OF FIGURES

Figure 1- 1) LR asymmetric organ placement and morphology is vital for human health ...... 41 Figure 1- 2) Four steps of initial LR symmetry breaking events during embryogenesis 42 Figure 1- 3) LR asymmetric organogenesis in the midgut and stomach ...... 44

Figure 2- 1) Early development of LR asymmetry in the Xenopus liver ...... 64 Figure 2- 2) Cellular level LR asymmetries in the early liver diverticulum ...... 66 Figure 2- 3) The Nodal pathway is required for liver asymmetry ...... 68 Figure 2- 4) Left-sided Pitx2c is required for LR liver asymmetry ...... 69 Figure 2- 5) Knockdown of Pitx2c increases left-sided apical constriction and cell elongation ...... 70 Figure 2- 6) Ectopic right-sided Pitx2c expression induces liver heterotaxia in the form of left isomerism ...... 72 Figure 2- 7) Ectopic right-sided Pitx2c expression reduces apical constriction and cell elongation ...... 73 Figure 2- 8) Model of the role of Pix2c in asymmetrical liver development ...... 75 Supplementary Figure 2- 1) The left and right cells of the liver diverticulum originate from the left and right sides of the early embryo ...... 76 Supplementary Figure 2- 2) Cell number and proliferation are not different between the left and right sides of the developing liver ...... 77 Supplementary Figure 2- 3) Pitx2 is asymmetrically expressed in the left mesoderm surrounding the liver diverticulum ...... 78 Supplementary Figure 2- 4) Pitx2c microinjection manipulation strategies ...... 79 Supplementary Figure 2- 5) Ectopic right-sided Pitx2c expression decreases diverticulum length ...... 80

Figure 3- 1) Possible downstream targets of Pitx2c ...... 114

Figure A- 1) The schedule of Lepidobatrachus laevis embryonic development (versus Xenopus laevis) ...... 152 Figure A- 2) Early cleavage and gastrulation patterns in L. laevis are similar to Xenopus ...... 153 Figure A- 3) Neurulation and organogenesis in L. laevis ...... 154 Figure A- 4) Craniofacial morphogenesis in L. laevis ...... 155 Figure A- 5) Thermal tolerance of L. laevis and X. laevis embryos ...... 157 Figure A- 6) L. laevis embryos are amenable to microinjection of exogenous reagents for fate-mapping and expression of synthetic mRNA ...... 158 Figure A- 7) L. laevis explants are amenable to animal cap explant culture ...... 159 Figure A- 8) Heart morphogenesis occurs at a larger scale in L. laevis ...... 160 Figure A- 9) Limb regeneration occurs at a rapid pace in L. laevis ...... 162

Figure B- 1) Wnt, RA, and FGF pattern the foregut ...... 204 Figure B- 2) Intestine lengthening involves Hedghog- and Wnt/PCP-mediated endoderm cell polarization, rearrangement and epithelial differentiation ...... 205 Figure B- 3) Altered RA signaling may have led to a novel foregut morphology ...... 207 Figure B- 4) Endoderm morphogenesis in ancestral versus direct-developing frog species ...... 208

Figure C- 1) Variation in tadpole feeding ecology correlates with variation in egg size (yolk reserves) and gut length ...... 227 Figure C- 2) All endodermal cells contribute to the gut epithelium in exotrophic and terrestrial, delayed herbivorous tadpoles, regardless of egg size ...... 229 Figure C- 3) The innermost endoderm cells do not contribute to the gut epithelium in rapidly-developing, exotrophic carnivorous tadpoles, regardless of egg size ...... 231 Figure C- 4) Endoderm rearrangement increases epithelial surface area, but not length, in endotrophic (direct developing) tadpoles ...... 233 Figure C- 5) Sonic Hedgehog signaling increases intestinal surface area and reduces apoptosis in cannibalistic tadpoles ...... 235 Figure C- 6) Changes in endoderm morphogenesis may underlie anuran gut evolution ...... 237

Figure D- 1) Gene Expression patterns of transverse cross sections at NF 35.36 ...... 240 Figure D- 2) Gene expression patterns of frontal sections at NF 37.38...... 243 Figure D- 3) Gene expression patterns of transverse cross sections at NF 37.38 ...... 246 Figure D- 4) Gene expression patterns of frontal sections at NF 39 ...... 249 Figure D- 5) Gene expression patterns of transverse cross sections at NF 39.40 ...... 252 Figure D- 6) Gene expression patterns of frontal sections at NF 40.41 ...... 254 Figure D- 7) Gene expression patterns of transverse cross sections at NF 40.41 ...... 257 Figure D- 8) Gene expression patterns of transverse cross sections at NF 41 ...... 260

CHAPTER 1: Introduction

Vertebrates are members of the group Bilateria, meaning they have symmetrical external body plans consisting of dorsal/ventral (DV), anterior/posterior (AP), and left/right

(LR) axes; however, vertebrate internal organs are LR asymmetrical in both anatomical location and morphology including the liver. The liver is positioned on the right side of the body cavity and liver morphology is LR asymmetric in the size and number of left and right lobes. In humans, the right side of the liver is 5-6 times larger than the left and consists of four lobes: the medial, lateral, caudate, and quadrate while the left only consists of the medial and lateral lobes (Gray, 1918). In addition to the liver, many other internal organs are LR asymmetrical. The stomach, heart, and spleen are all positioned on the left side of the body cavity opposite to liver placement. The right and left atria and ventricles, the heart valves, and the associated blood vessels, including the superior vena cava, aorta, and left and right pulmonary arteries, all have LR asymmetries vital for the separation of oxygenated and deoxygenated blood as well as to orient the direction of blood flow through the heart and lungs and throughout the rest of the body. The gastrointestinal tract always loops in a counter clockwise direction, a mechanism vital to packaging the long length of the intestines inside the body cavity efficiently (Burn and Hill, 2009). Like the liver, the lung has asymmetries in size and lobation as the right lung is larger and has 3 lobes: the superior, middle, and inferior lobes while the left lung only consists of superior and inferior lobes (Gray, 1918). The coordination of the left-right asymmetrical body axes within organs, between organs, and as they connect with the vascular system is vital for health and development.

As many as 1 in 10,000 human infants are born with defects in left-right asymmetry termed heterotaxia (Lin et al., 2000). The normal positioning of the organs within the body cavity is called situs solitus (Fig. 1.1A). The complete reversal of all organ placement and morphology is called situs inversus (Fig. 1.1B). Situs inversus occurs in 1 in 8,500 human births (Basu and Brueckner, 2008; Brueckner, 2007). These individuals are relatively normal and can live their entire lives without knowledge of their condition because the complete body axis is reversed and therefore functions correctly. Problems occur in cases of heterotaxy when one or more organs have defects in LR asymmetry that affect the function of the organ or the connection of that organ with other organs, vasculature, lymphatics, or ducts. Infants with heterotaxy syndrome have a fatal prognosis in 90-99% of cases (Lee et al., 2014).

Two of the most common heterotaxy associated diseases are right-isomerism (Fig.

1.1C) and left-isomerism (Fig. 1.1D). Infants with right isomerism often present with midline stomach, common atrioventricular canal, univentricular heart, transposition of the great arteries, and anomalous venous connection (Applegate et al., 1999). Individuals with left- isomerism have bilateral bi-lobed lungs, bilateral pulmonary atria, multiple spleens, stomachs in indeterminate positions, partial anomalous pulmonary venous return, atrial septal defects and common atrioventricular canal (Applegate et al., 1999). Over 50% of patients with heterotaxy syndrome also present with midline, symmetrical livers with abnormal lobation

(Burton et al., 2014). Another subset of heterotaxy patients have extrahepatic biliary atresia, extrahepatic portosystemic shunt, and preduodenal portal veins requiring surgical correction and in severe cases, neonatal liver transplantation (Gottschalk et al., 2016; Newman et al.,

2010; Pathak and Sarin, 2006). Even after surgical intervention, individuals continue to

3 present with intermittent biliary cirrhosis and require additional treatment and surgery throughout their lives (Kerkeni et al., 2015). The association of liver heterotaxia with other organ laterality defects suggests a common development program.

Research over the past several decades has given us insight into the etiology of heterotaxia including knowledge about initial left-right axis determination, cellular level mechanisms of LR organ morphogenesis, and genes required for this process like the transforming growth factor beta (TGF-growth factor, Nodal and its downstream target, the

Paired like homeodomain 2 (Pitx2) that is expressed on only the left side of many organs as they develop. Researchers are just beginning to understand the LR differences in tissue morphogenesis including cell shape changes, cell migration, and tissue remodeling that are required for organ laterality in the stomach, heart, and midgut; however, the cellular level differences required for the development of LR asymmetries in liver lobation are unknown.

Left-sided expression of Pitx2 is required for stomach curvature, intestinal rotation, heart looping, lung laterality, and asymmetric vasculature development (Davis et al., 2008;

Davis et al., 2017; Kitamura et al., 1999; Mahadevan et al., 2014). Pitx2 is expressed on the left side of the septum transversum mesenchyme (STM), a mesodermal tissue that surrounds the liver during development; however, it is unknown whether Pitx2 is required for asymmetric liver size and lobation (Shiratori et al., 2006). This introductory chapter is a comprehensive review on the initial establishment of LR asymmetry, general mechanisms of

LR asymmetrical organogenesis, and the known role of Pitx2 in this process.

1) The Establishment of Left-right Asymmetry in Development

The establishment of LR asymmetry consists of 4 major steps: (1) The initial symmetry breaking event in or near the embryonic “node” (Fig. 1.2A), (2) The transfer of these LR cues to the lateral plate mesoderm (LPM) by the asymmetric expression of the signaling molecule Nodal (Fig. 1.2B), (3) The establishment of a midline barrier (Fig. 1.2C), and (4) the induction of LR asymmetric organogenesis (Fig. 1.2D) (Ibanes and Izpisua

Belmonte, 2009; Raya and Izpisua Belmonte, 2006; Shiratori et al., 2006; Yamamoto et al.,

2003).

1.1) The initial left-right symmetry breaking event at the “node”

In all vertebrate species studied including zebrafish, Xenopus, chick, rabbit, and mouse, the presence of a LR organizer or node is required for LR axis determination:

Kupffer’s vesicle in zebrafish, Spemann’s Organizer in Xenopus, Henson’s node in chick, the posterior notochord in rabbits, and the node in mice (Basu and Brueckner, 2008; Blum et al.,

2014; Ibanes and Izpisua Belmonte, 2009; Raya and Izpisua Belmonte, 2006; Shiratori et al.,

2006). Although the timing and geometry of the LR organizer varies among species, all involve a shift from symmetrical expression of the TGF- growth factor, Nodal, to left-sided

Nodal expression surrounding the LR organizer (Basu and Brueckner, 2008). How left-sided expression of Nodal is first established has been a matter of much debate, however, recent research points towards a cilia driven, nodal flow hypothesis in which directional beating of cilia within the node creates an asymmetric flow of perinodal fluid containing LR

5 determinants that initiate left-limited Nodal expression (Basu and Brueckner, 2008, Blum et al., 2014).

The first evidence that cilia were required for the establishment of the LR axis came with the discovery of a correlation between human disorders involving ciliogenesis or ciliary function and LR defects. As many as 50% of patients with Kartagener syndrome or primary ciliary dyskinesia, a disorder caused by immotile cilia, have defects in LR asymmetrical organ development (Basu and Brueckner, 2008). Additionally, 13 independent mouse mutations that affect ciliary biogenesis and function also cause LR defects (Basu and

Brueckner, 2008). For example, mice with defects in microtubule associated motor proteins in which no cilia are present [kinesin-like protein (Kif3a/b) mutants], or cilia are immotile

[left-right dynein (Lrd) mutants], have abnormalities in LR asymmetric organogenesis (Basu and Brueckner, 2008; Shiratori et al., 2006; Wagner and Yost, 2000).

Upon further evaluation of the nodal flow hypothesis in mice, long, monocilia were found on the ventral columnar pit cells of the node (Basu and Brueckner, 2008; Ibanes and

Izpisua Belmonte, 2009; Shiratori et al., 2006). These cilia rotate in a counter clockwise direction with the appropriate tilt angle to generate leftward flow of embryonic fluid (Basu and Brueckner, 2008; Brennan et al., 2002; Blum et al., 2014; Raya and Izpisua Belmonte,

2006; Shiratori et. al., 2006; Wagner and Yost, 2000). The localization and tilt of the cilia within the cells of the node is most likely established by preexisting DV and AP axes, similar to the process of positioning hairs in the cuticles of Drosophila or in the vertebrate inner ear by the planar cell polarity (PCP) pathway (Shiratori et. al., 2006). In this manner, the tilt and

6 position of the cilium is established by the localization of the basal body within the cell

(Shiratori et. al., 2006).

All of the vertebrate LR organizers studied have motile primary cilia in the node except for the chick (Basu and Brueckner, 2008). It is possible that avian species have a different mechanism than “nodal flow” to establish asymmetric Nodal expression but it is more likely that researchers have not observed nodal cilia in the chick yet due to the imaging constraints of the species (Basu and Brueckner, 2008). In other vertebrates, the direction of cilia beating is conserved although the beating frequency varies, most likely due to the constraints of each embryonic strategy. In rabbits, the cells of the node itself do not have motile cilia but cells at the posterior segment of the notochord do have cilia and are surrounded by Nodal expression (Basu and Brueckner, 2008). Fluid flow has been observed in Xenopus, rabbit, zebrafish, and mouse (Basu and Brueckner, 2008). In Xenopus, increasing the viscosity of the fluid induces LR defects, possibly by preventing normal fluid flow.

Morpholino knockdown of the genes involved in cilia motility and assembly in zebrafish also causes LR defects (Basu and Brueckner, 2008). In general, nodal flow generates a LR difference within the node that is subsequently amplified as this signal is transferred to the left LPM (Shiratori et al., 2006).

All vertebrates need the embryonic node and the left-sided expression of Nodal around this node for LR cues. In mammals, nodal flow is the first evidence of laterality (Basu and Brueckner, 2008; Raya and Izpisua Belmonte, 2006). However, in other species, the first sign of an initial symmetry breaking event begins before the formation of the LR organizer.

In frogs (X. laevis), the first sign of LR asymmetry is the asymmetric localization of

7 maternally derived H+/K+ ATPase subunit transcripts (Ibanes and Izpisua Belmonte, 2009;

Raya and Izpisua Belmonte, 2006). In chicks, the first evidence is LR differences in ion fluxes or membrane voltage potential as the left side is more depolarized than the right. This is due to differences in the activity of the H+/K+ ATPase pump as pharmacological inhibition results in organ heterotaxia (Raya and Izpisua Belmonte, 2006). This mechanism is also found in zebrafish, urochordates, and echinoderms (Duboc et al., 2005; Raya and Izpisua

Belmonte, 2006). Therefore, no model of initial symmetry breaking is the same for all animals.

1.2) The transfer of left-right cues from the “node” to the lateral plate mesoderm (LPM)

In all vertebrates, the initial asymmetric left-sided Nodal expression surrounding the embryonic LR organizer or node is required for subsequent left-sided Nodal expression in the

LPM, a tissue located in the lateral region of the embryo at somite stages that eventually contributes to the mesenchyme of visceral organs (Raya and Izpisua Belmonte, 2006; Saijoh et al., 2003; Shiratori et al., 2006; Yamamoto et al., 2003). Ablation of Nodal in the region surrounding the LR organizer eliminates Nodal expression in the LPM (Shiratori et al.,

2006). However, how the initial Nodal signal is transferred from the node to the LPM is still unknown.

Nodal may act through cells that are located between the node and the LPM (e.g., perinodal cells, endodermal cells, paraxial mesodermal or intermediate mesoderm) to transfer a secondary signal to the LPM (Shiratori et al., 2006). In chick, Sonic Hedgehog (Shh) is also expressed asymmetrically in the node and activates the BMP inhibitor, Caronte, in the

8 paraxial mesoderm between the node and the LPM. Caronte induces Nodal within the left

LPM (Shiratori et al., 2006). It is unknown whether a similar mechanism exists in other vertebrates because a Caronte-like BMP antagonist has not been found (Shiratori et al.,

2006). Interestingly, however, in Xenopus, the chemical inhibition of Smad1 and Smad5

(BMP effectors) causes Nodal to be expressed bilaterally in the LPM, hinting that a similar mechanism is yet to be discovered.

There is evidence in mice that growth differentiation factor 1 (GDF-1), a TGF- related protein, is required for the transfer of the Nodal signal from the node to the LPM

(Raya and Izpisua Belmonte, 2006; Shiratori et al., 2006). GDF-1 is expressed in the perinodal region, in a similar expression pattern to Nodal. GDF1 mutant mice do not have

Nodal expression in the LPM and the organs of these animals are heterotaxic. GDF-1 is also expressed in the LPM prior to Nodal expression, suggesting that GDF-1 may make the LPM competent for Nodal signaling (Shiratori et al., 2006). Although these interesting correlations have been found between GDF-1 and Nodal expression in the LPM, additional signaling molecules must be involved as GDF-1 alone cannot activate Nodal signaling when overexpressed in cultured cells or frog embryos (Shiratori et al., 2006).

Another possible mechanism is that Nodal itself is transported from the node to the

LPM directly without intermediate signaling as ectopic Nodal expression in the right LPM induces endogenous Nodal signaling (Saijoh et al., 2003; Shiratori et al., 2006). More research is needed to uncover the mechanism by which the asymmetric Nodal signal is transferred from the LR organizer to the LPM. It likely that a combination of signaling pathways are involved that act independently or together and that some of these mechanisms

9 are species specific. What remains conserved between all vertebrates studied is that left-sided asymmetric expression of Nodal within the LPM is required for the establishment of the midline barrier and subsequent LR asymmetrical organogenesis.

1.3) The establishment of the midline barrier

Left-sided nodal expression in the LPM begins in a region at the level of the node and expands along the AP axis where Nodal is positively regulated by itself and induces the expression of LR determination factor-2 (Lefty-2) along the LPM and Lefty-1 at the midline

(Brennan et al., 2002; Saijoh et al., 2003; Shiratori et al., 2006; Yamamoto et al., 2003).

Lefty is a Nodal feedback inhibitor that acts to restrict the domain and duration of Nodal expression (Raya and Izpisua Belmonte, 2006; Saijoh et al., 2003; Shiratori et al., 2006;

Yamamoto et al., 2003). In mammals, Lefty-2 is expressed throughout the left LPM while

Lefty-1 is expressed at the midline and is required for midline barrier function (Yamamoto et al., 2003). The midline barrier sequesters Nodal expression to only the left side (Yamamoto et al., 2003). In mice mutants for Lefty-1 or Lefty-2, Nodal expression begins asymmetrically in the left LPM but then crosses over to the right, causing bilateral expression and heterotaxia of visceral organs (Shiratori et al., 2006). Other signaling molecules, like Sonic hedgehog

(Shh) and Indian hedgehog (Ihh), secreted from the notochord may also be required for this process (Shiratori et al., 2006). Future research is needed to understand the function of the midline barrier, other genes involved, and the mechanism of action of Lefty proteins.

1.4) The induction of left-right asymmetric organogenesis

Nodal expression in the left LPM is required for LR visceral organ development.

Nodal mutants are heterotaxic; stomach curvature is randomized, spleens are reduced in size, the lung is double right-sided, heart looping is randomized with transposition of the great arteries, atrial-septal defects, and common atrial chambers, there are abnormalities in liver lobation including clefts in the medial lobes, and gastrointestinal tract coiling is disorganized

(Saijoh et al., 2003). Although expression of Nodal within the LPM is a transient event, only lasting from the 2-6 somite stages in mice, a signaling cascade is initiated that sets up the entire LR internal body plan (Basu and Brueckner, 2008; Ibanes and Izpisua Belmonte, 2009;

Shiratori et al., 2006). Nodal induces the expression of the homeobox transcription factor,

Pitx2, in the left LPM (Basu and Brueckner, 2008; Ibanes and Izpisua Belmonte et al., 2009;

Raya and Izpisua Belmonte, 2006; Shiratori et al., 2006). Pitx2 expression is maintained, even after Nodal expression disappears, by the homeobox transcription factor, NK2 homeobox 5 (Nkx2.5) (Basu and Brueckner, 2008). Pitx2 is the only known downstream target of Nodal signaling still expressed during organogenesis and is also required for LR organ development as Pitx2 deficient mice phenocopy Nodal mutants in visceral organ heterotaxia (Gage et al., 1999; Kitamura et al., 1999; Lin et al., 1999; Liu et al., 2001; Lu et al., 1999; Raya and Izpisua Belmonte, 2006; Shiratori et al., 2006).

The downstream cellular mechanism by which Pitx2 directs organ laterality is just starting to be uncovered. Research over the past few decades has shown that left-sided Pitx2 expression is required for cell shape changes, epithelial remodeling, and/or cell rearrangements during intestinal looping and stomach curvature (Davis et al., 2008; Davis et

11 al., 2017). However, the mechanism by which Pitx2 confers morphological laterality in other organs like the lung, heart, spleen, and liver is still unknown.

1.5) The evolutionary conservation of LR asymmetry

The role of Nodal signaling and the left-sided expression of Pitx2 is well established in vertebrates. Interestingly, the LR asymmetry pathway is a conserved feature of all chordates as amphioxi (cephalochordates) and ascidians (urochordates) also show left-sided

Nodal and Pitx2 expression (Duboc et al., 2005). The Nodal signaling cascade can even be found in lophotrochozoans as mollusks and snails require the asymmetric expression of a

Nodal homolog for shell coiling (Grande and Patel, 2009). The Nodal signal was lost in ecdysozoans as no Nodal-like protein has been found (Blum et al., 2014). Drosophila instead utilizes actin filament based systems to orchestrate gut rotation (Okumura et al., 2008).

Homologs of Nodal, Pitx2, and Lefty are found asymmetrically expressed in echinoderms, however, this expression is on the right side, not the left (Blum et al., 2014; Duboc et al.,

2005). Adult echinoderms are radially symmetrical but evolved from a bilateral ancestor and have bilateral larvae. These larvae are LR asymmetric as the imaginal disc or rudiment is on the left side of the embryo (Duboc et al., 2005). Interestingly, Sry-related HMG box E (Sox

E), a protein related to Sry-related HMG box 9 (Sox 9), is expressed in the left coelomic pouch (Duboc et al., 2005). Inhibition of Nodal signaling eliminates right sided Nodal, Pitx2, and Lefty expression while Sox E expression becomes bilateral (Duboc et al., 2005).

Conversely, ectopic expression of Nodal with Activin treatment induces bilateral Nodal,

Pitx2 and Lefty expression while eliminating Sox9 expression (Duboc et al., 2005).

The conservation of Nodal signaling across multiple animal Phyla suggests that

Nodal evolved as a LR determination gene before the split between Protostomes and

Deuterostomes and that the role of Pitx2 as a downstream effector of Nodal evolved before the separation of echinoderms and chordates. It is very interesting that the expression of the

Nodal pathway is right sided in echinoderms and left sided in chordates (Duboc et al., 2005).

This could raise interesting questions as to whether our definition of left versus right in echinoderms is backwards or whether they have a complete inversion of the DV axes (Duboc et al., 2005).

1.6) The discovery and characterization of Pitx2

Pitx2 was first discovered for its role in Axenfeld-Riegar syndrome (ARS). ARS is an autosomal dominant, haploinsufficiency disorder characterized by a variety of birth defects in eye, tooth, pituitary, umbilicus, facial structure, intestinal tract, and abdominal wall development (Hjait and Semina, 2005). Thirty different mutations in the Pitx2 gene located at chromosomal locus 4q25 are associated with ARS; mostly point mutations in the homeodomain (Hjait and Semina, 2005). Pitx2 has several splicing and transcriptional variants including Pitx2 a, b, c, and d. Each isoform has alternatively spliced exons (Hjait and Semina, 2005; Schweickert et al., 2000). Pitx2a and b have different promoters than

Pitx2c and d (Hjait and Semina, 2005; Schweickert et al., 2000). All isoforms share their protein-protein interaction domain in the C-terminus (Hjait and Semina, 2005; Schweickert et al., 2000). Pitx2a has the shortest N-terminal region while Pitx2b and c have different, large

N-termini (Hjait and Semina, 2005; Schweickert et al., 2000). Pitx2d is a truncated version of

13 the protein with a nonfunctional homeodomain (Hjait and Semina, 2005). Pitx2a, b, and c are expressed widely throughout the body, particularly in the craniofacial region, eye, oral ectoderm, pituitary, body wall, central nervous system, and internal organs while Pitx2d is only found in the craniofacial region (Hjait and Semina, 2005, Liu et al., 2003). Pitx2c is the only isoform that has an asymmetrical expression, found on the left side of visceral organs

(Schweickert et al., 2000). In cell culture, each isoform has different gene promoter targets so they most likely have diverged functions (Cox et al., 2002; Hjait and Semina, 2005). Pitx2c but not Pitx2a or Pitx2c is required for asymmetric organ morphogenesis as misexpression induces heterotaxia of the lungs, heart, spleen, and gut (Liu et al., 2001; Kitamura et al.,

1999; Schweickert et al., 2000; Shiratori et al., 2006). In human heterotaxy patients, LR congenital liver and biliary defects are often linked with defects of the other visceral organs, therefore, it is likely that Pitx2 is also required for LR liver development, although this has never been described.

2) Left-right Asymmetric Organogenesis

The ultimate outcome of LR axis formation is the transformation of LR cues (like

Pitx2) into LR asymmetric organogenesis. The heart, lungs, stomach, gastrointestinal tract, liver, and spleen have LR asymmetric morphology. Although they use the same initial signaling pathways to develop laterality (i.e. Nodal), each of these organ systems undergoes differential development based on the tissues involved, the geometry and shape of the structures, and the functions vital for embryonic and adult survival. There are three main outcomes of asymmetric organ development: 1) directional looping as in tubal structures like

14 the heart, gut, and stomach, 2) LR differences in organ size and branching as in the lungs and liver, and 3) unilateral regression of organs like the spleen and structures like blood vessels

(Shiratori et al., 2006). Cellular level mechanisms of LR asymmetrical development have recently been discovered during heart looping, vasculature development, stomach curvature, and midgut rotation. Left-sided expression of Pitx2 is required for LR development in the heart, vasculature, lungs, spleen, intestine, and stomach (Liu et al., 2001; Kitamura et al.,

1999; Schweickert et al., 2000; Shiratori et al., 2006). Downstream cellular level mechanisms of Pitx2 have been described during vasculature, stomach, and midgut development but have yet to be discovered in other organ systems including the lungs and heart (Davis et al., 2008;

Davis et al., 2017; Mahadevan et al., 2014; Muller et al., 2003). Knowledge pertaining to the mechanisms of LR development and their link to Pitx2 will be valuable when determining how liver laterality is established. This section of the introductory chapter reviews mechanisms of LR development during organogenesis in a variety of different model organisms.

2.1) Heart looping

The heart is the first organ to develop with LR asymmetric morphology. There are 5 phases of cardiac development that are crucial for the proper form and function of the heart.

The first phase (“pre-looping”) is when cells are bilaterally specified to form the heart fields.

Heart precursors arise from bilaterally paired areas of the splanchnic mesoderm (Amand et al., 1998; Bakkers et al., 2009; Manner, 2009). The two heart fields then fuse at the ventral embryonic midline to form a straight, symmetric tube (Amand et al., 1998; Bakkers et al.,

2009; Manner, 2009). The second phase is “dextral looping” and occurs as the straight tube forms a C-shaped loop with the convex portion towards the right side of the embryo

(Manner, 2009). The third phase is the “early phase of S-looping” when the arterial and venous poles shorten, causing the C-shape to transform into an S-shape and the ventricular bend to move from a position cranial to the atria to a more caudal position (Manner, 2009).

During phase four, the “late S-phase”, the proximal outflow tracts move leftward (Manner,

2009). At this time, the arterial trunk anlage develops at the distal portion of the outflow tract and the atrial and ventricular portions of the heart balloon outward (Manner, 2009). During phase five, “cardiac septation”, septal structures first appear, dividing the heart lumens and arterial trunks into left and right compartments (Manner, 2009). The proper establishment of

LR asymmetry is vital for all phases of cardiac morphogenesis.

The earliest evidence of LR asymmetry during heart morphogenesis is in gene expression patterns in the primary heart field prior to fusion at the midline (Ramsdell, 2005).

Extracellular matrix (ECM) proteins exhibit LR asymmetries in expression levels; Fibrillin-2 is higher on the right side while heart specific lectin-associated matrix protein-1 (hLamp-1) and Flectin are expressed higher on the left (Ramsdell, 2005). In fact, the asymmetric expression of Flectin is maintained even after initial heart tube fusion (Ramsdell, 2005). In addition to gene expression asymmetries, cardiomyocyte differentiation and myofibril formation begin slightly sooner in the right early heart field (Ramsdell, 2005).

During the next stages of organogenesis, cells from the original left or right heart fields contribute to different portions of the developing heart, indicating asymmetries in cell rearrangements and migration (Gormley and Nascone-Yoder, 2003; Ramsdell, 2005). The

16 anterior portion of the heart is larger on the right side while the posterior portion is larger on the left as cells derived from the right and left heart fields contribute to the anterior and posterior regions differently (Ramsdell, 2005). A larger proportion of cells originating from the right heart field contribute to the posterior or inflow region (Ramsdell, 2005). Fate- mapping experiments in Xenopus laevis reveal that, later in development, the left side contributes to the inner curvature and the ventral face of the loop while the right side contributes to the outer curvature and dorsal portion of the loop (Gormley and Nascone-

Yoder, 2003). The localization of the left-sided cells correlates to the expression pattern of

Pitx2, indicating that the Nodal-Pitx2 pathway may be involved in patterning of these regions

(Gormley and Nascone-Yoder, 2003).

Pitx2 is expressed in the entire left side of the heart throughout early heart morphogenesis and loop formation and is later expressed in the left atrioventricular canal, left atrium, interatrial septum, and left caval vein (Amand et al., 1998; Gormley and Nascone-

Yoder, 2003). Nodal and Pitx2 are required for normal LR asymmetrical heart development as mutations in both genes cause LR congenital defects such as double outlet right ventricle, transposition of the great arteries, atrial and ventricular septal defects, anomalous venous return, right atrial isomerism, persistent truncus arteriosus, and aortic arch anomalies

(Bakkers et al., 2009; Manner, 2009; Ramsdell, 2005; Saijoh et al., 2003). The downstream mechanisms of Pitx2 during LR asymmetrical heart morphogenesis remain mostly unknown, however, research over the past several decades has shed light on the early processes of dextral looping, using the zebrafish as a model system.

In zebrafish, the adult heart is relatively simple, consisting of sinus venous, atrium, ventricle, and bulbous arteriosus segments (Hu et al., 2000). During development, there are two asymmetric events: a leftward displacement of the developing heart termed “heart jogging”, and the formation of the c-shaped curvature, “dextral looping” (Manner, 2009). In zebrafish, like other vertebrates, the heart is derived from bilaterally symmetrical mesoderm populations that undergo organized, collective movements towards the ventral embryonic midline where they fuse together to form the cardiac cone (Bakkers et al., 2009; Smith et al.,

2008). The ECM protein, Fibronectin, is crucial for this midline migration as natter mutants that lack Fibronectin develop cardia bifidia due to a failure of coordinated cell movements towards the midline (Bakkers et al., 2009). Fibronectin is important in providing the correct path of migration (Bakkers et al., 2009). After the fusion of these cells at the midline, jogging occurs due to displacement of the heart cone to the left side of the embryo (Bakkers et al.,

2009). Both the mesocardium and endocardium, formed from different precursor populations, are displaced left at the same time although the endocardium population is dispensable in this asymmetric displacement as cloche mutants that lack endothelial cells still display cardiac jogging (Bakkers et al., 2009).

Following the initial left-ward displacement of the heart, dynamic cell rearrangements in the cardiac cone contribute to the dextral looping or c-shaped curvature that follows

(Bakkers et al., 2009). Myocardial cells on the right side of the cardiac cone move anteriorly and ventrally towards the left (Bakkers et al., 2009). These cells will eventually form the ventral part of the cardiac tube while the left side contributes to the dorsal roof of the tube, changing previous RL locations to DV respectively (Bakkers et al., 2009). Differential

18 migration behaviors of specific regions of the heart tube itself contribute to the directionality of rotation (Bakkers et al., 2009; Smith et al., 2008). By tracking individual cells during this migration process, Smith et al. (2008) found that all myocardial cells were linearly displaced to the left and anterior but that the speed and consistency of movement depended on whether the cells were anteriorly or posteriorly derived. Posterior cells did not move at a constant speed but rather started at the same speed as anterior cells before doubling their speed, eventually having a higher displacement rate (Bakkers et al., 2009; Smith et al., 2008). The anterior cells moved at a constant speed the entire time (Bakkers et al., 2009; Smith et al.,

2008). The inconsistencies of cell movement causes the cardiac cone to rotate in a clockwise direction when observed from a dorsal view (Bakkers et al., 2009; Smith et al., 2008).

Hyaluronic synthase 2 (Has2), a protein that produces the ECM protein, hyaluronic acid (HA), and is asymmetrically expressed on the embryonic left side during asymmetric heart development, is required for the cell migration events during both heart jogging and dextral looping as has2 mutants have a failure of leftward cardiac cone displacement as well as a failure of cardiac rotation (Bakkers et al., 2009; Smith et al., 2008). In has2 mutants, anterior and posterior cells of the cardiac cone no longer migrate at different speeds (Bakkers et al., 2009; Smith et al., 2008). Posterior cells instead migrate at the same constant speed as anterior cells, disrupting normal cardiac rotation events (Bakkers et al., 2009; Smith et al.,

2008). While Has2 appears to direct the differential speed of migration, bone morphogenetic protein (Bmp) interacts with Has2 to direct the path and directionality of the migration

(Bakkers et al., 2009; Smith et al., 2008).

Bmp4 is expressed asymmetrically in the left LPM during cardiac looping events

(Bakkers et al., 2009; Smith et al., 2008). This asymmetric expression is required for dextral looping as bmp4 mutants have reduced heart rotation and no cardiac jogging (Bakkers et al.,

2009; Smith et al., 2008). During the cell migration that initiates dextral looping events, bmp4 mutants have irregular cellular movements with a high meandering index (Bakkers et al., 2009; Smith et al., 2008). Interestingly, the addition of Bmp soaked beads directs the path of cell migration towards the Bmp signal (Bakkers et al., 2009; Smith et al., 2008).

Therefore, Bmp instructs migration by acting as a directional cue while Has2 acts to create the rotational events by affecting differential speeds of migrating cells (Bakkers et al., 2009;

Smith et al., 2008).

The downstream mechanism by which BMP and Has2 act to coordinate both cardiac jogging and dextral looping is still unknown but may require the actions of the actin cytoskeleton. Has 2 is known to stimulate Ras-related C3 botulinum toxin substrate 1 (Rac1) in cultured cells, activating lamellipodia formation and cell migration behaviors (Smith et al.,

2008) Interestingly, lamellipodial extensions are found on heart cells during cardiac jogging events and therefore, may contribute to cell migration (Smith et al., 2008). Has2 is also required for cardiac looping in mice (Bakkers et al., 2009; Camenisch et al., 2000). Further research is needed to determine other mechanisms by which Has2 and Bmp act to direct cardiac looping events and whether they interact with other members of the LR signaling pathway like Pitx2. Heart looping requires coordinated cellular rearrangements and migration likely through reorganization of the actin cytoskeleton. It is possible that the development of liver laterality also involves asymmetries in these same cellular level processes.

2.2) Asymmetric regression of blood vessels closely associated with the heart

Many of the blood vessels that connect to the heart are asymmetric in both location and function. For example, the superior and inferior vena cava return deoxygenated blood from the body to the left atrium. The pulmonary artery and veins bring blood to and from the lungs for the oxygenation process. The aorta takes freshly oxygenated blood from the right ventricle to the rest of the body. Additionally, many vessels and capillary beds must provide blood and oxygen to the heart tissue itself. The proper separation and formation of each of these vessels is vital for the proper functioning of the heart.

At the beginning of heart blood vessel formation, there are six pairs of aortic arch arteries and the aorta (Ramsdell, 2005). The first and second pairs regress into capillary beds

(Ramsdell, 2005). The third pair becomes paired common carotid arteries. However, the fourth pair exhibits asymmetries as the left side becomes part of the aortic arch while the right side contributes to the subclavian artery (Ramsdell, 2005). The fifth pair regresses while the sixth pair only regresses on the right side as the left side contributes to the pulmonary artery and truncus arteriosus (Ramsdell, 2005). Pitx2 is most likely required for the asymmetric development of these blood vessels has Pitx2 deficient mice commonly have congenital defects affecting their development including: transposition of the great arteries, anomalous venous return, persistent truncus arteriosus, and aortic arch anomalies (Ramsdell,

2005). However, the cellular and molecular mechanism by which Pitx2 confers left-right asymmetries to these blood vessels is unknown.

2.3) Gut looping

Like the heart, the midgut also loops in a stereotypical direction required for proper gastrointestinal tract function. In chick and mice, the primitive gut tube first arises during body folding events as the endoderm forms an elongated cylinder that becomes segregated into the foregut, midgut, and hindgut (Davis et al., 2008). As the midgut elongates, a hairpin loop develops in the small intestine (Davis et al., 2008). This loop eventually extends outside of the body cavity and undergoes a 90 degree counterclockwise rotation before retracting back inside the body and rotating another 180 degrees counterclockwise (Burn and Hill,

2009; Davis et al., 2008). These rotation events are required in order to fit the entire length of the intestine inside the body cavity. The LR signaling pathway is required for proper midgut looping as Nodal and Pitx2 mutants display disorganized gut coiling (Saijoh et al., 2003).

Research over the past several decades using chick, mouse, zebrafish, and Xenopus has given us a glimpse into the mechanisms required for midgut coiling.

In chicks and mice, LR asymmetries within the dorsal mesentery (DM) are responsible for initiating the leftward tilt of the midgut by providing the torque needed to drive subsequent gut rotation (Burn and Hill, 2009; Davis et al., 2008; Kurpios et al., 2008;

Welsh et al., 2014). The DM is derived from a part of the LPM called the splanchnopleural mesoderm and functions to suspend the primitive gut tube from the body wall (Burn and Hill,

2009; Davis et al., 2008; Kurpios et al., 2008; Welsh et al., 2014). The DM consists of both mesenchymal and epithelial compartments (Burn and Hill, 2009; Davis et al., 2008; Kurpios et al., 2008; Welsh et al., 2014). LR asymmetries in gene expression, epithelial cell shape, mesenchymal density, cell adhesion, ECM composition, and actomyosin contractility within

22 both compartments of the DM are downstream of Pitx2 and required for midgut looping

(Burn and Hill, 2009; Davis et al., 2008; Kurpios et al., 2008; Welsh et al., 2014).

In order to discover genes involved in LR asymmetrical development of the gastrointestinal tract and find gene targets downstream of Pitx2, Welsh et al. (2014) performed laser capture microdissection, comparing gene expression in the right and left sided compartments of the DM. Isl1, a LIM homeodomain containing transcription factor, N- cadherin (Ncad), and components of the Wnt signaling pathway, including Frizzled (Fzd) receptors Fzd4 and Fzd8, the heparin sulfate proteoglycan, Gpc3, and disheveled associated activator of morphogenesis (Daam), were found to be expressed solely on the left side

(Welsh et al., 2014). Conversely, the T-box containing transcription factor, Tbx18, and Wnt signaling inhibitors, secreted frizzled related proteins, Sfrp1 and Sfrp2, were expressed on the right side (Welsh et al., 2014). Pitx2 is required for the asymmetric expression of many of these genes as ectopic right-sided expression of Pitx2 eliminates Tbx18 expression and induces the expression of Ncad, Daam2, and Isl1 on the right side whereas Pitx2 mutants display loss of the expression of Isl1 and Daam2 (Welsh et al., 2014). Interestingly, ectopic expression of Isl1 also induces bilateral expression of Pitx2, suggesting that Isl1 functions in a positive regulatory loop with Pitx2 (Welsh et al., 2014). Genes expressed asymmetrically within the DM and genes downstream of Pitx2 are likely responsible for any LR differences in cellular level morphogenesis required for midgut looping and may also be candidates for downstream targets of Pitx2 in other organ systems. Knowledge about how Pitx2 and its downstream effectors like Daam1 and Ncad orchestrate LR asymmetry during midgut looping may give us clues to the development of laterality in other organs like the liver.

LR differences in cellular architecture downstream of the Pitx2 pathway causes the

DM to adopt a trapezoidal shape, tilting the gut tube to the left (Davis et al., 2008; Kurpios et al., 2008; Welsh et al., 2014). In the epithelial compartment, differences in cell shape contribute to this leftward tilt as cells on the left side are more columnar in morphology while cells on the right are flattened and cuboidal (Fig. 1.3A) (Davis et al., 2008; Kurpios et al.,

2008). Additionally, cells on the left side are more polarized than cells on the right, as indicated by higher levels of actin staining on the apical border and the presence of a basement membrane; right-sided actin is more diffuse and no basement membrane is apparent (Davis et al., 2008; Kurpios et al., 2008). Pitx2, Isl1, and Ncad expression are required for these cellular asymmetries as ectopic right-sided expression of any of these genes causes the right sided cells to become more columnar in shape, preventing the normal tilting of the DM (Davis et al., 2008; Kurpios et al., 2008; Plageman Jr. et al., 2011). In Pitx2 mutant mice, in which Isl1and Ncad expression is eliminated, cells on the left side become shorter and more cuboidal, similar to normal right-sided morphology (Davis et al., 2008;

Kurpios et al., 2008; Plageman Jr. et al., 2011). Ncad, expressed asymmetrically on the left side of the DM, cooperates with Shroom3 to regulate left-sided cell shape (Plageman Jr. et al., 2011). Shroom3 induces apical F-actin and Myosin II, increasing apical constriction and cell elongation on the left side (Plageman Jr. et al., 2011). Interestingly, cell shape changes occur only on the right side, (i.e., both sides of the epithelium have a columnar shape prior to the initial tilting process), indicating that Shroom3 and Ncad interactions may act downstream of Pitx2 to prevent cells from becoming cuboidal in shape through actin-myosin contractility (Davis et al., 2008).

In the mesenchymal compartment, there are no LR differences in cell shape, however; there are differences in cellular condensation. Left-sided cells are more densely packed together than right-sided cells (Fig. 1.3A) (Davis et al., 2008; Kurpios et al., 2008). Pitx2 and

Ncad are required for this difference in mesenchymal compaction as ectopic right sided expression of either gene induces cell condensation (Davis et al., 2008; Kurpios et al., 2008).

The left versus right composition of ECM proteins accounts for this difference. The left mesenchyme compartment consists of more Glycosaminoglycans (GAGs) while the right has more Hyaluronic acid (HA) (Kurpios et al., 2008). HA is a big protein that occupies a larger volume of space than GAGs, therefore, asymmetric expression of HA in the right mesenchymal compartment would cause expansion of the right side (Kurpios et al., 2008).

Downstream of Pitx2, Ncad is expressed asymmetrically within the DM on the left side. This asymmetric expression is required for LR differences in mesenchymal compaction as Ncad alters ECM production of HA and GAGs. Ectopic expression of Ncad in the right

DM induces mesenchymal cells to become more densely packed due to an increase in GAGs and a decrease in HA (Kurpios et al., 2008). Additionally, the expression of a dominant negative Ncad on the left side increases HA and decreases GAGs, causing mesenchyme cells to adopt a sparse organization (Kurpios et al., 2008). Therefore, asymmetric expression of

Ncad in the left mesenchymal compartment downstream of Pitx2 inhibits the deposition of

HA, instead providing a favorable environment for GAGs.

Daam2, a formin protein that is downstream of Pitx2 on the left side, is also required for left-sided mesenchymal condensation by facilitating the maintenance of cell to cell contacts via the formation of cadherins based junctions (Welsh et al., 2014). Formins, like

Daam2, typically polymerize unbranched F-actin to form stress fibers critical for cytoskeletal rearrangements, cell polarity, and adhesion (Welsh et al., 2014). Ectopic expression of

Daam2 on the right side of the DM induces mesenchymal condensation due to an increase in cell contacts as a result of an expansion in the length of cell junctions (Welsh et al., 2014).

Conversely, Daam2 mutants have disrupted left-sided condensation due to a loss of F-actin and abnormal intercellular adhesion as a result of a decrease in the length of cell junctions

(Welsh et al., 2014). Interestingly, Daam2 physically interacts with a cell to cell adhesion protein, -catenin, as they co-immunoprecipitate together (Welsh et al., 2014). Therefore,

Daam2 downstream of Pitx2 directs the formation and size of cadherin based junctions required for left-sided mesenchymal condensation through its effect on F-actin organization and possibly through its physical interaction with cell adhesion proteins (Welsh et al., 2014).

In the chick and mouse, asymmetries in the DM are required for midgut looping, however; in zebrafish and frog, there is no dorsal mesentery. In zebrafish, asymmetries in cell movements in the LPM orchestrate gut laterality. In this species, the gut originates from a solid rod of endodermal cells found at the ventral midline (Horne-Badovinac et al., 2003;

Yin et al., 2010). Looping of the gut occurs at 26 hours post fertilization as the gut tube curves to the left (Horne-Badovinac et al., 2003). The LPM surrounding the developing gut arise from two distinct left and right populations. Both sides of the LPM have a highly polarized epithelium, however, asymmetries exist in the morphology and position of cells

(Horne-Badovinac et al., 2003; Yin et al., 2010).

The left LPM is dorsal to the endoderm with columnar cells organized on the ventral side whereas the right LPM is ventrolateral to the endoderm with columnar cells on the

26 dorsal side (Horne-Badovinac et al., 2003; Yin et al., 2010). Before the onset of gut looping, the LPM is symmetrical, however, during looping, the right side undergoes ventrolateral migration while the left side moves dorsal to the endoderm (Fig. 1.3B) (Horne-Badovinac et al., 2003; Yin et al., 2010). Cell tracking experiments show that more ventrally located cells of the right LPM roll dorsally, moving the entire LPM on this side to a more ventrolateral position (Horne-Badovinac et al., 2003; Yin et al., 2010). Thus, the right-sided movement of the LPM actively pushes the gut leftward (Horne-Badovinac et al., 2003; Yin et al., 2010).

Gut displacement is autonomous to the LPM as bonnie and clyde mutants (bon) that lack endoderm, still have asymmetric migration of the LPM (Horne-Badovinac et al., 2003).

Interestingly, the proper polarity of cells within the LPM compartment is vital to this event as embryos with defects in nagie oko, a gene required for epithelial polarity, have defects in gut looping (Horne-Badovinac et al., 2003). In these mutants, the epithelial structure of the LPM is disrupted and the ventrolateral migration of the right LPM is perturbed as these cells migrate dorsally, similar to the normal pattern of the left LPM (Horne-Badovinac et al.,

2003). Proper epithelial polarity is likely necessary for coordinated and directional cell migration events required for gut laterality.

ECM proteins are integral to many aspects of organogenesis due to their ability to not only provide organ support but also to regulate cell to cell communication, differentiation, proliferation and migration (Yin et al., 2010). The ECM protein, Laminin, is required for asymmetric migration during zebrafish gut looping as Laminin mutants display symmetric, dorsally located LPM arrangement (Hochgreb-Hagele et al., 2013, Yin et al., 2010). Hand2, a transcription factor that regulates Laminin degradation, coordinates Laminin deposition along

27 the migratory path of rearranging cells (Yin et al., 2010). Normally, Laminin expression is degraded in the direction of cell movement; however, in Hand2 mutants, Laminin persists, preventing proper migration (Yin et al., 2010). Hand2 regulates Laminin deposition by regulating the activity of matrix metalloproteases (MMPs) (Yin et al., 2010). MMPs are proteolytic enzymes that aide in the remodeling of the ECM by cleaving ECM proteins like

Laminin (Yin et al., 2010). Hand2 mutants have reduced MMP proteolytic activity due to the increased expression of MMP inhibitors Timp2a and b (Yin et al., 2010).

Hand2 is only expressed in the ventral portion of the LPM and this expression is regulated by Bmp2 as ectopic expression of Bmp2b induces Hand2 expression throughout the LPM (Yin et al., 2010). Bmp itself is restricted to the ventral portion of the LPM due to the expression of Bmp antagonists in the somites (Yin et al., 2010). Inhibition of Bmp also eliminates Hand2 expression (Yin et al., 2010). Hand2 mutants, as well as embryos treated with Bmp inhibitors, have gut laterality defects (Yin et al., 2010). Therefore, Hand2 downstream of Bmp2b regulates cellular rearrangements by altering the activity of MMP through the expression of MMP inhibitors, creating a path of migration and subsequent cellular rearrangement by remodeling the ECM of the LPM to initiate gut looping events

(Yin et al., 2010). Gastrointestinal laterality in zebrafish is downstream of the Nodal signaling pathway as Nodal knockdowns display abnormalities in gut looping and have randomized LPM migration (Horne-Badovinac et al., 2003). However, it is unknown whether the Pitx2 pathway influences the expression of polarity genes, or BMP and Hand2 to orchestrate asymmetric migration of the LPM.

The model frog Xenopus laevis also lacks a well-defined DM region. However, midgut looping is not a consequence of LR differences in cellular migration of the LPM but rather due to differences in cellular rearrangements within the endoderm of the gut tube itself

(Muller et al., 2003). During the peak of midgut curvature, the right side of the gut tube becomes two times longer than the left, inducing a right-sided convex surface as opposed to the concave left side (Muller et al., 2003). This differential LR gut elongation is driven by

Pitx2c as ectopic expression induces midgut malrotation (Muller et al., 2003). It is likely that

Pitx2c influences cell shape and cell rearrangement patterns, ultimately orchestrating differential convergent extension and radial intercalation movements that are required for the process of gut elongation, but this is not well understood (Reed et al., 2009).

The morphological mechanism by which midgut looping is established varies between species studies. In the mouse, cell shape changes and differences in ECM deposition are required for midgut tilting (Davis et al., 2008; Kurpios et al., 2008). In the zebrafish, cellular rearrangements within the LPM surrounding the gut endoderm are required to initiate looping events (Horne-Badovinac et al., 2003; Yin et al., 2010). During midgut looping in the frog (X. laevis), cellular rearrangements intrinsic to the gut endoderm itself are necessary for LR asymmetries (Muller et al., 2003). These mechanisms are known to be downstream of

Pitx2 in the mouse and frog, however, it is unknown if Pitx2 orchestrates these cellular level asymmetries during zebrafish gut looping (Davis et al., 2008; Muller et al., 2003). It is also possible that similar cellular level mechanisms are required for liver laterality.

2.4) Vascular and Lymphatic asymmetries in the dorsal mesentery

The proper development and placement of the vascular system that supplies the gastrointestinal tract is important for providing blood supply to the intestine as well as transporting materials absorbed to the liver for detoxification. Arteries connect to the dorsal portion of the gut from the aorta while veins located ventrally drain into the hepatic portal system (Mahadevan et al., 2014). This segregation of arteries and veins is vital as defects in this process induce portosystemic shunts and metabolic imbalances as the material drained from the gut bypasses the detoxification system within the liver (Mahadevan et al., 2014).

Many of the arteries that supply the gastrointestinal tract develop within the dorsal mesentery, the tissue that suspends the gut tube from the body wall and directs intestinal rotation (Mahadevan et al., 2014). Interestingly, vascular patterning develops asymmetrically from the endothelial plexus within the DM as the forming arterial cords are present only on the left side within the densely packed mesenchyme (Mahadevan et al., 2014). Prior to intestinal rotation, the endothelial plexus is bilaterally symmetric in both the left and right mesenchymal compartments of the DM (Mahadevan et al., 2014). At the first appearance of cellular asymmetries within the DM that are required for intestinal rotation, the endothelial cells leave the right side and cross over to the left (Mahadevan et al., 2014). As the gut tilts leftward, endothelial cords are only found on the left side. Pitx2, expressed in the left mesenchyme surrounding the endothelial cords, is required for this asymmetrical regression as Pitx2 deficient mice have decreased arterial cord formation (Mahadevan et al., 2014).

Interestingly, the c-x-c motif chemokine (Cxcl12) and its receptor, chemokine receptor type 4 (Cxcr4) are also required for asymmetrical vascular development as mice

30 deficient for either gene display defective DM arteriogenesis (Mahadevan et al., 2014).

Cxcl12 is expressed asymmetrically in the left side of the dorsal mesentery in the tissue that surrounds endothelial cells while Cxc4 is expressed bilaterally within the endothelial cells themselves (Mahadevan et al., 2014). Pitx2 is required for asymmetric Cxcl12 expression as ectopic right-sided Pitx2 induces ectopic Cxcl12 expression and also causes the vasculature to development bilaterally, preventing right-sided vascular regression (Mahadevan et al.,

2014). Additionally there are binding sites for Pitx2c in the promoter region of Cxcl12

(Mahadevan et al., 2014). Asymmetric Cxcl12 expression, induced by Pitx2c, likely acts as a chemotactic agent, guiding the Cxc4+ endothelial cells to the left side (Mahadevan et al.,

2014).

Asymmetric vascular development also initiates asymmetric lymphatic development within the DM (Mahadevan et al., 2014). As the arterial cords are regressing to the left side, lymphatic precursors also move to the left (Mahadevan et al., 2014). The Pitx2-Cxcl12 pathway is required for this asymmetry as both mutations in Pitx2 and inhibition of Cxcl12 causes the lymph vessels to not develop (Mahadevan et al., 2014). Therefore, Pitx2 is required to set up the proper location of both the vasculature and the lymphatic system within the dorsal mesentery of the gastrointestinal tract (Mahadevan et al., 2014). The mechanism by which both vascular and lymphatic precursors cells migrate within the mesenchyme or how much the mechanics of this migration are also influenced by Pitx2 is unknown.

The adult liver is an extremely vascular organ as each lobule and functional unit

(portal triad) consists of hepatocytes surrounded by sinusoidal capillaries originating from the portal vein and hepatic artery (Si-Tayeb et al., 2010). The close development and

31 interplay between liver lobation and vasculature infiltration could suggests that LR asymmetries also exist during blood vessel development as they intermingle with the liver parenchyma. It is unknown if any asymmetries exist in liver associated vasculature development although portosystemic shunts and preduodenal portal veins are commonly associated with heterotaxy syndrome.

2.5) Stomach Curvature

The earliest event of LR asymmetrical morphogenesis in the gastrointestinal tract is the bulging of the foregut to the left to create the J-shaped curvature of the stomach with the longer, “greater” curvature on the left and the shorter, “lesser curvature” on the right (Davis et al., 2017; Burn and Hill, 2009). Pitx2c is expressed in the left endoderm and mesoderm of the Xenopus stomach and this expression is required for LR asymmetric stomach curvature as mutations of Pitx2c cause reversed or midline stomach orientation (Davis et al., 2017).

Research published just this year reveals downstream mechanisms of Pitx2c during stomach curvature.

In both mouse and frog, the left endodermal wall of the stomach becomes longer, expanding more than the right, causing the tissue to bend outward while also skewing the lumen leftward (Fig. 1.2C) (Davis et al., 2017). The left wall of the stomach becomes thinner and has fewer cell layers than the right (Fig. 1.2C) (Davis et al., 2017). There are no differences in proliferation or cell number, suggesting cellular rearrangement as a driving mechanism for leftward elongation (Davis et al., 2017). The left epithelial layer is also more polarized than the right, a requirement for subsequent cellular rearrangement (Davis et al.,

2017). The Nodal-Pitx2c pathway is required for differential elongation as chemical inhibition of Nodal and misexpression of Pitx2c in Xenopus causes the left wall to increase in cell layers, with abnormal architecture due to failed cellular rearrangement that causes a straightened stomach as the left wall fails to thin (Davis et al., 2017). Knock-down of Pitx2c in either the mesodermal or endodermal layer also affects adjacent, non-targeted tissue suggesting reciprocal signaling between the two compartments (Davis et al., 2017).

Interestingly, cellular mechanisms upstream of midgut looping, in the DM, occur extrinsic to the gut tube whereas both midgut looping and stomach curvature in the frog requires a mechanism intrinsic to the organ itself (Davis et al., 2008; Davis et al., 2017).

Although Pitx2 is required for both stomach curvature and intestinal looping, the cellular level morphological result is contrasting as Pitx2 drives tissue expansion in the left stomach wall and tissue condensation in the mesenchymal compartments of the dorsal mesentery of the midgut (Davis et al., 2008; Davis et al., 2017). Additionally, in the midgut of the frog, the right wall of the epithelium elongates instead of the left, opposite to events during stomach curvature (Muller et al., 2003). Therefore, the role of Pitx2 during LR asymmetric organogenesis is likely tissue specific and depends on the final geometrical shape of the organs affected. Determining how LR asymmetries are established in other organ systems, like the liver, will give us further insight into conserved and divergent mechanisms of all aspects of LR organogenesis.

2.6) Asymmetric regression of the spleen

The spleen is asymmetrically located on the left side of the body cavity. In fact, the placement of the spleen is commonly used for the diagnosis of heterotaxia. The presence of bilateral spleen indicates left-isomerism while the absence of the spleen or splenic hypoplasia indicates right isomerism. During spleen development in X. laevis, there is initially a bilateral pool of precursor cells indicated by the expression of Nkx2.5 (Burn and Hill, 2009; Patterson et al., 2000). The left sided precursors develop into the spleen while the right sided precursors adopt a different developmental fate and are incorporated into other organs due to the loss of Nkx2.5 (Burn and Hill, 2009; Patterson et al., 2000). The Nodal-Pitx2 pathway is required for this process as Nodal and Pitx2 mutants have a reduction in spleen size (i.e. right isomerism) or loss of the spleen (Burn and Hill, 2009; Patterson et al., 2000). Pitx2 most likely acts to maintain Nkx2.5 expression in left precursor cells, however, more research is needed into the cellular mechanisms of asymmetric splenic development (Burn and Hill,

2009; Patterson et al., 2000).

2.7) Lung Lobation

The lung is one of the few organs that has a symmetrical placement within the body cavity while also exhibiting asymmetric morphology. The trachea is centered along the backbone and the left and right lung lobes bud off their respective bronchi to either side.

However, like the liver, the lungs exhibit laterality in the size and number of lobes between the left and right sides as the right lung is larger, and has more lobes than the left (Gray,

1918). For example, in mice, there are four right lung lobes while the left side only has two

(Kitamura et al., 1999) while, in humans, the right lung has three lung lobes and the left lung has two (Gray, 1918). The Nodal-Pitx2 pathway is required for this LR asymmetrical morphology as Pitx2 mutant mice show lung right isomerism (i.e., both the left and right lungs have four lobes) (Gage et al., 1999; Kitamura et al., 1999; Lin et al., 1999; Liu et al.,

2001; Lu et al., 1999; Shiratori et al., 2006). However, the underlying cellular mechanism of

LR lung morphogenesis is unknown. Knowledge about the development of the liver including cellular level mechanism of differential lobation will also give us clues about the formation of lung laterality.

2.8) Liver lobation

The role of Pitx2 in LR asymmetrical organogenesis is just beginning to be elucidated in organs like the stomach and midgut, however, the role of Pitx2 in orchestrating liver laterality is unknown. The liver is LR asymmetric in both location within the body (on the right side) and morphology (the right lobe of the liver is generally larger than the left)

(Abdel-Misih and Bloomston, 2010; Gray, 1918). In humans, the right side of the liver consists of four lobes, the right medial, right lateral, quadrate, and caudate, while the left side consists of two lobes, left medial and lateral (Gray, 1918).

The liver is initially specified from a region on the ventral surface of the developing foregut (Zorn and Wells, 2009). Future hepatoblasts bud off from the endoderm, eventually intermingling with the surrounding mesoderm and endothelial cells to form the liver cords and sinusoids of the mature liver (Zorn and Wells, 2009). We know that the left and the right lobes of the liver arise from two distinct left and right cell populations that connect at the

35 ventral midline (Weiss et al., 2016), however, the mechanisms that produce left-right asymmetries in the shape and lobation patterns of the liver are not well understood.

In zebrafish, there is no LR asymmetry within the liver itself (no differences in LR lobal morphology), however, the placement of the liver with respect to the gastrointestinal tract displays laterality as the liver always buds off of the left side of the developing foregut

(Cayuso et al., 2016). Future hepatoblasts are initially symmetrically located at the embryonic midline (Cayuso et al., 2016). Shortly after initial specification, the liver bud shifts to the left side of the midline due to cell shape changes and cellular migration patterns

(Cayuso et al., 2016). During active liver bud formation and leftward outgrowth, hepatoblasts become elongated and change their orientation towards anterior-leftward outgrowth, moving in a consistent and coordinated direction, exchanging positions with their neighbors (Cayuso et al., 2016). Anteriorly positioned cells move posterior and leftward while posterior cells move anterior and leftward before a second phase of more consistent, angular displacement to the left (Cayuso et al., 2016). Cytoskeletal components like filamentous actin (F-actin) are required for this process as F-actin depolymerizing drugs cause hepatoblasts to remain at the midline (Cayuso et al., 2016). Hepatoblasts have distinct F-actin rich cellular protrusions including flat, sheet like protrusions (lamellipodia) and thin extensions (filopodia) (Cayuso et al., 2016). Lamellipoda are typically seen at the leading edge of migrating cells while filopodia act to direct the path of migration through their sensing capabilities (Mejillano et al., 2004). Interestingly, the LPM surrounding the migrating hepatoblasts also forms basal protrusions, interconnecting the tissues, suggesting LPM involvement in liver laterality

(Cayuso et al., 2016). Indeed, asymmetries are also seen in the movement of the LPM as the

36 left side moves dorsal to the underlying liver endoderm while the right moves ventrolaterally

(Cayuso et al., 2016). This suggests that signaling factors within the mesoderm and endoderm coordinate zebrafish liver laterality.

In fact, EphrinB1 in the endoderm interacts with the Ephrin receptor, Ephb3b, expressed asymmetrically in the right lateral plate mesoderm to orchestrate leftward hepatoblast migration (Cayuso et al., 2016). Ephrin B1 in the endoderm regulates hepatoblast directional migration through its interaction with EphB3b controlling the polarity and movement of the LPM (Cayuso et al., 2016). Ephb3b actually repels hepatoblasts inducing their leftward movement into the liver bud (Cayuso et al., 2016).

Cellular protrusions mediated by Eph/Ephrin signaling affect hepatoblast motility

(Cayuso et al., 2016). EphrinB1 mediates F-actin cellular extensions while Epb3b induces extension stabilization (Cayuso et al., 2016). Both EphrinB1 and Ephb3b are required for leftward liver bud migration as knockout of either gene induces laterality defects due to impaired actin protrusive activity and defects in cellular rearrangement (Cayuso et al., 2016).

Currently, there is no evidence that Ephrin or Ephrin receptors are downstream of the Nodal-

Pitx2 pathway but the EphrinB1/Ephb3b interaction provides evidence that zebrafish liver laterality depends largely on reciprocal interactions between the mesoderm and endoderm

(Cayuso et al., 2016). Additional studies are needed to determine how liver lobal morphology in other animals becomes asymmetrical (i.e. size and number of left and right lobes) and whether Pitx2 is required for this process.

2.9) Conclusions

Although the final shape and geometry of LR asymmetric organs varies widely based on the position within the body cavity and the functions required, many factors of LR asymmetric morphogenesis remain conserved. An initial symmetry breaking event during neurulation at the organizer or “node” sets up the embryonic LR axes by cilia driven leftward flow that induces the left-sided expression of Nodal (Basu and Brueckner, 2008; Blum,

2014; Ibanes and Izpisua Belmonte, 2009; Raya and Izpisua Belmonte, 2006; Shiratori et al.,

2006). Nodal expression, limited to the left LPM by the midline barrier, is vital for LR organ development (Brennan et al., 2002; Saijoh et al., 2003; Shiratori et al., 2006; Yamamoto et al., 2003). During organogenesis, a variety of morphogenetic events are responsible for laterality including LR asymmetric migration/cell rearrangement events, modifications to the actin cytoskeleton, cell shape changes, and the distribution of ECM proteins. Importantly, the homeobox transcription factor, Pitx2, downstream of the initial Nodal signal, is the driving force behind these asymmetrical events during morphogenesis in many organs and model systems studied. However, additional downstream mechanisms by which Pitx2 orchestrates these asymmetries are still relatively unknown. Additionally, the role of Pitx2 in the LR asymmetric development of many organs like the liver have yet to be elucidated.

3. Rationale and Hypothesis

As discussed in previous sections, many internal organs of the body are LR asymmetrical in anatomical location and morphological configuration including the liver.

The liver is positioned on the right side of the body cavity. Additionally, the right liver lobe

38 is larger than the left and the two lobes have different morphologies. As many as 1 in 10,000 humans are born with defects in LR asymmetry that often involve severe anomalies in liver laterality including hepatic duct and biliary tree malformations, yet the mechanism by which this asymmetry develops and the genes involved are largely unknown (Lin et al., 2000).

In other LR asymmetric organs, LR differences in cell rearrangements, cell shape changes, reorganization of the actin cytoskeleton, and redistribution of ECM proteins orchestrate laterality. For example, cellular rearrangements in the cardiac cone orchestrated by cytoskeletal lamellipodial extensions are required for cardiac jogging and dextral heart looping in zebrafish (Bakkers et al., 2009, Smith et al., 2008). During X. laevis midgut looping, asymmetries in cellular rearrangements of the gut endoderm initiate gut curvature

(Muller et al., 2013). In mice, asymmetries in ECM composition and cell shape in the dorsal mesentery create the leftward tilt required to initiate midgut rotation (Davis et al., 2008). In this context, cells on the right side of the DM become cuboidal and flattened in shape while cells on the left are more elongated and apically constricted due to apical accumulation of F- actin and Myosin II (Davis et al., 2008; Plageman Jr. et al., 2011). In zebrafish, liver morphology itself is not asymmetric but the placement of the liver is located to the left side of the midline (Cayuso et al., 2016). Cellular rearrangements driven by cell shape changes and F-actin mediated protrusions are required for future liver cells to migrate to the left side of the midgut to initiate liver bud outgrowth (Cayuso et al., 2016). In humans and many other vertebrates, the liver is asymmetric in both location within the body cavity and morphology of the left and right lobes, however, the cellular level mechanism by which this asymmetric

39 morphology develops is unknown. It is likely that mechanisms employed by other LR asymmetric organs to initiate laterality are also used for LR liver development.

Pitx2c, a homeobox transcription factor, expressed on the left side of many LR asymmetric organs, is required for proper asymmetric morphogenesis of the heart, lung, gastrointestinal tract, and stomach (Gage et al., 1999; Kitamura et al., 1999; Lin et al., 1999;

Liu et al., 2001; Lu et al., 1999; Raya and Izpisua Belmonte, 2006; Shiratori et al., 2006).

Pitx2 is required for apical F-actin and Myosin II accumulation that drives cell shape changes in the dorsal mesentery of the gut (Plagemen Jr. et al., 2011). Pitx2 is also required for cellular rearrangements during stomach curvature (Davis et al., 2017).

Pitx2c is expressed on the left side of the septum transversum mesenchyme, a mesodermal tissue that surrounds the developing liver, however, the requirement for Pitx2c in LR asymmetric liver development is unknown. I hypothesize that left-limited Pitx2c is required for LR asymmetrical development of the liver and that Pitx2c orchestrates these LR differences through asymmetries in cellular level epithelial morphogenesis including cell shape changes and cellular rearrangements.

In the model frog Xenopus laevis, the right liver lobe is larger and more elongate while the left liver lobe is small and spherical. Using the experimental amenability of

Xenopus, and a combination of chemical and molecular techniques, I tested my hypothesis with the following specific aims:

Specific Aim 1: Identify the gross morphological and cellular level LR asymmetries that arise during liver development.

Specific Aim 2: Determine the requirement for left-limited Pitx2c expression during LR asymmetrical development of the liver.

Specific Aim 3: Determine the cellular level mechanism by which Pitx2c orchestrates LR asymmetries in the liver.

My results reveal surprising contrasts between mechanisms that shape LR liver morphology and those that generate laterality in other organs like the gastrointestinal tract and stomach. I describe asymmetries in cell shape and behavior downstream of the transcription factor Pitx2c that are required for liver laterality. This is the first study to document the cellular level morphological asymmetries downstream of Pitx2c during liver development.

Figure 1- 1) LR asymmetric organ placement and morphology is vital for human health The normal positioning of the organs within the body is termed situs solitus (A). A complete reversal of the all organs within the body is called situs inversus (B). Forms of heterotaxia include right isomerism (C) and left isomerism (D) in which one or more organ systems are mirror images of either the right or the left side. Many infants are born with heterotaxia associated diseases that require intervention and surgical correction, leading to substantial hardship and morbidity.

Figure 1- 2) Four steps of initial LR symmetry breaking events during embryogenesis The establishment of LR asymmetry during embryonic development consists of four major steps. A) The initial LR symmetry breaking event at the embryonic node or LR organizer. Cilia driven leftward flow of embryonic fluid induces the left-sided expression of Nodal surrounding the node. B) This initial LR cue is transferred to the left LPM. Left-sided asymmetric expression of Nodal migrates from the perinodal region to the left LPM where Nodal expression expands anteriorly and posteriorly. C) The establishment of the midline barrier. Nodal induces the expression of Lefty, a gene required to restrict Nodal expression to the left side of the embryo by maintaining a midline barrier. D) The induction of LR asymmetric organogenesis. Nodal induces the expression of Pitx2 in the left LPM. The expression of Pitx2 is maintained during organogenesis where it surrounds the left side of organ primordia throughout development. A: anterior, P: posterior, R: right, L: left, LPM: lateral plate mesoderm.

Figure 1- 3) LR asymmetric organogenesis in the midgut and stomach During midgut looping in mice, cell shape changes in both the mesenchymal and epithelial compartments of the dorsal mesentery induce curvature. Cells in the left epithelium remain columnar in morphology while those on the right become flattened and cuboidal. In the left mesenchymal compartment, cells remain compact and close together while those in the right mesenchyme become sparse and less condensed. B) During zebrafish gut looping, asymmetric migration of the LPM pushes the gut tube to the left to initiate curvature. The cells of the right LPM undergo ventrolateral migration while the cells of the left LPM move dorsal to the endoderm. C) During asymmetric stomach development, cellular level asymmetries within the endoderm itself induce curvature. Cells on the left side become more polarized and undergo cellular rearrangements to organize into a single layered epithelium, increasing length on that side. Cells on the right side do not rearrange and remain a multilayered, unpolarized epithelium.

References

Abdel-Misih, S.R.Z., and Bloomston, M. (2010). Liver Anatomy. Surg Clin North Am. 90, 643-653.

Amand, T.R.S., Ra, J., Zhang, Y., Hu, Y., Baber, S.I., Qiu, M., and Chen, Y. (1998). Cloning and expression pattern of chicken Pitx2: a new component in the SHH signaling pathway controlling embryonic heart looping. Biochem. Biophys. Res. Commun. 247, 100- 105.

Applegate, K.E., Goske, M.J., Pierce, G., and Murphy, D. (1999) Situs-revisited: imaging of the heterotaxy syndrome. Radiographics. 19.

Bakkers, J., Verhoeven, M.C., and Abdelilah-Seyfried, S. (2009). Shaping the zebrafish heart: from left-right axis specification to epithelial tissue morphogenesis. Dev Biol. 330, 213-220.

Basu, B., and Brueckner, M. (2008). Chapter six cilia: multifunctional organelles at the center of vertebrate left–right asymmetry. Curr. Top. Dev. Biol. 85, 151-174.

Blum, M., Schweickert, A., Vick, P., Wright, C.V.E., and Danilchik, M.V. (2014). Symmetry breakage in the vertebrate embryo: When does it happen and how does it work? Dev. Biol. 393, 109-123

Brennan, J., Norris, D.P., and Robertson, E.J. (2002). Nodal activity in the node governs left-right asymmetry. Genes Dev. 16, 2339-2344.

Brueckner, M. (2007). Heterotaxia, congenital heart disease, and primary ciliary dyskinesia. Circulation 115, 2793-2795.

Burn, S.F., and Hill, R.E. (2009). Left-right asymmetry in gut development: what happens next? Bioessays 10, 1026-1037.

Burton, E.C., Olson, M., and Rooper, L. (2014). Defects in laterality with emphasis on heterotaxy syndrome with asplenia and polysplenia: an autopsy case series at a single institution. Pediatr. Dev. Pathol. 17, 250-264.

Camensich, T.D., Spicer, A.P., Brehm-Gibson, T., Biesterfeldt, J., Augustine, M.L., Calabro Jr., A., Kubalak, S., Klewer, S.E., and McDonald, J.A. (2000). Disruption of hyaluronan synthase-2 abrogates normal cardiac morphogenesis and hyaluronan-mediated transformation of epithelium to mesenchyme. J. Clin. Invest. 106, 349-360.

Cayuso, J., Dzementsei, A., Fischer, J.C., Karemore, G., Caviglia, S., Bartholdson, J., Wright, G.J., and Ober, E.A. (2016). EphrinB1/EphB3b coordinate bidirectional epithelial- mesenchymal interactions controlling liver morphogenesis and laterality. Dev. Cell. 39, 316- 328.

Cox, C.J., Espinoza, H.M., McWilliams, B., Chappell, K., Morton, L., Hjalt, T.A., Semina, E.V., and Amendt, B.A. (2002). Differential regulation of gene expression by Pitx2 isoforms. Journal of Biological Chemistry. 277, 25001-25010.

Davis, A., Amin, N.M., Johnson, C., Bagley, K., Ghashghaei, T., and Nascone-Yoder, N. (2017) Stomach curvature is generated by left-right asymmetric gut morphogenesis. Development.

Davis, N.M., Kurpios, N.A., Sun, X., Gros, J., Martin, J.F., and Tabin, C.J. (2008). The chirality of gut rotation derives from left-right asymmetric changes in the architecture of the dorsal mesentery. Developmental cell 15, 134-145.

Duboc, V., Rottinger, E., Lapraz, F., Besnardeau, L., and Lepage, T. (2005). Left-right asymmetry in the sea urchin embryo is regulated by nodal signaling on the right side. Dev. Cell. 9, 147-58.

Gage, P.J., Suh, H., and Camper, SA. (1999). Dosage requirement of Pitx2 for development in multiple organs. Development. 126, 4643-4651.

Gormley, J.P., and Nascone-Yoder, N.M. (2003). Left and right contributions to the Xenopus heart: implications for asymmetric morphogenesis. Dev. Genes Evol. 213, 390-398.

Gottschalk, I., Stressig, R., Ritgen, J., Herberg, U., Breuer, J., Vorndamme, A., Strizek, B., Willruth, A., Geipel, A., Gembruch, U., and Berg, C. (2016). Extracardiac anomalies in prenatally diagnosed heterotaxy syndrome. Ultrasound Obstet. Gynecol. 47, 443-449.

Grande, C., and Patel, N.H. (2009). Nodal signaling is involved in left-right asymmetry in snails. Nature. 457, 1007-1011.

Gray, H. (1918). Anatomy of the Human Body. Philadelphia: Lea & Febiger, Bartleby.com, 2000. www.bartleby.com/107/.

Hjalt, T.A., and Semina, E.V. (2005). Current molecular understanding of Axenfeld-Rieger Syndrome. Expert Rev. Mol. Med. 7, 1-17.

Hochgreb-Hagele, T., Yin, C., Koo, D.E., Bronner, M.E., and Stainier, D.Y. (2013). Laminin 1a controls distinct steps during the establishment of digestive organ laterality. Development. 140, 2734-2735.

Horne-Badovinac, S., Rebagliati, M., and Stainier, D.Y.R. (2003). A cellular framework for gut-looping morphogenesis in zebrafish. Science. 302, 662-665.

Hu, N., Sedmera, D., Yost, J.H., and Clark, E.B. (2000). Structure and function of the developing zebrafish heart. Anat. Rec. 260, 148-157.

Ibanes, M., and Izpisua Belmonte, J.C. (2009). Left-right axis determination. Wiley Interdiscip. Rev. Syst. Biol. Med. 1, 210-219.

Kerkeni, Y., Ksia, A., Zitouni, H., Belghith, M., Lassad, S., Krichene, I., Mekki, M., and Nouri, A. (2015). Biliary atresia and polysplenia syndrome. Tunis. Med. 93, 494-496.

Kitamura, K., Miura, H., Miyagawa-Tomita, S., Yanazawa, M., Katoh-Fukui, Y., Suzuki, R., Ohuchi, H., Suehiro, A., Motegi, Y., Nakahara, Y., Kondo, S., and Yokoyama, M. (1999). Mouse Pitx2 deficiency leads to anomalies of the ventral body wall, heart, extra- and periocular mesoderm and right pulmonary isomerism. Development 126, 5749-5758.

Kurpios, N.A., Ibañes, M., Davis, N.M., Lui, W., Katz, T., Martin, J.F., Belmonte, J.C.I., and Tabin, C.J. (2008). The direction of gut looping is established by changes in the extracellular matrix and in cell: cell adhesion. Proc Natl Acad Sci U S A. 105, 8499-8506.

Lee, J.M., Lee, H.S., and Kim, C.D. (2014). Situs ambiguous with left-sided isomerism. Digestive and Liver Disease. 46, 658.

Lin, A.E., Ticho, B.S., Houde, K., Westgate, M.N., and Holmes, L.B. (2000). Heterotaxy: associated conditions and hospital-based prevalence in newborns. Genet. Med. 2, 157-172.

Lin, C.R., Kioussi, C., O'Connell, S., Briata, P., Szeto, D., and Liu, F. (1999). Pitx2 regulates lung asymmetry, cardiac positioning and pituitary and tooth morphogenesis. Nature 401, 279-282.

Liu, C., Liu, W., Lu, M., Brown, N.A., and Martin, J.F. (2001). Regulation of left-right asymmetry by thresholds of Pitx2c activity. Development 128, 2039-2048.

Liu, W., Selever, J., Lu, M., and Martin, J.F. (2003). Genetic dissection of Pitx2 in craniofacial development uncovers new functions in branchial arch morphogenesis, late aspects of tooth morphogenesis and cell migration. Development. 130, 6375-6385.

Lu, M.F., Pressman, C., Dyer, R., Johnson, R.L., and Martin, J.F. (1999). Function of Riegar syndrome gene in left-right asymmetry and craniofacial development. Nature. 401, 276-278.

Mahadevan, A., Welsh, I.C., Sivakumar, A., Gludish, D.W., Shilvock, A.R., Noden, D.M., Huss, D., Lansford, R., Kurpios, N.A. (2014). The left-right Pitx2 pathway drives organ-specific arterial and lymphatic development in the intestine. Dev. Cell. 31, 690-706.

Manner, J. (2009). The anatomy of cardiac looping: a step towards the understanding of the morphogenesis of several forms of congenital cardiac malformations. Clin. Anat. 22, 21-25.

Mejillano, M.R., Kojima, S., Applewhite, D.A., Gertler, F.B., Scitkina, T.M., and Borisy, G.G. (2004). Lamellipodial versus filopodial mode of the actin nanomachinery: pivotal role of the filament barbed end. Cell. 118, 363-373.

Muller, J.K., Prather, D.R., and Nascone-Yoder, N.M. (2003). Left-right asymmetric morphogenesis in the Xenopus digestive system. Dev Dyn. 228, 672-682.

Newman, B., Feinstein, J.A., Cohen, R.A., Feingold, B., Kreutzer, J., Patel, H., and Chan, F.P. (2010). Congenital extrahepatic portosystemic shunt associated with heterotaxy and polysplenia. Pediatr. Radiol. 40, 1222-1230.

Okumura, T., Utsuno, H., Kuroda, J., Gittenberger, E., Takahiro, A., and Matsuno, K. (2008). The development and evolution of left-right asymmetry in vertebrates: lessons from drosophila and snails. Dev. Dyn. 237, 3497-3515.

Pathak, D., and Sarin, Y.K. (2006). Congenital duodenal obstruction due to a preduodenal portal vein. Indian J. Pediatr. 73, 423-435.

Patterson, K.D., Drysdale, T.A., and Krieg, P.A. (2000). Embryonic origins of spleen asymmetry. Development. 127, 167-175.

Plageman Jr., T.F., Zacharias, A.L., Gage, P.J., and Lang, R.A. (2011). Shroom3 and a Pitx2-N-cadherin pathway function cooperatively to generate asymmetric cell shape changes during gut morphogenesis. Dev. Biol. 357, 227-234.

Ramsdell, A.F. (2005). Left-right asymmetry and congenital cardiac defects: getting to the heart of the matter in vertebrate left-right axis determination. Dev. Biol. 288, 1-20.

Raya, A., and Izpisua Belmonte, J.C. (2006). Left-right asymmetry in the vertebrate embryo: from early information to higher-level integration. Nature Reviews Genetic. 7, 283- 293.

Reed, R.A., Womble, M.A., Dush, M.K., Tull, R.R., Bloom, S.K., Morckel, A.R., Devlin, E.W., and Nascone-Yoder, N.M. (2009). Morphogenesis of the primitive gut tube is generated by Rho/ROCK/myosin II-mediated endoderm rearrangements. Dev Dyn. 238, 3111-3125.

Saijoh, Y., Oki, S., Ohishi, S., and Hamada, H. (2003). Left-right patterning of the mouse lateral plate requires nodal produced in the node. Dev. Biol. 256, 160-172.

Schweickert, A., Campione, M., Steinbeisser, H., and Blum M. (2000). Mech. Dev. 90, 41-51.

Shiratori, H., Yashiro, K., Shen, M.M., and Hamada, H. (2006). Conserved regulation and role of Pitx2 in situs-specific morphogenesis of visceral organs. Development 133, 3015- 3025.

Si-Tayeb, K., Lemaigre, F.P., and Duncan, S.A. (2010). Organogenesis and development of the liver. Dev. Cell. 18, 175-189.

Smith, K.A., Chocron, S., von der Hardt, S., de Pater, E., Soufan, A., Bussmann, J., Schulte-Merker, M., Hammerschmidt, M., and Bakkers, J. (2008). Rotation and asymmetric development of the zebrafish heart requires directed migration of cardiac progenitor cells. Dev. Cell. 14, 287-297.

Wagner, M.K., and Yost, H.J. (2000). Left-right development: the roles of nodal cilia. Curr. Biol. 10, 149-151.

Weiss, M.C., Le Garrec, J.F., Coqueran, S., Strick-Marchand, H., and Buckingham, M. (2016). Progressive developmental restriction, acquisition of left-right identity and cell growth behavior during lobe formation in mouse liver development. Development. 143, 1149-1159.

Welsh, I.C., Thomsen, M., Gludish, D.W., Alfonso-Parra, C., Martin, J.F., and Kurpios, N.A. (2014). Integration of L-R Pitx2 transcription and Wnt signaling drives asymmetries gut morphogenesis via Daam2. Dev. Cell 26, 629-644.

Yamamoto, M., Mine, N., Mochida, K., Sakai, Y., Saijoh, Y., and Meno, C. (2003). Nodal signaling induces the midline barrier by activating Nodal expression in the lateral plate. Development 130, 1795-1804.

Yin, C., Kikuchi, K., Hochgreb, T., Poss, K., and Stainier, D.Y.R. (2010). Hand2 regulated extracellular matrix remodeling essential for gut-looping morphogenesis in zebrafish. Dev. Cell. 18, 973-984.

Zorn, A.M., and Wells, J.M. (2009). Vertebrate Endoderm Development and Organ Formation. Annu. Rev. Cell Dev. Biol. 25, 221-251.

CHAPTER 2: The left-right asymmetry of liver lobation is generated by Pitx2c- mediated asymmetries in the hepatic diverticulum

Mandy Womble1, Nirav Amin1, Adam Davis†, and Nanette Nascone-Yoder1 1 Department of Molecular Biomedical Sciences, College of Veterinary Medicine, North Carolina State University, Raleigh, NC 27607. † Current: Department of Biology and Physical Sciences, Gordon State College, Barnesville, GA 30204

Abstract: Both the anatomical location and morphology of vertebrate internal organs are left- right (LR) asymmetric including the heart, gastrointestinal tract, and liver. Organs that undergo branching morphogenesis, such as the lungs and liver, exhibit differential lobation on L vs R sides, but almost nothing is known as to how these asymmetries develop. Defects in liver laterality are manifest as abnormal organ position, aberrant lobe morphology and/or extrahepatic biliary tree malformations, yet the morphogenetic mechanisms underlying the development of hepatobiliary asymmetries are largely unknown. Here we describe the cellular and molecular events that generate asymmetrical liver lobation, using the experimentally amenable embryos of the frog, Xenopus laevis. We found that, during early liver morphogenesis, endoderm cells on the right side of the hepatic diverticulum become taller and more apically constricted while cells on the left side become rounder and stratified. Consequently, right side cells form an epithelial tissue with fewer layers that expands to occupy a greater surface area and, ultimately, forms a larger, more elongated lobe. Pharmacological inhibition of Nodal, or side-specific misexpression of Pitx2c, leads to anomalies in both epithelial architecture and final lobe morphology, indicating that these early morphogenetic asymmetries are specified by conserved LR patterning cues. Our results shed new light on the diversity of mechanisms that shape different types of LR asymmetries in different organs.

Introduction

Numerous internal organs acquire a left-right (LR) asymmetric anatomical position during development (e.g., spleen) or exhibit chiral rotation (e.g. intestine); however, many organs also manifest with an asymmetric morphology, such as distinct left and right atrial chambers of the heart. In branching organs, e.g., lungs and liver, asymmetry is manifest as differential size/lobation of left and right counterparts. How these distinct types of asymmetries are formed, and whether they use similar mechanisms is unknown. In most vertebrates, the liver becomes positioned in the upper right quadrant of the visceral cavity where the right side of the organ grows to be 5-6 times larger than the left, with a distinct morphology and more complex lobation pattern (Abdel-Misih and Bloomston, 2010; Gray, 1918). In individuals with defective LR asymmetry, termed heterotaxy (Lin et al., 2000), liver asymmetry is often disrupted. For example, the liver may be located at the midline (Brueckner, 2007; Shiraishi and Ichikawa, 2012), and liver lobation may be ambiguous or indeterminate (Burton et al., 2014). Both human heterotaxy patients and mouse models of heterotaxy (e.g., INV mutant) (Mazziotti et al., 1999; Shimadera et al., 2007) also exhibit defects/discontinuities in the biliary tree, including extrahepatic biliary atresia (EHBA), a common cause of liver transplantation (Gottschalk et al., 2016; Zhu et al., 2005). Understanding the cellular and molecular events that underlie the development of liver asymmetry may be vital for ascertaining the etiology of EHBA and other laterality- related congenital birth defects. Yet, the morphogenetic events which generate liver asymmetries remain almost entirely unknown. In many vertebrate embryos, LR asymmetry is initiated by cilia driven left-ward flow of extraembryonic fluid within the LR organizer or node, establishing LR asymmetric expression of Nodal, a TGF growth factor, in the left lateral plate mesoderm (LPM) (Basu and Brueckner, 2008; Blum et al., 2014; Brennan et al., 2002; Kawasumi et al., 2011; Yoshiba and Hamada, 2014). Nodal signaling then activates the transcription of the homeobox transcription factor, Pitx2c (Blum et al., 2014; Faucourt et al., 2001; Kawasumi et al., 2011; Raya et al., 2006). Both Nodal and Pitx2c activity are required for proper organ

52 laterality (Campione et al., 1999; Gage et al., 1999; Kitamura et al., 1999; Lin et al., 1999; Liu et al., 2001; Lu et al., 1999; Shiratori and Hamada, 2006) but Pitx2c expression is retained on the left side of developing organs and has been shown to cause specific changes in cell properties and behaviors that ultimately generate tissue-level asymmetries. For example, Pitx2 expression in the left side of the stomach produces changes in cell polarity and rearrangement that drive leftward curvature (Davis et al., 2017), while Pitx2 expression on the left side of the dorsal mesentery elicits changes in cell adhesion and tissue condensation that initiate chiral rotation of the attached midgut (Davis et al., 2008; Kurpios et al., 2008). Pitx2c has been found to be expressed in the left side of the LPM-derived septum transversum mesenchyme (STM) surrounding the developing liver (Shiratori and Hamada et al., 2006), suggesting that it could influence liver laterality, but the specific role of Pitx2c in the development of hepatobiliary asymmetry has not been investigated. Here we describe the differences in cell shape and behavior in the left versus right sides of the developing liver that underlie the LR asymmetric lobation of the mature organ. Using the highly accessible embryos of the frog, Xenopus laevis, that allow pharmacological induction of heterotaxy, as well as side- and tissue-specific modulations of gene function, we show that these liver-specific asymmetries are downstream of conserved LR patterning events. Our results reveal surprising contrasts between morphogenetic mechanisms that generate liver laterality and the events that shape asymmetry in other organs.

Materials and Methods

Animals

All animals were used in accordance with the IACUC regulations of NCSU. X. laevis in vitro fertilization, rearing, and staging were as previously described (Nieuwkoop and Faber, 1995; Sive et al., 1998)

Immunohistochemistry

Embryos were fixed by washing with Dents (80% methanol/20% DMSO) 8 times before overnight storage at -20C, and then processed for cryo-sectioning as previously described (Dush et al., 2013). Sections were post-fixed with 4% PFA [100 mM Hepes (pH 7.4), 100 mM NaCl, 4% paraformaldehyde] for 2 min, washed with PBT, and blocked for 1 hour as previously described (Reed et al., 2009). Immunohistochemical staining was performed overnight at 4C with blocking buffer containing the following primary antibodies: anti-integrin (Developmental Studies Hybridoma Bank (DSHB), created by the NICHD of the NIH and maintained at The University of Iowa; 8C8, 1:1000), anti-mCherry (Clontech, 632543; 1:1000), phospho-histone H3 (Millipore, 06- 570; 1:500), or GFP (ThermoFisher, 6455 1:1000). Slides were washed with PBT and secondary antibody staining was performed as described (Davis et al., 2017; Dush et al., 2013). Slides were again washed with PBT and PBS before staining with Topro-3 (ThermoFisher, T3604, 1:1000) in PBS for 5 min. at RT. Slides were washed with PBS and mounted with Prolong Gold (ThermoFisher, P36930), cured overnight in the dark, and visualized with a Leica DM 2500 confocal microscope. At least 3 embryos for each stage and experimental condition were analyzed.

RNA in situ hybridization

Whole embryos or dissected foreguts were fixed in MEMFA for 3 hours, or PFA overnight, at room temperature (RT) before being gradually dehydrated in methanol and stored at -20C. Digoxigenin (DIG)-labeled riboprobes were synthesized from linearized plasmids of coding regions of X. laevis hhex and fibrinogen as described (Lipscomb et al., 2006). In situ hybridizations (ISH) were performed as previously described (Dush et al., 2013). ISH on tissue sections was performed as previously described (Butler et al., 2001) with the following modifications. Following rehydration of slides, sections were refixed with MEMFA for 30 minutes. After probe hybridization overnight, slides were washed with SSPE before being treated with 20ug/mL Rnase A in 4X SSPE for 30 min. at 37C. Slides were

54 blocked with 0.5% blocking reagent (Roche; 11096176001) in Buffer 1 for 2 hours at RT before incubating with anti-DIG-AP Fab fragments (Roche; 11093274910, 1:5000) overnight at 4C. At the conclusion of the color reaction, slides were fixed for 15 min with 4% PFA before being washed with PBT, dehydrated to Xylene, and mounted with Permount and a coverslip.

X. laevis Loss- and Gain-of-function experiments

Dosing of X. laevis embryos from neurula stage (NF 19/20) through NF 32 with SB505124 was as previously described (Dush et al., 2011; Davis et al.; 2017). SB505124 at a concentration of 12.5 m or the same volume of DMSO (solvent control) was added to embryos in culture medium (0.1X MMR). Microinjection manipulation of Pitx2 activity using CRISPR Cas9, morpholino, and hormone-inducible synthetic mRNA was performed as previously described (Davis et al., 2017). Embryos were anesthetized and fixed at NF 35 and 46 for phenotyping and fixed at NF 35 for immunohistochemical analysis.

Measurements and Statistics

All Pitx2 perturbation experiments were performed at least in triplicate. When quantifying gross phenotypes (e.g., the length to width ratio of the right and left liver lobes), at least 10 livers were evaluated for each condition. To evaluate the LR ratio of hhex expression, between 7 and 23 embryos were measured. All morphometric measurements were made using Image J software (NIH); for each parameter, at least 3 different sections were measured in at least 3 different embryos. Statistical significance was analyzed by one-way Anova with post-hoc Tukey HSD.

Results

The liver diverticulum is left-right asymmetric In all vertebrates, the liver originates as a bud from the ventral foregut, which forms the hepatic diverticulum (Bort et al., 2006; Tremblay and Zaret, 2005; Zorn and Wells, 2009). In humans, the original diverticulum has been reported to bifurcate into left and right buds almost immediately (Heisler, 1907). In mammals, the endoderm of each liver bud delaminates and migrates into the surrounding septum transversum mesenchyme, where it forms liver cords and sinusoids, accelerating proliferation and growth and becoming highly vascularized, creating a complex branching morphology that is critical to the proper function of the liver (Bort et al., 2006; Hunter et al., 2007; Kaestner, 2005; Margagliotti et al., 2008; Tremblay, 2011; Tremblay and Zaret, 2005; Zorn and Wells, 2009). In humans, differences in the size of the right and left lobes are not reported before the 4th month (Heisler, 1907). In other vertebrates, it is unknown at which phase of liver development LR asymmetries arise. To determine when LR asymmetry is first established during liver morphogenesis, we characterized the size and shape of the developing liver at multiple stages in X. laevis (Fig. 1A-I). Surprisingly, morphometric measurements of the area occupied by the right and left halves of the liver diverticulum (relative to the ventral midline and position of the forming gall bladder) revealed that, by NF 33 (p<.0001; Fig. 1B,K), the right side of the organ is already larger in area than the left. During subsequent stages of development, the nascent right lobe of the liver continues to enlarge while the left lobe remains small (Fig. 1C-I). By tadpole stages (e.g., 42), the right lobe is visibly larger and elongated, with a significantly larger length to width ratio than the left lobe, which is more spherical (p<.0001; Fig. 1J). These results demonstrate that the morphological asymmetry of the liver arises early in development, initiating with the unequal expansion of the right side of the hepatic diverticulum itself.

Cellular left-right asymmetries in the early liver diverticulum To begin to ascertain the morphogenetic differences that underlie the differential growth of the contralateral sides of the liver, we first defined the distribution of left versus right cells during Xenopus liver development. In these experiments, membrane gfp (mgfp) and membrane cherry (mcherry) was injected into the left and right vegetal dorsal cells of the blastula to label progeny that will contribute to each side of the prospective foregut. Cells from the right or left side of the embryo were found to contribute to the right or left lobe of the mature liver, respectively (Supp. Fig.1A, D), confirming that each side remains separated and executes distinct developmental programs, as previously shown in mouse (Weiss et al., 2016). Interestingly, we found that, earlier in development when morphological asymmetry is becoming evident in the early diverticulum (e.g., stage 35), right-labelled hepatic endoderm cells occupy a broader region of the diverticulum than left labelled cells, with a significantly greater frontal and cross-sectional surface area (Figure 2J; Supp. Fig. 1B-C), suggesting that cell number or proliferation may differ on the contralateral sides of the early diverticulum. However, we observed no LR difference in total cell number, proliferation, (Sup. Fig.2), or cell death (data not shown) during the emergence of morphological asymmetry, suggesting that the observed differences in the distribution of right side cells are more likely due to LR differences in morphogenetic remodeling. To explore this idea further, we compared tissue architecture in the left and right hepatic endoderm. Prior to asymmetry (NF 32), the entire diverticulum is composed of a pseudostratified epithelium (Fig. 2A), with no significant LR differences in cell shape (Fig. 2G-H) or in the number of cell layers along the apical-basal axis (Fig. 2I). However, we observed that, by NF 33 (Fig. 2B, C), the cells in the right side of the diverticulum possess a higher length-to-width ratio (Fig. 2E, H) and become apically constricted (Fig. 2 E, G), cell properties often associated with the initiation of invagination, tube elongation and/or branching morphogenesis in other foregut organs (e.g., pancreas, Drosophila trachea, chick lung) (Sawyer et al., 2010; Varner and Nelson, 2014). In contrast, the hepatic endoderm cells on the left side of the diverticulum become rounded in shape (Fig. 2F, G-H).

Subsequently (NF 37), the left side of the diverticulum forms a compact, stratified epithelial bud, while the right side forms a pseudo-stratified epithelium comprising a greater cross-sectional surface area with nascent involutions and folds (Fig. 2D,I). The appearance of these differences correlates closely with the observed gross anatomical differences between the left and right lobes (see Fig. 1). Thus, the differential morphology of the left and right liver lobes can be attributed to differences in hepatic endoderm morphogenesis in the left versus right sides of the early liver diverticulum.

Liver asymmetry is under the control of the left-right asymmetry pathway The Nodal signaling pathway is required for LR asymmetry of multiple organ systems (Campione et al., 1999; Gage et al., 1999; Kitamura et al., 1999; Lin et al., 1999; Liu et al., 2001; Lu et al., 1999; Shiratori and Hamada, 2006). To determine if this critical LR- determining pathway directs liver asymmetry, we exposed Xenopus embryos to a small molecule chemical inhibitor of TGFsignaling, SB505124, at stages known to induce heterotaxy (Dush et al 2011; Davis et al 2017). In control DMSO treated embryos, the heart and gut show normal laterality (Fig. 3A) and the liver is asymmetric, with the right lobe larger and more elongate than the left (Fig. 3B, Fig 4G). In contrast, in embryos treated with the nodal inhibitor, the heart and gut are heterotaxic (Fig. 3C,E) and the livers exhibit a right- isomerism phenotype, where the left lobe of the liver is now larger and more elongate, similar to typical right-sided morphology (Fig. 3D,F; Fig. 4G). Thus, LR asymmetries in liver lobe morphology and size are downstream of conserved LR patterning cues.

Pitx2c is required for left lobe morphogenesis At stages 33-39, pitx2c is expressed asymmetrically in the mesoderm surrounding the left side of the liver diverticulum (Supp. Fig. 3). To determine the requirement for pitx2c in liver morphogenesis, we diminished pitx2c activity via CRISPR-Cas9 genome editing (Supp. Fig. 4A, D, G) (see Materials and Methods, Davis et al 2017). While control embryos (Fig. 4A-B) showed normal liver morphology, in which the right liver lobe is larger and more elongate than the left (p<.0001) (Fig. 4H), the livers of F0 embryos with CRISPR-mediated

58 pitx2c mutations (see Davis et al., 2017 for sequence analysis of indels) display right isomerism (32.8%; n=61, Fig. 4G), i.e., the left lobe of the liver is larger and more elongate, similar to normal right sided morphology (Fig. 4C,E). To determine the side- and tissue-specific role of Pitx2c in liver morphogenesis, we employed a pitx2c morpholino (Supp. Fig. 4B, E, H), previously validated to efficiently knock down Pitx2c activity (Davis et al., 2017), targeted to left side of liver, and chose embryos primarily limited to injections of the surrounding mesoderm. In these experiments, the left liver lobes of pitx2c-MO injected embryos were found to have a significantly larger length to width ratio than embryos injected with the control morpholino (p=.002) (Fig. 4D, F, H). These results suggest that left-sided Pitx2c expression is required for normal left side liver lobation by repressing the tissue expansion normally seen in the right lobe. We next sought to define the influence of Pitx2 on the morphogenesis of hepatic endoderm. Transverse sections through embryos injected to target the left side of the diverticulum with a control morpholino (COMO) showed normal LR asymmetries in hepatic endoderm cell shape and the number of apical to basal nuclei layers at stage 35 (p<.0001) (Fig. 5A,C,E,G-I). However, injection of a pitx2c MO caused left hepatic endoderm cells to become more apically constricted and elongated with a decrease in the number of apical to basal nuclei layers, similar to right side cellular morphology (Fig. 5B,D,F, G-I). These results suggest that Pitx2c is required to inhibit right-side morphogenetic events and allow left-side hepatic endoderm to from a stratified epithelial architecture in order to maintain a more compact lobal morphology.

Ectopic R-sided Pitx2c induces heterotaxy To determine whether Pitx2c is sufficient to inhibit right side liver lobation, and/or induce left-sided lobe morphology, we ectopically expressed Pitx2c on the right side of the developing liver diverticulum using an mRNA construct encoding a glucocorticoid-inducible variant of Pitx2c (Pitx2cGR)(Supp. Fig. 4C,F,I)(Chung et al., 2011, Davis et al 2017). In the presence of dexamethasone (see Materials and Methods), ectopic Pitx2 translocates into the nucleus to become active. Uninduced control embryos (-Dex; Fig.6A) had normal LR

59 asymmetry where the right lobe is larger and more elongate with a significantly larger length to width ratio than the left (p<.0001) (Fig. 6E). However, following dexamethasone induction, ectopic right-sided Pitx2c expression (+Dex) (Fig. 6B-C) induced left-isomerism where the right liver lobe was smaller and more spherical with a reduced length to width ratio, similar to the normal left-lobe morphology (Fig. 6D, E). Moreover, ectopic right-sided Pitx2c expression decreases surface area as the right side diverticulum length is the same as the left side (Supp. Fig. 5A). Therefore, Pitx2c is sufficient to induce left-sided liver lobe morphology.

Pitx2c is sufficient to induce left-sided cell character To determine the effect of ectopic Pitx2c on cell properties and behavior, we performed immunohistochemistry on transverse cross sections of embryos with ectopic right- sided Pitx2c expression limited to the right mesoderm (Fig.7B). Control embryos (st 35)(Fig. 7A) exhibit LR asymmetries in the number of apical to basal nuclei layers (Fig. 7K) and cell shape, with taller, apically constricted right side cells (p<.0001) (Fig. 7C,E,G, I-J). In contrast, right side cells adjacent to ectopic Pitx2c activity become less apically constricted and more rounded, similar to normal left-side morphology (Fig. 7F, H). In fact, in these embryos, there is no significant difference between the left and right cell shapes (Fig. 7I-J). Concomitantly, the number of cells layers within the right epithelium increases in the presence of ectopic Pitx2c (Fig. 7K). Taken together, our results suggest that Pitx2c expression is sufficient to alter cell shape and rearrangement patterns within the liver diverticulum that inhibit apical constriction and expansion, resulting in differential organ lobation.

Discussion

Pitx2c is necessary and sufficient to induce left-sided epithelial characteristics

The findings presented here provide the first molecular and cellular model of the morphogenetic mechanisms that generate asymmetrical liver lobation (Fig 8). At the beginning of liver morphogenesis (NF 30, Fig 8), the diverticulum is symmetrically composed of a single-layered columnar epithelium (Bort et al., 2006; Zorn and Wells, 2009). However, as the diverticulum begins to enlarge, endoderm cells on the left side become rounder and form a compact, multilayered tissue architecture while the contralateral cells on the right continue to contribute to an expanding pseudostratified epithelium (NF 33-35, Fig. 8). Consequently, the cross-sectional surface area of the left side of the liver diverticulum remains unchanged while the right side expands and begins to form folds that portend the future liver cords (NF 37, Fig. 8). This early difference in diverticular architecture precedes and correlates with the ultimate formation of an enlarged right organ lobe compared to the left lobe (NF 46, Fig. 8). Our data indicate that asymmetric expression of Pitx2c is required for diverticular asymmetry, as left-sided knockdown of Pitx2c causes cells in the left diverticulum to become more elongated and apically constricted, with expanded tissue area, ultimately resulting in a liver with a large and elongate left lobe resembling normal right lobe morphology (i.e., right isomerism). Conversely, ectopic right-sided Pitx2c expression induces cells in the right diverticulum to become rounder and form a more compact, multilayered tissue architecture, ultimately resulting in an organ with a small and spherical right lobe that resembles normal left lobe morphology (i.e., left isomerism). Taken together, these results suggest that left- sided Pitx2c expression orchestrates the cellular events that underlie left-right asymmetrical liver lobation.

Pitx2c’s influence on cell behavior is organ specific

The cellular events influenced by Pitx2 in the liver contrast with the changes induced by Pitx2 in other regions of the gastrointestinal tract. For example, LR differences in the architecture of the dorsal mesentery (DM) asymmetrically distort the attachment of the gut tube to the body wall, thus biasing the directionality of intestinal rotation. Pitx2c expression within the left side of dorsal mesentery has been found to elicit specific cellular changes in this tissue, including cell elongation, apical constriction, and polarization (Davis et al., 2008; Kurpios et al., 2008). Strikingly, these changes are opposite to those that we see in the liver diverticulum, where Pitx2 expression inhibits apical constriction and cell elongation. In the stomach, LR asymmetries in cell properties and behaviors underlie the lengthening of the left side of the organ compared to the right, resulting in the formation of its greater curvature (Davis et al., 2017). Pitx2 has been found to orchestrate this process by eliciting changes in the apicobasal polarity and rearrangements of endoderm cells that thin the layers of the left stomach wall, resulting in overall tissue expansion. However, in the liver diverticulum, Pitx2 expression causes endoderm cells to become rounder and form a more compact, multilayered tissue architecture. In both the mesentery and the liver, Pitx2 expression elicits tissue condensation. However, in the stomach, Pitx2 expression engenders tissue extension. Thus, it appears that the morphogenetic mechanisms that elicit different types of morphological asymmetries are organ-dependent. The readout of asymmetry in each organ may depend, in part, on the expression patterns of Pitx2. For example, Pitx2 is expressed throughout the left side of the dorsal mesentery (both epithelial and mesenchymal layers/zones). Likewise, Pitx2c is expressed in both the endoderm and mesoderm of the stomach (Davis et al., 2017). There is evidence that there is interaction between tissue layers in this context, suggesting that Pitx2 acts both autonomously and non-cell-autonomously to orchestrate the formation of morphological asymmetry (Davis et al., 2017). In contrast to mesentery and stomach, Pitx2 is only expressed in the mesoderm layer surrounding the liver diverticulum and therefore cannot directly influence cell shape in the underlying diverticular endoderm.

Therefore, it is tempting to speculate that mesodermal Pitx2c activates the transcription of a diffusible ligand to influence the development of the underlying endoderm in the left side of the diverticulum. For example, hepatocyte growth factor (HGF) is required for liver development and promotes tubulogenesis of the salivary and mammary glands (Barros et al., 1995; Bryant and Mostov, 2008; Ikari et al., 2003; Zhang and Vande Woude, 2003). Additionally, there are Pitx2 binding sites in the promoter region of HGF (Eng et al., 2010). It is possible that Pitx2c represses the transcription of HGF in the mesoderm of the left side of the liver diverticulum, preventing the formation of elongated branches and liver sinusoids in this lobe. Although the zebrafish liver does not develop differential lobation, reciprocal Eph- Ephrin mediated interactions between the mesoderm and endoderm are necessary for asymmetric liver budding and positioning, which also involves changes in endoderm cell shape and rearrangements (Cayuso et al., 2016). There is no evidence currently that Ephrin or Ephrin receptors are downstream of Pitx2c but their communication provides further indication that interactions between mesodermal and endodermal signaling ligands and their receptors influence endodermal cellular behaviors during LR liver development. Alternatively, Pitx2c may influence asymmetric liver organogenesis by regulating the deposition and/or remodeling of extracellular matrix proteins that are vital for growth factor signaling or tissue architecture/epithelial morphogenesis. In the DM of the chick and mouse, Pitx2 induces asymmetric Wnt signaling via Daam2 activation (Welsh et al., 2014). This Wnt signaling is positively regulated by left-sided expression of GPC3, a heparin sulfate proteoglycan extracellular matrix protein that is involved in multiple signaling pathways including Wnt, FGF, SHH, and BMP (Welsh et al., 2014). Interestingly, mutations in GPC3 are associated with biliary atresia, a congenital defect commonly found in patients with heterotaxy syndrome (Cui et al., 2013). Additionally, there are Pitx2c binding sites in the GPC3 promoter (Welsh et al., 2014). It is possible that Pitx2c expression in the mesoderm influences asymmetric liver organogenesis by regulating the expression of extracellular matrix proteins, like GPC3, that are vital for growth factor signaling. Discovery of the

63 downstream organ-specific targets of Pitx2c will provide further molecular insight into the mechanisms of LR asymmetric morphogenesis in the liver and other organs.

Ramifications for human health

Heterotaxia is a congenital defect that affects 1 in 10,000 human infants and causes substantial hardship and morbidity (Lin et al, 2000). Knowledge about the cellular mechanisms by which LR asymmetric organs develop is vital to understanding the etiology of this disease. We know that Pitx2c is required for the asymmetric development of many organs including the heart, lungs, and gastrointestinal tract. Our study provides the first evidence that Pitx2c is also required for asymmetric liver development. The scientific community is just beginning to uncover downstream targets of Pitx2c that drive asymmetric morphogenesis. Here we reveal asymmetries in cell shape and epithelial character that are driven by left-sided Pitx2 expression. Importantly, our study provides evidence that the role of Pitx2 to influence asymmetrical morphology is organ dependent and may depend on the constraints of the tissue. For example, the morphogenetic mechanism by which Pitx2 confers asymmetry to tube-like organs including the heart, gut, and vasculature, may be a very different mechanism than in organs that have a branching or lobation morphology, like the liver, pancreas, and lungs. Answers to these questions will be important as we move forward with genetic therapies to combat diseases like heterotaxy. Additionally knowledge about how organs develop their shape and size will be vital for new technologies like growing organ transplants from stem cells.

Figure 2- 1) Early development of LR asymmetry in the Xenopus liver A-I) In situ hybridization was performed to detect the liver specific markers, hhex, in whole Xenopus embryos (A-E, ventral views) or fibrinogen in dissected gastrointestinal tracts (F-I) at the indicated NF stages. Orange and yellow outlines demarcate the borders of the right and left lobes, respectively (based on the midline and position of the gall bladder) (Scale bars = 200µm). The liver first becomes asymmetric at NF 33, when the right lobe becomes larger and more elongated while the left lobe remains spherical. J) Box and whiskers plot of the length to width ratios of the right and left lobes at NF 46. The right lobe is more elongated than the left, with a higher length to width ratio (p<.0001; n=23). K) Box and whiskers plot of the right/left ratios of the surface area of hhex expression. Ratios close to one (red line) indicate symmetry. Beginning at NF 33, the right/left ratio is significantly greater than one, indicating that the right side of the liver has a greater area. Asterisks indicate significance of p<.05 (*) or p<.0001 (**). NS: no significance. R=right; L=left.

Figure 2- 2) Cellular level LR asymmetries in the early liver diverticulum Transverse sections through the liver at NF 32 (A), 33 (B), 35 (C), and 37 (D) were stained with integrin (int, green) and topro-3 (blue) to reveal cellular morphology. The right side of the diverticulum is outlined in orange and the left in yellow (based on left-right fate mapping, see Fig. S1). Dotted lines on cartoon embryos indicate the plane of section. E-F) Higher magnification of boxed regions in C. Cell shapes are outlined in orange (right, E) or yellow (left, F). G-J) Line graphs displaying apical to basal width ratio (larger numbers indicate apical constriction) (G), length to width ratios (smaller numbers indicate cell elongation) (H), the number of apical to basal cell layers (I), and the cross-sectional surface area on each side of the diverticulum (J). Closed circles indicate the mean value. Error bars = S.E. Asterisks indicate significance of p<.05 (*) or p<.0001 (**) between the left and right sides. NS: no significance. At NF 32 (A), the liver diverticulum is symmetric with a pseudostratified epithelium on both sides. As development proceeds, differences between the left and right sides become apparent. Cells on the left side become rounded in shape compared to elongated, apically constricted cells on the right (E-H). The number of cell layers within the left epithelium increases on the left side, but remains constant on the right side (I). The right liver diverticulum progressively increases in surface area compared to the left (J). R=right; L=left.

Figure 2- 3) The Nodal pathway is required for liver asymmetry Embryos were treated with either DMSO (control) or the Nodal receptor inhibitor, SB505124 (SB505), from NF 19 to 32 and harvested at NF 46 to determine if the Nodal pathway is required for liver asymmetry. Control embryos (A) have normal asymmetry with C-shaped heart curvature (red line) and counter clockwise gut looping (white line). SB505 embryos (C, E) have heterotaxia in both heart curvature and gut looping. In DMSO controls (B), the right liver lobe (orange) is larger and more elongate than the left (yellow). SB505 livers (D, F) have right isomerism, where the left lobe is large and elongate, similar to normal right-sided morphology. Scale bars= A, C, E: 200m B, D, F: 100m. R=right, L=left.

Figure 2- 4) Left-sided Pitx2c is required for LR liver asymmetry In order to determine if left-sided Pitx2c is required for left-right liver asymmetry, two different knockdown strategies were used, Pitx2c CRISPR Cas-9 (Pitx2c CRISPR) or Pitx2c morpholino (Pitx2cMO)(see Fig. S4). Liver morphology was analyzed at NF 46 using in situ hybridization, probing for the liver marker, fibrinogen (R: orange line, L: yellow line) (A-F). In both CRISPR control (A) and control morpholino (COMO) (B), the right lobe of the liver is larger and more elongate than the left. In both Pitx2c CRISPR (C, E) and Pitx2cMO (D, F), the liver is heterotaxic with a right isomerism (R-Iso) phenotype where the left lobe of the liver is larger and elongate, similar to the normal right sided morphology. G) Graphical representation of the percentage of phenotypes (Control CRISPR: 72% normal, 4% R-Iso, 22% left-isomerism (L-Iso), 1.4% reversal (Rev), n=68; Pitx2c CRISPR: 40% normal, 32% R-Iso, 21% L-iso, 5% Rev, n=61; DMSO: 100% normal, n=5; SB505124: 22% normal, 64% R-iso, 4% L-iso, 9% Rev, n=22; CoMo: 95% normal, 4% R-Iso, n=24; Pitx2cMo: 11% normal, 77% R-Iso, 8% L-Iso, 5% Rev, n=66). Cartoon illustrations show each liver phenotype. H) Box and whiskers plot of the length to width ratios of the right and left lobe of the liver at NF 46. In embryos injected with the COMO, the right lobe of the liver has a significantly larger length to width ratio (p<.0001). In embryos injected with the Pitx2cMO, the left lobe has a significantly larger length to width ratio than the normal CoMo left lobe, approaching the normal right sided ratio (p=0.007). Scale bars= 200m. Error bars= S.E. Asterisks indicate significance of p<.05 (*) or p<.0001 (**). NS: no significance. R=right, L=left.

Figure 2- 5) Knockdown of Pitx2c increases left-sided apical constriction and cell elongation . In order to determine the requirement of Pitx2c for normal left sided cellular level morphology, a Pitx2c morpholino (Pitx2c MO) or a control morpholino (COMO) were injected into the future left mesoderm of the liver diverticulum (see Fig. S4). Transverse cross sections at NF 35 were stained with -integrin (green) (A-F). The right side of the diverticulum is outlined in orange and the left in yellow. C-F) Zoomed in views of the liver diverticulum as indicated by white boxes (A-B). G-H) Box and Whiskers plot of the apical/basal cell width ratio (apical constriction) (G) and the cellular length to width ratio (cell elongation) (H) at NF 35. I. Bar graph of the mean number of apical to basal nuclei layers within the epithelium. Error bars= S.E. In COMO injected embryos, cells on the right side (C) have significantly increased apical constriction (G) and cell elongation (H) compared to left sided cells (E) (p<.0001). There is a significant difference between the number of nuclei layers within the epithelium as the left side is stratified and the right is a pseudostratified (I) (p<.0001). Morpholino knockdown of Pitx2c increases left-sided (F) apical constriction (G) and cell elongation (H) and the epithelium becomes pseudostratified with no significant difference compared to right side morphology (C, D). R=right, L=left. Scale bars=100m. Asterisks indicate significance of p<.0001 (**). NS: no significant.

Figure 2- 6) Ectopic right-sided Pitx2c expression induces liver heterotaxia in the form of left isomerism In order to determine if left-limited Pitx2c is required for normal left-right asymmetrical liver development, Pitx2c was ectopically expressed by microinjection of a dexamethasone (Dex) inducible Pitx2cGR construct at the 8 cell stage targeting the right liver diverticulum (see Fig. S4). A-C) in situ hybridization of livers at NF 46, probing for the liver marker, fibrinogen. Orange and yellow outlines demarcate the borders of the right and left lobes, respectively. In control (Dex-) embryos (A), the right lobe is larger and more elongate than the left. In embryos with ectopic right-sided Pitx2c expression (+Dex), the liver is heterotaxic with a left isomerism (L-Iso) phenotype where the right lobe of the liver is smaller and less elongate, similar to the normal left sided morphology (B-C). D) Graphical representation of the percentage of phenotypes (Control, Pitx2cGR-Dex: 88% normal, 2% R-Iso, 5% left- isomerism (L-Iso), 3% reversal (Rev), n=90; Pitx2c GR+Dex: 14% normal, 1% R-Iso, 81% L-iso, 2% Rev, n=74). Cartoon illustrations show each liver phenotype. E) Box and whiskers plot of the length to width ratios of the right and left lobe of the liver at NF 46. In control embryos, the right lobe of the liver has a significantly larger length to width ratio (p<.0001). In embryos with ectopic right-sided Pitx2c, this asymmetry is eliminated (p=0.12). The right lobe has a significantly smaller length to width ratio than the control right lobe (p=0.0001). Asterisks indicate significance of p<.0001 (**). NS: no significant. R=right, L=left. Scale bars= 200m.

Figure 2- 7) Ectopic right-sided Pitx2c expression reduces apical constriction and cell elongation . In order to determine if Pitx2c expression is sufficient to induce left-sided epithelial character, a dexamethasone (Dex) inducible Pitx2cGR construct was co-injected with mGFP as a lineage tracer into the future right mesoderm of the liver diverticulum (see Fig. S4). Transverse cross sections at NF 35 were stained with GFP (red) (A-B) or -integrin (green) (C-H). The right side of the diverticulum is outlined in orange and the left in yellow in C-D. E-H) Zoomed in views of cells in the liver diverticulum indicated by white boxes (C-D). Cell shapes are outlined in orange (right, E-F) or yellow (left, G-H). I-J) Box and Whiskers plots of the apical/basal cell width ratio (apical constriction) (I) and the cellular length to width ratio (cell elongation) (G) at NF 35. K) Bar graph of the mean number of apical to basal nuclei layers within the epithelium. Error bars= S.E. In controls (-Dex) (C), cells on the right side (E) have significantly increased apical constriction (G) and cell elongation (H) compared to left sided cells (G) (p<.0001). There is a significant difference between the number of nuclei layers within the epithelium as the left side is stratified and the right is pseudostratified (I) (p<.0001). In embryos with ectopic right-sided Pitx2c expression (Dex+) (D,F,H), cells on the right side (F) are less elongated (J) and apically constricted (I), with no significant difference compared to left-sided morphology (G,H). In Dex+ embryos, the right side of the epithelium becomes stratified with an increased number of nuclei layers similar to normal left-sided epithelial character (K). Scale bars= 100m. R=right, L=left.

Figure 2- 8) Model of the role of Pix2c in asymmetrical liver development During liver morphogenesis, pitx2c (pink) is expressed in the mesoderm surrounding the left side of the liver diverticulum. The diverticulum is initially symmetrical, comprised of a simple columnar epithelium (NF 30). At NF 33 the epithelium becomes pseudostratified on both sides. At NF 33 and 35, cells on the right side of the diverticulum become elongated and apically constricted while cells on the left side become more rounded in shape. By NF 37, the right side of the diverticulum has expanded into a thin, single layered epithelium, increasing apical and basal surface area, while the left side remains a thickened, multi-layered, stratified epithelium. At NF 46, the right liver lobe is larger and more elongate than the left. The expression of Pitx2c in the left mesoderm surrounding the liver diverticulum is necessary and sufficient for left-right asymmetries at the cellular and gross morphological level.

Supplementary Figure 2- 1) The left and right cells of the liver diverticulum originate from the left and right sides of the early embryo In order to determine the origin of cells in the left and right sides of the liver diverticulum, we performed fate-mapping microinjection experiments. A. Cartoon illustration of fate- mapping experiment where membrane GFP (mgfp) was microinjected into the left vegetal dorsal cell and membrane cherry (mcherry) was microinjected into the right vegetal dorsal cell of the 8 cell embryo. Transverse cross sections through these embryos at NF 35 (B) and 37 (C), staining for mcherry (red) and mgfp (green), reveal a distinct separation between the left and right precursors where right sided injections only contributed to the right lobe of the liver diverticulum and left sided injections only contributed to the left side of the diverticulum. At NF 46 (D), it is apparent that the right lobe is only red and the left lobe is only green.

Supplementary Figure 2- 2) Cell number and proliferation are not different between the left and right sides of the developing liver Transverse sections through the liver at stage 33 (A), 35 (B), 37 (D) and 39 (D) were stained to reveal individual nuclei (blue, topro-3) and proliferating cells (magenta, phospho-histone 3; green, integrin). Orange and yellow lines outlines the right and left liver diverticulum respectively. Box and Whiskers plots indicate there is no significant difference in the total number of cells (E) or mitotic indices (F) between the right and left sides of the liver at any stage. Scale bars= 200m.

Supplementary Figure 2- 3) Pitx2 is asymmetrically expressed in the left mesoderm surrounding the liver diverticulum In order to determine if Pitx2 is expressed asymmetrically in the liver diverticulum, in situ hybridization was performed on whole embryos (A-D) and transverse serial cross sections (E-H) probing for the liver specific transcription factor, hhex (A-B, E-F), and the homeobox transcription factor, pitx2 (C-D, G-H). Cartoon illustrations demonstrate expression patterns in the whole embryo (I) and in section (J). Pitx2 is expressed asymmetrically on the left side of the liver diverticulum in the mesoderm (white arrows). (R=right, L=left). Scale bar= 200m.

Supplementary Figure 2- 4) Pitx2c microinjection manipulation strategies A-C. Cartoon illustrations of Pitx2c manipulation experiments. Pitx2c CRISPR guide RNA, Cas9 mRNA, and membrane GFP (mgfp) mRNA were microinjected at the 1 cell stage (A). Pitx2c morpholino (MO) and mgfp mRNA was microinjected at the 8 cell stage into the future left liver mesoderm (B). A hormone inducible Pitx2cGR construct was microinjected at the 8 cell into the future right liver mesoderm and activated by dexamethasone (Dex) prior to liver organogenesis (C). D-F) Gross morphological phenotypes at NF 46. Orange and yellow outlines demarcate the borders of the right and left lobes, respectively. Pitx2c CRISPR (D) and Pitx2cMO (E) injected embryos display right isomerism. Pitx2cGR embryos (F) activated with Dex display left isomerism phenotypes. Regions of Pitx2c CRISPR (G), Pitx2cMO (H), and Pitx2cGR (I) expression are represented by mgfp (green). Scale bars= 200m. R=right; L=left.

Supplementary Figure 2- 5) Ectopic right-sided Pitx2c expression decreases diverticulum length

A. Box and Whiskers plot of the liver diverticulum length (m) at NF 35 of control embryos (EtOH) or embryos with ectopic right-sided Pitx2c expression (Pitx2cGR). In control embryos, the right side of the liver diverticulum is significantly longer than the left (p<.0001). Ectopic R-sided expression of Pitx2c eliminates this right/left difference (p=0.65) and creates a significant difference compared to the control right side (p<.0001).

References

Abdel-Misih, S.R.Z., and Bloomston, M. (2010). Liver Anatomy. Surg Clin North Am. 90, 643-653.

Barros, E.J.G., Santos, O.F.P., Matsumoto, K., Nakamura, T., and Nigam, S.K. (1995). Differential tubulogenic and branching morphogenetic activities of growth factors: Implications for epithelial tissue development. Proc. Natl. Acad. Sci. U S A. 92, 4412-4416.

Basu, B., and Brueckner, M. (2008). Chapter six cilia: multifunctional organelles at the center of vertebrate left–right asymmetry. Curr. Top. Dev. Biol. 85, 151-174.

Blum, M., Schweickert, A., Vick, P., Wright, C.V.E., and Danilchik, M.V. (2014). Symmetry breakage in the vertebrate embryo: When does it happen and how does it work? Dev. Biol. 393, 109-123.

Bort, R., Signore, M., Tremblay, K., Barbera, J.P.M., and Zaret, K.S. (2006). Hex homeobox gene controls the transition of the endoderm to a pseudostratified, cell emergent epithelium for liver bud development. Dev. Biol. 290, 44-56.

Brennan, J., Norris, D.P., and Robertson, E.J. (2002). Nodal activity in the node governs left-right asymmetry. Genes Dev. 16, 2339-2344.

Brueckner, M. (2007). Heterotaxia, congenital heart disease, and primary ciliary dyskinesia. Circulation 115, 2793-2795.

Bryant, D.M., and Mostov, K.E. (2008). From cells to organs: building polarized tissue. Nature Reviews: Mol Cell Biol. 9, 887-901.

Butler, K., Zorn, A.M., and Gurdon, J.B. (2001). Nonradioactive in situ hybridization to Xenopus tissue sections. Methods 23, 303-312,

Campione, M., Steinbeisser, H., Schweickert, A., Deissler, K., van Bebber, F., Lowe, L.A., Nowotschin, S., Viebahn, C., Haffter, P., Kuehn, M.R., and Blum, M. (1999). The homeobox gene Pitx2: mediator of asymmetric left-right signaling in vertebrate heart and gut looping. Development 126, 1225-1234.

Chung, M.I., Nascone-Yoder, N. M., Grover, S. A., Drysdale, T. and Wallingford, J. B. (2011). Direct activation of Shroom3 transcription by Pitx proteins drives epithelial morphogenesis in the developing gut. Development 137, 1339-1349.

Cui, S., Leyva-Vega, M., Tsai, E.A., Eauclaire, S.F., Glessner, J.T., Hakonarson, H., Devoto, M., Haber, B.A., Spinner, N.B., and Matthews, R.P. (2014). Evidence from human and zebrafish that GPC1 is a biliary atresia susceptibility gene. Gastroenterology 144, 1107-1115.

Davis, A., Amin, N.M., Johnson, C., Bagley, K., Ghashghaei, T., and Nascone-Yoder, N. (2017) Stomach curvature is generated by left-right asymmetric gut morphogenesis. Development.

Dush, M.K., and Nascone-Yoder, N.M. (2013). Jun N-terminal kinase maintains tissue integrity during cell rearrangement in the gut. Development 140, 1457-1466.

Dush, M.K., McIver, A.L., Parr, M.A., Young, D.D., Fisher, J., Newman, D.R., Sannes, P.L., Hauck, M.L., Dieters, A., and Nascone-Yoder, N. (2011). Heterotaxin: a TGF- signaling inhibitor identified in a multi-phenotype profiling screen in Xenopus embryos. Chem. Biol. 18, 252-226.

Eng. D., Campbell, A., Hilton, T., Leid, M., Gross, M.K., and Kioussi, C. (2010). Prediction of regulatory networks in mouse abdominal wall. Gene 469, 1-8.

Faucourt, M., Houliston, E., Besnardeau, L., Kimelman, D., and Lepage, T. (2001). The Pitx2 homeobox protein is required early for endoderm formation and nodal signaling. Dev. Biol. 229, 287-306.

Gage, P.J., Suh, H., and Camper, SA. (1999). Dosage requirement of Pitx2 for development in multiple organs. Development. 126, 4643-4651.

Gray, H. (1918). Anatomy of the Human Body. Philadelphia: Lea & Febiger, Bartleby.com, 2000. www.bartleby.com/107/.

Heisler, J.C. (2008). A text-book of embryology: for students of medicine. The University of California, Saunders, 1907. Digitized: 2008.

Hunter, M.P., Wilson, C.M., Jiang, X., Cong, R., Vasavada, H., Kaestner, K.H., and Bogue, C.W. (2007). The homeobox gene Hhex is essential for proper hepatoblast differentiation and bile duct morphogenesis. Dev. Biol. 308, 355-367.

Ikari, T., Hiraki, A., Seki, K., Sugiura, T., Matsumoto, K., and Shirasuna, K. (2003). Involvement of hepatocyte growth factor in branching morphogenesis of the murine salivary gland. Dev Dyn. 228, 174-184.

Kaestner, K.H. (2005). The making of the liver: competence in the foregut endoderm and induction of liver-specific genes. Cell Cycle 4, 1146-1148.

Kawasumi, A., Nakamura, T., Iwai, N., Yashiro, K., Saijoh, Y., and Belo, J.A. (2011). Left-right asymmetry in the level of active Nodal protein produced in the node is translated into left-right asymmetry in the lateral plate of mouse embryos. Dev. Biol. 353, 321-330.

Lin, A.E., Ticho, B.S., Houde, K., Westgate, M.N., and Holmes, L.B. (2000). Heterotaxy: associated conditions and hospital-based prevalence in newborns. Genet. Med. 2, 157-172.

Lin, C.R., Kioussi, C., O'Connell, S., Briata, P., Szeto, D., and Liu, F. (1999). Pitx2 regulates lung asymmetry, cardiac positioning and pituitary and tooth morphogenesis. Nature 401, 279-282.

Lipscomb, K., Schmitt, C., Sablyak, A., Yoder, J.A., and Nascone-Yoder, N. (2006). Role for retinoid signaling in left-right asymmetric digestive organ morphogenesis. Dev. Dyn. 235, 2266-75.

Liu, C., Liu, W., Lu, M., Brown, N.A., and Martin, J.F. (2001). Regulation of left-right asymmetry by thresholds of Pitx2c activity. Development 128, 2039-2048.

Lu, M.F., Pressman, C., Dyer, R., Johnson, R.L., and Martin, J.F. (1999). Function of Riegar syndrome gene in left-right asymmetry and craniofacial development. Nature. 401, 276-278.

Margagliotti, S., Clotman, F., Pierreux, C.E., Lemoine, P., Rousseau, G.G., Henriet, P., and Lemaigre, F.P. (2008). Role of metalloproteinases at the onset of liver development. Dev. Growth Differ. 50, 331-338.

Mazziotti, M.V., Willis, L.K., Heuckeroth, R.O., LaRegina, M.C., Swanson, P.E., Overbeek, P.A., and Perlmutter, D.H. (1999). Anomalous development of the hepatobiliary system in the inv mouse. Hepatology 30, 372-378.

Nieuwkoop, P.D., and Faber, J. (1994). Normal Table of Xenopus laevis (Daudin). New York: Garland Publishing Inc.

Raya, A., and Izpisua Belmonte, J.C. (2006). Left-right asymmetry in the vertebrate embryo: from early information to higher-level integration. Nat. Rev. Genet. 7, 283-293.

Sawyer, J.M., Harrell, J.R., Shemer, G., Sullivan-Brown, J., Roh-Johnson, M., and Goldstein, B. (2010). Apical constriction: a cell shape change that can drive morphogenesis. Dev. Biol. 341, 5-19.

Shimadera, S., Iwai, N., Deguchi, E., Kimura, O., Fumino, S., and Yokoyama, T. (2007). The inv mouse as an experimental model of biliary atresia. J. Pediatr. Surg. 42, 1555-1560.

Shiraishi, I., and Ichikawa, H. (2012). Human heterotaxy syndrome. From molecular genetics to clinical features, management and prognosis. Circ. J. 76, 2066-2075.

Shiratori, H., Yashiro, K., Shen, M.M., and Hamada, H. (2006). Conserved regulation and role of Pitx2 in situs-specific morphogenesis of visceral organs. Development 133, 3015- 3025.

Sive, H.L., Grainger, R.M., and Harland, R.M. (1998). Early Development of Xenopus laevis. Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press.

Tremblay, K.D. (2011). Inducing the liver: Understanding the signals that promote murine liver budding. J. Cell. Physiol. 226, 1727-1731.

Tremblay, K., and Zaret, K.S. (2005). Distinct populations of endoderm cells converge to generate the embryonic liver bud and ventral foregut tissues. Dev. Biol. 280, 87-99.

Varner, V.D., and Nelson, C.M. (2014). Cellular and physical mechanisms of branching morphogenesis. Development. 141, 2750-2759.

Yoshiba, S., and Hamada, H. (2014). Roles of cilia, fluid flow, and Ca2+ signaling in breaking of left-right symmetry. Trends Genet. 30, 10-17.

Zhang, Y.W., and Vande Woude, G.F. (2003). HGF/SF-met signaling in the control of branching morphogenesis and invasion. J Cell Biochem. 88, 408-417.

Zhu, L., Belmont, J.W., and Ware, S.M. (2005). Genetics of human heterotaxias. Eur. J. Hum. Genet. 14, 17-25.

Zorn, A.M., and Wells, J.M. (2009). Vertebrate Endoderm Development and Organ Formation. Annu. Rev. Cell Dev. Biol. 25, 221-251.

CHAPTER 3: Discussion

As in humans and other vertebrates, in our model frog, X. laevis, the right lobe of the liver is larger and more elongated than the left lobe which remains small and spherical in shape. I found that LR asymmetries in cell shape, independent of cell proliferation and apoptosis, downstream of the homeobox transcription factor, Pitx2c, are required for this laterality. In general, the shape and size of organs of the body is determined by the delicate interplay between cell movement, cell division/death, cell signaling, ECM composition, and changes in cell size and shape (Auman et al., 2007; Lecuit and Lenne, 2007). LR differences in any of these processes could lead to morphological asymmetries in organ systems. Below,

I discuss: the conserved role and possible mechanisms by which Pitx2c induces (1) cell shape changes, (2) cellular rearrangement, and tissue remodeling in the liver and other LR asymmetric organs, (3) endodermal and mesodermal interactions that mediate organ morphogenesis, (4) possible downstream targets of Pitx2c during the development of liver laterality, and (5) implications for human health.

1) Pitx2 induces left-right asymmetries in cell shape

We found that cells on the right side of the liver diverticulum become progressively more elongated on their apical to basal axis and more apically constricted than the cells on the left that are rounded in morphology. Pitx2c is required to prevent cell shape changes in the left side of the liver diverticulum as knockdown of Pitx2c in the left lobe causes endodermal liver cells to become more elongated and apically constricted, while ectopic

Pitx2c in the right lobe causes these cells to become rounded in morphology. In other

87 asymmetric organs like the heart and midgut, cell shape changes are also required for organ laterality (Auman et al., 2007; Davis et al., 2008). Our findings are consistent with the idea that cell shape change is a conserved aspect of LR asymmetric organogenesis downstream of

Pitx2. It is likely that proteins required to orchestrate cell shape changes including cytoskeletal components and cell adhesion molecules act downstream of Pitx2 during the development of liver laterality. Our results also reveal that the localization (right versus left) of specific cell shape changes (i.e. apical-basal elongation and apical constriction vs. cuboidal and rounded) is organ specific and may depend on the geometry of each individual organ system.

1.1) Pitx2’s influence on Apical-Basal Cell Elongation and Apical Constriction

Cell shape changes are a conserved aspect of LR asymmetric organ development, however, the exact morphological shape of left sided cells induced by Pitx2 expression depends on the organ system. For example, cell shape changes during zebrafish heart chamber development are similar to my findings during liver development as cells on the outer curvature (right side) of the heart ventricle become elongated, aligning with one another in an organized fashion, while cells on the inner curvature (left side) maintain a cuboidal morphology (Auman et al., 2007). On the other hand, in the dorsal mesentery of the midgut, Pitx2 is required for cells in the left epithelial compartment to remain columnar and apically constricted or bottle-shaped in morphology while cells on the right side become more flattened and cuboidal, opposite to side specific cell shapes observed in the heart and in the liver (Davis et al., 2008). These asymmetries in cell shape induce organ laterality as

88 apical to basal elongation and apical constriction on the left side of the DM causes the gut tube to tilt leftward (Davis et al., 2008), while right sided cell elongation generates chamber curvature in the heart (Auman et al., 2007) and produces an apical to basal expansion of the right liver lobe. It is likely, that the specific gene expression profile within individual organs is specialized to orchestrate cell shape changes necessary to evoke distinct organ morphologies.

Cell shapes during embryogenesis are influenced by surface tensions and forces generated by interactions between molecular motors, the cytoskeleton, and cell-cell adhesions (Lecuit, and Lenne, 2007; Sawyer et al., 2010). Therefore, Pitx2c likely influences the expression of genes involved in these processes and the expression of other organ specific genes that “fine tune” the type of cell shape change generated to orchestrate LR asymmetric morphology. For example, apical constriction is a driving force behind tissue bending mechanics during Xenopus gastrulation, Drosophila tracheal and salivary gland development, chick inner ear formation, and lung branching morphogenesis in multiple species (Lecuit and

Lenne, 2007, Sawyer et al., 2010; Varner and Nelson, 2014). F-actin and Myosin II cooperation is vital for apical constriction to occur in all of these contexts (Sawyer et al.,

2010, Varner and Nelson, 2014). Apical constriction is driven by contraction of apical F- actin in cooperation with Myosin motors (Sawyer et al., 2010). Myosin II is phosphorylated by RoK which is activated by RhoGEFs and Rho kinases (Lecuit and Lenne, 2007). Pitx2c likely regulates the expression of genes that influence actin-myosin cooperation within the left side of the liver diverticulum to prevent apical constriction.

Although we found that Pitx2 inhibits apical constriction during liver development,

Pitx2 is known to induce apical constriction in the DM of the midgut (Plageman Jr. et al.,

2011). In this context, N-cadherin, downstream of Pitx2 cooperates with Shroom3 to increase apical constriction on the left side (Plageman Jr. et al., 2011). In fact, there is an asymmetrical localization of F-actin and Myosin in the left epithelial compartment of the DM during midgut looping events (Plageman Jr. et al., 2011). Additionally, Pitx2a is known to activate RhoGTPases, Rac1 and RhoA, in cell culture by inducing the expression of Trio, a guanine nucleotide exchange factor (Wei and Adelstein, 2002). Therefore, Pitx2 may influence the expression of molecules that regulate cell shape changes in multiple contexts; however, whether Pitx2 induces or restricts apical constriction or apical to basal cell elongation appears to be organ specific. It will be important to determine the expression and localization patterns of Shroom 3, F-actin, and Myosin II within the liver diverticulum as well as to determine if any of the molecules involved in the activation of myosin, like Rho kinases and Rho GEFs, are influenced by Pitx2c. The organization of the actin cytoskeletal is also influenced by cell adhesion properties within the DM of the gut, therefore, cell adhesion proteins are another likely target downstream of Pitx2 during LR liver development as discussed below.

1.2) The role of cell adhesion in cell shape change

Cell shape can also be influenced by cell adhesion properties. It is possible that LR asymmetries in cell shape observed in the developing liver diverticulum are the result of differential cell adhesion. Cell shape is controlled by the organization of the plasma

90 membrane into surface contacts with other cells and the extracellular environment (Lecuit and Lenne, 2007). Cells typically organize in clusters to minimize their surface area of contact with the environment and each other while being stabilized by cell to cell adhesion molecules (Lecuit and Lenne, 2007). Cell membranes will contract due to myosin motors acting on actin filaments and stronger contractions in one location in the cell will change cell shapes (Lecuit and Lenne, 2007). Cells will minimize total surface area of contact by assuming a default rounded or hexagonal shape, similar to left side cell shapes in the liver diverticulum. E-cadherin mediated cell to cell adhesions are required to overcome this surface tension to increase the surface area of contact between cells to induce an elongated shape (Lecuit and Lenne, 2007). Cells on the right side of the liver diverticulum have an elongated shape so it is possible that there is an increase in the expression or localization of

E-cadherin on the right side. It is likely that differences in cell to cell adhesive properties and cytoskeletal rearrangements account for LR asymmetries in cell shape during liver morphogenesis including LR differences in apical constriction and apical to basal elongation.

In other contexts, like the DM of the midgut, N-cadherin and the formin protein,

Daam1, are asymmetrically expressed on the left side (Kurpios et al., 2008, Welsh et al.,

2014). N-cadherin is a cell to cell adhesion protein while formin proteins, like Daam1, polymerize F-actin to form stress fibers necessary for cell adhesion (Welsh et al., 2014). The asymmetric expression of these genes is downstream of Pitx2 in the DM and required for left-sided cell elongation and apical constriction in the epithelial compartment that is necessary for midgut looping (Kurpios et al., 2008, Welsh et al., 2014). As mentioned previously, we found that Pitx2 has an opposite effect on cell shape during the development

91 of liver laterality as compared to midgut looping. This suggests that cells on the left side of the liver instead of the right may minimize their surface area of contact by becoming more rounded while cells on the right overcome this tension due to cell adhesion mediated increases in their length to width ratio. Cell adhesions are stabilized by -catenin, -catenin, and actin filaments (Lecuit and Lenne, 2007). Therefore, it would be interesting to determine if the expression or localization of cadherins and associated molecules like catenins are elevated on the right side compared to the left in the developing liver diverticulum. I found that there are no differences in the localization or intensity of -catenin during X. laevis liver diverticulum development (data not shown), however, the localization pattern of N-cadherin and E-cadherin remains to be investigated. Importantly, the opposing effect of Pitx2 on cell shape in the liver as compared to the dorsal mesentery suggests that, while the role of Pitx2 in orchestrating cell shape is conserved, the mechanism of action and final consequence of this action is tissue- and organ-specific.

2) Pitx2c directs left-right asymmetrical epithelial remodeling

Our study also uncovered LR asymmetries in the number of nuclei within the epithelial layer of the developing liver diverticulum. At the beginning of liver development, the diverticulum is symmetrical, with a pseudostratified epithelium on both sides. At a stage in which morphological asymmetries are first apparent, the right side of the diverticulum remains a single layered pseudostratified epithelium while the left side demonstrates cell stacking as the tissue is multiple layers of nuclei thick and stratified. In the absence of left-

92 right differences in cell proliferation, this suggests that the left and right epithelial walls of the liver diverticulum are undergoing differential remodeling events.

2.1) Cellular rearrangement

Cellular rearrangements are a conserved feature of embryogenesis and are a necessary mechanism for increasing length and surface area in a variety of organs (Pilot and Lecuit,

2005; Reed et al., 2009). We found that the right side of the developing liver diverticulum expands its surface area, resulting in an increased length to width ratio of the mature right lobe. However, the left side of the liver diverticulum increases girth at the expense of surface area as cells stack on top of one another and become stratified, resulting in a smaller, round liver lobe. This suggest that the right side of the liver may undergo more cellular rearrangement than the left during LR organogenesis. In other organs like the Drosophila trachea, epithelial cells undergo cell intercalation causing dorsal elongation of the organ primordium (Pilot and Lecuit, 2005). Interestingly, apical constriction precedes the cell junction remodeling that is required for this elongation (Pilot and Lecuit, 2005; Varner and

Nelson, 2014). It is possible that the right-sided increase in apical constriction we discovered in the liver diverticulum is a necessary precursor to expansion of the organ’s right side.

During gastrointestinal development, the primitive gut tube also undergoes tubal elongation as it is remodeled from a multilayered group of cells to a single-layered epithelium lining the gut tube (Reed et al., 2009). Cells in the middle of the gut tube reorganize due to changes in polarity and cell shape, radially intercalating between neighboring cells (Reed et al., 2009). The Rho/Rock/Myosin II pathway is required for this

93 process as inhibition prevents cellular rearrangement and subsequent elongation (Reed et al.,

2009). It is intriguing that the Rho/Rock/Myosin II pathway is required for both apical constriction and cell rearrangement processes in multiple organ systems as we see both processes occurring asymmetrically in the right liver diverticulum. A promising future direction for understanding asymmetrical epithelial organization within the liver diverticulum would be to determine the expression and role of Rho, Rock, and Myosin II in liver laterality.

In the stomach, another asymmetric organ, cell rearrangement is required for differences in the length of the greater curvature on the left and the lesser curvature on the right (Davis et al., 2017). Pitx2c induces the left wall of the stomach epithelium to become thinner than the right even though the total number of cells remains the same between the two sides (Davis et al., 2017). The endoderm of the right stomach wall is stratified with increased nuclei layers while the left stomach wall is highly polarized and pseudostratified

(Davis et al., 2017). The role of Pitx2c in orchestrating asymmetrical tissue reorganization is conserved during both stomach and liver development; however, once again we see opposing downstream effects. In the liver, the right side undergoes epithelial thinning while the left side becomes more stratified; in the stomach, the opposite tissue changes occur. Our findings show that the molecular tools used to create asymmetries may be conserved between organs, but the means by which these tools are used is dependent on the tissue involved and the final geometry of the organ.

The mechanism by which cellular rearrangements occur asymmetrically downstream of Pitx2 within the liver diverticulum is yet to be elucidated but likely involves the same genes that induce cell shape changes including cytoskeletal components and adhesion

94 molecules. Another possible mechanism could involve LR differences in the reincorporation of dividing cells into the developing liver epithelium as discussed below.

2.2) Interkinetic nuclear migration

During liver morphogenesis in the mouse, the hepatic diverticulum is initially a single layered columnar epithelium that undergoes interkinetic nuclear migration (INM) while transitioning to a pseudostratified liver bud (Zorn and Wells, 2009). INM is a hallmark of cell division in pseudostratified epithelia and is found in the embryonic neural tube, developing retina, and elongating gastrointestinal tract (Grosse et al., 2011, Kosodo et al.,

2012). During INM, nuclei adjacent to the basement membrane begin to synthesize DNA before moving apically during the G2 phase where they undergo mitosis before moving back towards the basal membrane during the G1 phase (Grosse et al., 2011; Kosodo et al., 2012).

INM within pseudostratified epithelia allows for the packing of a greater density of cells within a confined space and promotes rapid tissue expansion (Lee et al., 2013). Additionally, epithelial remodeling following cell division occurs differently in pseudostratified epithelium versus stratified epithelium as pseudostratification allows for the expansion of luminal surfaces through cell shape changes (apical-basal cell elongation and apical constriction) as cells are reincorporated into the epithelium while stratification allows for thickening of the tissue by the addition of dividing cells into layers and de novo polarization of central cells to form lumina (Grosse et al., 2011). Therefore, pseudostratified and stratified tissues form different geometric organ shapes through different epithelial remodeling events. During LR liver development, we found that cells on the left side become stratified while cells on the

95 right are pseudostratified. This difference in epithelial organization following apical cell division could account for the LR morphological asymmetries we observed.

In other organs like the gastrointestinal tract, INM is responsible for remodeling the epithelium to form finger-like projections called villi (Grosse et al., 2011). Interestingly, the liver diverticulum also forms finger-like projections called sinusoids or hepatic cords. Like the liver, the early intestinal pseudostratified epithelium undergoes INM and increases surface area by apical basal cell elongation. This remodeling event requires Shroom3-driven microtubule and actomyosin activity to drive both cell shape change and re-integrate apically dividing cells into a single pseudostratified layer, similar to epithelial morphology that we observed in the right liver diverticulum (Grosse et al., 2011). In the absence of Shroom3, apical surfaces of the intestine become disorganized as the dividing cells remain rounded in shape, with expanded apical surfaces, as they become trapped at the luminal surface, creating a stratified epithelium, similar to cellular morphology observed in the left liver diverticulum

(Grosse et al., 2011). It is possible that remodeling events observed in the context of intestinal villi formation are occurring asymmetrically on the right side of the developing liver. Shroom3 is required for cell incorporation following the division cycle within the gastrointestinal tract through its effect on microtubule and actomyosin activity. Interestingly,

Pitx1 directly activates Shroom3 during neuronal development (Grosse et al., 2011).

Although no direct link between Shroom3 and Pitx2 has been discovered, as mentioned previously, Shroom3 cooperates with the Pitx2 target, Ncad, for left-sided cell shape changes during midgut looping events (Plageman Jr. et al., 2011). Therefore, it is possible that Pitx2c influences Shroom3 and its downstream targets during LR liver development to influence

INM remodeling events as well as cell shape changes that ultimately influence final LR asymmetrical liver morphology.

In our study, we found that both sides of the liver diverticulum display similar amounts of cell proliferation and that this proliferation occurs at the apical surface (data not shown), however, the right side remains a pseudostratified epithelium while the left side becomes stratified. This suggests that cells on the right side are reincorporated into a pseudostratified epithelium after INM-mediated proliferation, becoming apically constricted and elongating on their apical to basal axis and thus expanding the surface area of this lobe similar to the increased surface area observed following INM in the developing intestine. In contrast, cells on the left side may not become reincorporated after mitosis, remaining rounded, and apically expanded at the luminal surface, thus increasing the thickness and girth of this tissue but not increasing surface area. The final morphological result of this asymmetry is to increase the length to width ratio of the right lobe, making it larger and more elongate compared to the left lobe which remains rounded in shape.

Future research should include studies observing the movement of cells during the division cycle and the effect of left-sided Pitx2c expression on this movement. By staining cells during the S phase with BrdU, we could observe the movement of cells during the subsequent division cycle. I would predict that BrdU+ cells on the right side would divide at the lumen and become reincorporated into the pseudostratified epithelium, while those on the left would divide and remain at the lumen.

Pitx2c could also influence the cell cycle itself during INM as Pitx2 controls proliferation in pituitary development and epidermal keratinocytes by controlling G1 phase

97 cell cycle genes like p21 and cyclin D1 (Kioussi et al., 2002; Shi et al., 2010). However, this is unlikely as the number of dividing cells and the location of this division does not change between the left and right sides of the diverticulum. It is more likely that the movement and reincorporation of these cells after division is asymmetric.

3) Pitx2c expression in the mesoderm influences endodermal tissue architecture

We found that Pitx2c is expressed in the mesoderm surrounding the left side of the developing liver endoderm. This mesodermal expression is necessary and sufficient to drive

LR differences in the cellular architecture of the developing liver diverticulum that results in asymmetries in the size and shape of the left and right liver lobes. Knockdown of Pitx2c in the left mesoderm influences cell shapes and epithelial organization in the diverticulum, and causes final liver lobe morphology to resemble the normal right side. Ectopic Pitx2c expression in the right mesoderm induces right side diverticulum cells to adopt left sided shape, tissue architecture, and final lobal morphology. Thus, the asymmetric expression of a transcription factor in the neighboring mesoderm influences endodermal tissue morphogenesis. How can a mesodermally derived transcription factor influence epithelial morphogenesis? Although Pitx2c cannot directly influence endodermal development, it can inhibit or activate the transcription of genes (e.g., signaling ligands and/or ECM components) required for the mesodermal-endodermal interactions that influence cell shape and tissue architecture. Examples of known mesodermal and endodermal interactions and implications for Pitx2c signaling during LR liver development are discussed below.

3.1) Endodermal and Mesodermal interactions mediated by signaling ligands/receptors

Reciprocal endodermal and mesodermal interactions are required for the morphogenesis of many organs including the liver. During zebrafish liver development and mouse DM vasculature formation, signaling between two opposing tissue compartments is required for LR asymmetry (Cayuso et al., 2016; Mahadevan et al., 2014). It is likely that asymmetries in the expression of signaling ligands or their receptors downstream of Pitx2c orchestrates the interactions between the mesoderm and endoderm that are critical for X. laevis liver laterality.

Zebrafish liver laterality is different from other animals in that the location of the liver is LR asymmetric (left of the midline) but there are no differences in lobal morphology as seen in Xenopus, humans, and mice. However, knowledge about signaling interactions between the endoderm and mesodermal that are necessary for asymmetric liver placement in zebrafish may give us insight into LR liver lobation in other species. For example, during liver morphogenesis in zebrafish, reciprocal interactions between the mesoderm and endoderm coordinate the migration of the hepatic endoderm towards its asymmetric placement left of the midline (Cayuso et al., 2016). EphrinB1 in the endoderm interacts with its receptor EphB3b in the lateral plate mesoderm to influence endodermal polarity and directional migration by controlling actin based protrusions needed for hepatoblast motility

(Cayuso et al., 2016). EphrinB1 mediates actin based extension formation while EpB3b interactions insure extension stabilization (Cayuso et al., 2016). Therefore, signaling interactions between epithelial and mesenchyme tissues coordinate zebrafish liver laterality.

However, it is unknown if Pitx2 is required for the expression of EphrinB1 or EphB3b during

99 this process or whether similar signaling molecules are required for asymmetric lobation in other species.

During vasculature development in the DM of the mouse, Pitx2 expression in the mesenchyme influences the asymmetric left-sided migration of endothelial cells in which

Pitx2 expression is absent (Mahadevan et al., 2014). Pitx2 promotes the transcription of

CxCl12 in the mesenchymal compartment that signals to endothelial cells through its receptor Cxcr4, which is expressed in the endothelium (Mahadevan et al., 2014). Therefore, during Xenopus LR asymmetrical liver development, it is possible that Pitx2c promotes or inhibits the transcription of a signaling molecule or receptor in the mesoderm to interact with and influence endodermal morphogenesis. Future research should focus on such factors as downstream targets of Pitx2.

3.2) Mesodermal and Endodermal interactions mediated by ECM components

The extracellular matrix (ECM) is a compartment composed of proteins, polysaccharides, and water that offers several different functions during tissue morphogenesis including providing structure to define tissue boundaries, acting as an adhesive substrate for migratory cells, presenting, sequestering, and storing growth factors, and sensing/transducing mechanical signals (Frantz et al. 2010, Rozario and Desimone,

2010). The ECM can interact with cell surface receptors to initiate signal transduction and regulate gene transcription between tissue layers like the mesoderm and endoderm (Frantz et al., 2010). Therefore, it is possible that LR asymmetries in ECM composition, downstream of

100

Pitx2 may mediate mesodermal and endodermal tissue interactions to orchestrate the development of liver laterality.

In other organs, ECM components are required to orchestrate proper organogenesis.

For example, the ECM is important for the migration of cardiac precursors and neural crest, the invasion of epithelial buds into the surrounding mesenchyme during branching morphogenesis in multiple tissues, and is implicated in the development of the mammary gland, salivary gland, kidney, gut, and lung (Rozario and Desimone, 2010). Additionally, the

ECM protein Syndecan2 is required for LR asymmetric organ development as knockdown of this gene in Xenopus induces heterotaxia of the gut and heart (Rozario and Desimone, 2010).

In zebrafish, the ECM protein, Laminin, is required for the laterality of the liver and pancreas as laminin mutants display midline livers and bilateral pancreas structures (Hochgreb-Hagele et al., 2013). During zebrafish gut looping, ECM remodeling is required for the asymmetric migration of the LPM which displaces the gut tube to the left (Yin et al., 2010). During zebrafish heart development, interactions between cardiac precursors and the ECM protein,

Fibronectin, are required for heart jogging and looping events (Bakkers et al., 2009).

Additionally, the ECM protein, Hyaluronic acid (HA), downstream of Has2, is required for the left-anterior myocardial cell migration events necessary for leftward heart jogging

(Bakkers et al., 2009, Smith et al., 2008)). It is unknown whether any of these ECM proteins are expressed LR asymmetrically within the liver diverticulum or whether Pitx2 controls

ECM composition in any of the contexts described above. However, Pitx2 is known to control LR asymmetries in the ECM during midgut rotation in mice.

101

During midgut looping in mice, asymmetries in the ECM deposition within the dorsal mesentery, downstream of Pitx2 and Ncad, are required for gut looping (Kurpios et al.,

2008). The mesenchymal compartment on the left side of the DM has a more condensed morphology than the right (Kurpios et al., 2008). The left mesenchyme has higher levels of

GAGs while the right mesenchyme consists of more HA (Kurpios et al., 2008). HA is a large protein that is negatively charged and attracts a lot of water, causing interstitial fluid to swell, preventing strong cell to cell contacts, and inducing the right side of the DM to be less condensed than the left (Kurpios et al., 2008). Misexpression of both Pitx2 and its downstream target Ncad affects the composition of ECM proteins within each compartment.

Therefore, it is possible that Pitx2 influences the composition of the ECM during the development of liver laterality.

ECM proteins can also influence cell shape due to biomechanical and structural properties (Frantz et al. 2010, Rozario and DeSimone, 2010). It is possible that Pitx2c could influence the expression of ECM molecules within the liver diverticulum to influence LR asymmetrical liver shape and remodeling through changes in the composition of the ECM or by coordinating mesenchymal to epithelial signaling events to ultimately direct liver laterality. Future research should focus on ECM proteins that may act as downstream targets of Pitx2c during LR liver development.

3.3) Differential expression of Pitx2 in organ tissues layers

We found that Pitx2c is expressed in only the mesodermal layer surrounding the developing liver endoderm, suggesting that the effect of Pitx2 on the underlying epithelium is

102 indirect. However, in the dorsal mesentery (DM) of the midgut, Pitx2 is expressed within the cells that adopt asymmetric morphologies; therefore, the morphogenetic events downstream of Pitx2 are likely direct in this context. In the stomach, Pitx2c is expressed in the endoderm and in the surrounding mesoderm, so, in this case, Pitx2 most likely acts both directly and indirectly.

In the gastrointestinal tract and stomach, the effect of Pitx2 on epithelial morphology during LR organogenesis is also opposite to that seen in the liver. For example, in the DM of the midgut, cells on the left side become columnar and apically constricted. Conversely left- sided cells in the developing liver diverticulum are rounded in morphology and right sided cells are apically constricted and elongated. In the developing stomach, the left side of the epithelium becomes polarized and undergoes cellular rearrangements to form a single layered epithelium. In the liver diverticulum, right sided cells remodel into a single layered epithelium while the epithelium on the left side becomes stratified. Therefore, the differences in the location of Pitx2 expression and the tissue affected by this expression may account for the somewhat opposing morphogenetic events observed downstream of Pitx2. Moving forward, it will be important to determine how Pitx2 acts differently through direct and indirect mechanisms. In other LR asymmetric organs in which the effect of Pitx2 on the development of laterality has yet to be elucidated (e.g., lungs), it will also be important to determine the tissue layer in which Pitx2 is expressed in order to predict cellular level mechanisms.

103

4) Downstream targets of Pitx2c during left-right liver development

We identified the cellular level morphological differences that are orchestrated by

Pitx2c in the developing liver diverticulum to shape final organ laterality; however, the specific downstream molecular targets of Pitx2c have yet to be elucidated. It is likely that

Pitx2c, as a transcription factor, either inhibits or promotes the transcription of genetic pathways required to signal from the mesoderm to the endoderm and induce changes in cell shape and cellular rearrangements. Possible targets include members of the Wnt signaling pathway, extracellular matrix proteins, soluble growth factors, and chemokines.

4.1) Components of the Wnt signaling pathway

Wnt signaling is a conserved pathway controlling many aspects of morphogenesis including cell proliferation, differentiation, polarization, and movement (rearrangement and migration) (Dale et al., 2009, Yang et al., 2012). Wnt signaling is known to promote the later stages of liver development including liver bud outgrowth and proliferation (Si-Tayeb et al.,

2010). It is possible that some Wnt signaling components act asymmetrically, as downstream targets of Pitx2c, to orchestrate the many morphogenetic aspects of liver laterality.

The Wnt/-catenin (-cat) pathway or canonical pathway acts to stabilize nuclear - cat and regulate cell proliferation and differentiation (Yang et al., 2012). In the absence of

Wnt ligands, -cat is phosphorylated by GSK-3 within the destruction complex and is degraded (Yang et al., 2012). However, the binding of Wnt ligands to their heterodimeric receptors, Frizzled (Frz) and LRP5/6, induces a receptor conformational change, causing Frz to bind and activate Dishevelled (Dsh) (Clevers and Nusse, 2012). These downstream

104 signaling events inactivate or degrade the -cat destruction complex, stabilizing and translocating -cat to the nucleus (Yang et al., 2012). Within the nucleus, -cat binds to Lef and TCF factors to regulate gene expression (Yang et al., 2012). Interestingly, Pitx2 and the

-cat binding partner, Lef-1, show overlapping expression during tooth development

(Vadlamudi et al., 2005). Pitx2 deficient mice show reduced Lef-1 expression and Pitx2 can activate the human Lef-1 promoter (Vadlamundi et al., 2005). In this context, Pitx2 binds directly with Lef-1 and -cat to synergistically regulate downstream gene expression

(Vadlamudi et al., 2005). Pitx2 is also known to interact with B-cat and mir200 during dental epithelial differentiation (Sharp et al., 2014). Additionally, Pitx2 increases the expression of

Lef-1 during breast cancer metastasis (Pillai et al., 2015). Pitx2 is also known to regulate Wnt ligands during human ovarian cancer by directly binding to their promoter regions (Basu and

Roy, 2013). Pitx2’s interactions with members of the Wnt/-cat pathway during tooth and cancer morphogenesis highlight the role of Pitx2 in regulating differentiation and proliferation in these tissues. However, our study discovered no differences in mitosis or liver specification/differentiation in the left versus right liver diverticulum suggesting that it is unlikely that Pitx2 acts through the canonical Wnt signaling pathway to initiate liver laterality.

The Wnt/Planar Cell Polarity or non-canonical pathway is a -cat independent pathway that mediates cytoskeleton reorganization critical for cell movement and cell polarization during morphogenesis (Dale et al., 2009). Wnt5 and Wnt 11 ligands bind to the

Frz receptor activating downstream signaling partners: Vangl, Prickle, Dsh, Daam1, RhoA and Rac1. RhoA interacts with Rock to induce cytoskeletal changes (Yang et al., 2012). Rock

105 acts to phosphorylate regulators of the actin cytoskeleton including myosin light chain kinase

(Spiering and Hodgson, 2011). The morphogenesis of many different organs requires the

Wnt/PCP pathway to orchestrate apical constriction, cell shape changes, and cellular rearrangements (Sawyer et al., 2010).

In the dorsal mesentery (DM) of the gut many different Wnt/PCP components are expressed asymmetrically. Frz 4/8 and Daam2 are expressed on the left side of the DM while

Prickle and the Wnt inhibitors Sfrp 1/2 are expressed on the right (Welsh et al., 2014).

Daam2, a formin protein and intracellular effector of Wnt signaling transduction, polymerizes unbranched actin filaments (F-actin) and forms the stress fibers required for cytoskeletal rearrangements, adhesion, and cell polarity (Welsh et al., 2014). Daam2 functions downstream of Pitx2 in the DM to induce left-sided mesenchymal condensation required for gut tube tilting by regulating actin based filopodial extensions and the size and formation of cadherin based junctions (Welsh et al., 2014). In fact, there are binding sites for

Pitx2 in the Daam 2 promoter (Welsh et al., 2014). Surprisingly, Daam1 is strongly expressed on just the left side of the developing mouse liver, suggesting that Daam1 is a possible downstream target of Pitx2c during LR liver development (Nakaya et al., 2004).

Pitx2c could also influence the expression of other Wnt/PCP effectors during LR liver development that influence cell shape like Rho GTPases, Rac1 and RhoA. Ectopic expression of Pitx2a in HeLa cells induced the expression of Trio, a protein that interacts with Rac1 and RhoA to reorganize the actin cytoskeleton and strengthen cell to cell adhesive contacts (Wei and Adelstein, 2002). Gelsolin, an actin binding protein that is a key regulator of actin filament assembly and a possible downstream effector of the Wnt/PCP pathway, has

106

Pitx2 binding sites in its promoter region (Eng et al., 2010). Therefore, Rac1, RhoA, Trio, and Gelsolin should be investigated as potential downstream targets of Pitx2c during liver organogenesis that could influence cellular elongation and apical constriction.

During LR liver development, we found asymmetries in apical constriction, cell shape changes, and cellular rearrangements downstream of Pitx2. These processes are regulated by the Wnt/PCP pathway in other organs. Therefore, it is likely that the components of the Wnt/PCP pathway are downstream of Pitx2c during LR liver development. Future research should focus on the role of asymmetric Daam1 expression on liver laterality. Additionally, other components of the Wnt pathway that are asymmetrically expressed in the DM of the gut should be investigated like Frz receptors. There are 11 different Wnt ligands and 8 different Frz receptors that are expressed during liver organogenesis that may provide additional clues about the downstream mechanism of Pitx2c during liver development (Zeng et al., 2006).

4.2) ECM components

The ECM functions as an adhesive substrate for migrating cells, defines tissue boundaries, and presents growth factors to their receptors. Therefore, ECM proteins are ideal targets downstream of Pitx2 in the mesoderm to transfer signals to the endoderm during asymmetric liver morphogenesis. As mentioned previously, asymmetries in the composition of ECM proteins on the left and right side of dorsal mesentery (DM) act downstream of Pitx2 to influence differences in mesenchymal condensation, a requirement for midgut looping

(Kurpios et al., 2008). Another ECM protein, the heparin sulfate proteoglycan, Glypican 3

107

(GPC3), was also found to be expressed solely on the left side of the DM in a microarray screen (Welsh et al., 2014). In fact, there are Pitx2 binding sites in the GPC3 promoter (Eng et al., 2010; Welsh et al., 2014). Interestingly, in humans, there is a correlation between mutations in the GPC3 gene and biliary atresia, a condition commonly found in patients with laterality associated congenital birth defects (Cui et al., 2013). Additionally, GPC3 was found to coordinate interactions between Wnt ligands and their Frizzled receptors in hepatocellular carcinoma (Capurro et al., 2014). Taken together, this suggests that GPC3 could act downstream of Pitx2c to influence Wnt signaling during LR organogenesis of the liver.

Other ECM proteins expressed during liver development include: Nidogen,

Fibrinogen, type IV Collagen, Tenascin, and Laminin (Handa et al., 2014; Si-Tayeb et al.,

2010). Nidogen is a basement membrane ECM protein whose degradation by matrix metalloproteases (MMPs) is required for cellular remodeling events during branching morphogenesis in the developing lung (Warburton et al., 2010). The expression of

Fibrinogen is also important for branching morphogenesis in multiple organs including the lung, mammary, and salivary gland (Warburton et al., 2010). The liver itself is a branching organ as hepatocytes intermingle with mesenchymal and endothelial cells to form liver cords and sinusoids during later stages of morphogenesis. Interestingly, we found LR differences in the number and surface area of liver sinusoids in the developing liver (data not shown).

Therefore, Pitx2c may influence later stages of liver morphogenesis by promoting or inhibiting Fibronectin and Nidogen dependent remodeling events.

There are also Pitx2 binding sites in the promoter region of the ECM protein,

Tenascin-C. Tenascin-C is a nonstructural ECM protein highly expressed during tissue

108 remodeling events in morphogenesis through its effects on cell adhesion, motility, differentiation, growth control, and ECM organization (Imanaka-Yoshida and Aoki, 2014).

Tenascin interacts with multiple cell surface receptors, including integrins and syndecans, to influence cell signaling events (Imanaka-Yoshida and Aoki, 2014). Therefore, it is possible that Tenascin acts downstream of Pitx2 during liver morphogenesis by acting as a signaling mediator between the mesoderm and endoderm.

Laminin is a basement membrane protein required for LR asymmetric gut looping in zebrafish due to its effects on cellular remodeling in the lateral plate mesoderm (Hochgreb and Hagele et al., 2013; Yin et al., 2010). There are Pitx2 binding sites in the Laminin gamma 1 promoter region (Eng et al., 2010). Laminin is also known to signal through

Integrin to activate Rac1 and induce changes in the actin cytoskeleton in embryonic stem cells (Suh and Han et al., 2010). Therefore, possible LR asymmetries in laminin during LR liver development downstream of Pitx2, could influence cell shape change through its effect on Rac1 and the actin cytoskeleton.

During liver morphogenesis, a laminin-rich basement membrane surrounds the developing diverticulum. At the initiation of sinusoid and liver cord formation, this laminin layer is degraded by MMPs so that hepatoblasts can intermingle with surrounding mesenchyme and endothelial cells to form liver cords. Interestingly, there are also Pitx2 binding sites in the promoter region of the MMP inhibitor, Timp2 (Eng et al., 2010). In zebrafish, Timp2a/b expression is required for gut looping (Yin et al., 2010). It is possible that Pitx2c influences laminin deposition, stability, or degradation, or Laminin-Integrin-Rac1 signaling during asymmetric liver development in other vertebrates by directly affecting the

109 expression of laminin, or by influencing the expression of MMPs and MMP inhibitors like

Timp.

4.3) Soluble Growth Factors

ECM proteins also influence the availability of soluble growth factors and can assist in presenting growth factors to their receptors. Many different soluble growth factors are required for proper liver morphogenesis and represent possible downstream targets of Pitx2c during the development of liver laterality. For example, Fibroblast Growth Factor (Fgf) and

Bone Morphogenetic Protein (BMP), expressed in the septum transversum mesenchyme surrounding the liver, are required for the induction of the hepatic gene program (Si-Tayeb et al., 2010). Bmp4 deficient mice have a delay in the expansion of the liver bud, therefore,

Bmp may be required for asymmetric right-sided diverticulum expansion (Si-Tayeb et al.,

2010), however it is unknown if Bmp is expressed asymmetrically within the developing liver. Another growth factor required for liver development, Fgf, is also required for heart development, as Fgf orchestrates ECM formation during outflow tract development.

Interestingly, Pitx2 mutants have reduced Fgf and Bmp signaling in the heart (Ma et al.

2013). During cecum morphogenesis, Fgf10 is downstream of mesenchymally expressed

Pitx2 (Alam et al., 2012). Additionally, Pitx2 and Wnt act synergistically to induce the expression of Fgf16 during ovarian cancer metastasis through Fgf16 regulation of MMPs

(Basu et al., 2014). Fgf signaling is also required for the branching morphogenesis of the lungs, collecting ducts of the kidneys, mammary glands, and salivary glands (Varner and

Nelson, 2014). Therefore, it is possible that Pitx2 promotes or inhibits the expression of Fgf

110 or Bmp ligands in the mesoderm to facilitate signaling to the endoderm during LR asymmetric liver development, including asymmetric liver branching.

Another soluble signaling protein of interest is Hepatocyte Growth Factor (HGF) which is required for liver bud outgrowth. HGF mutant mice show drastic reductions in liver size (Schmidt et al., 1995). HGF is also required for branching morphogenesis in the salivary gland by regulating interactions between the mesenchyme and branching epithelium (Ikari et al., 2003). Interestingly, Pitx2 binding sites have been found in the promoter region of HGF

(Eng et al., 2010). HGF signals through its receptor, MET, to induce Rho mediated cytoskeletal reorganization and may orchestrate cell shape changes during LR liver organogenesis (Parikh et al., 2014). Therefore, soluble growth factors that influence branching events in other organs like Fgf, Bmp, and Hgf are good candidates for factors downstream of Pitx2c during liver development.

4.4) Chemokines

The chemokine, Cxcl12, is a downstream target of Pitx2 during angiogenesis in the dorsal mesentery where it acts to guide the direction of migration of endothelial cells

(Mahadevan et al., 2014). There is a Pitx2 binding site in the promoter region of Cxcl12 (Eng et al., 2010). Cxcl12 also plays a role in pancreatic induction and is expressed during liver hepatocellular carcinoma (Katsumoto and Kume, 2013, Shibuta et al., 2002). Cxcl12 binds to its receptor Cxcr4 to influence Rac/Rho mediated cytoskeletal rearrangements needed for cell migration (Cojoc et al., 2013). During liver development, Cxcl12 is expressed in the developing vascular bed and is known to interact with GAG ECM proteins like GPC3

111

(Franco et al., 2009, Friand et al., 2009). It is possible that Pitx2 induces the asymmetric expression of Cxcl12 in the left mesenchyme to influence liver laterality, but it is more likely that there are asymmetries in vasculature development similar to those seen in the DM that are driven by a Pitx2-Cxcl12 interaction independent of LR diverticulum morphology.

4.5) Conclusions

During liver development, Pitx2 is necessary and sufficient for LR asymmetries in cell shape change and epithelial remodeling required for larger scale morphological laterality

(lobation). There are several genes with Pitx2 binding sites in their promoter regions that are possible downstream targets of Pitx2c (Fig. 4.1). Many of these genes can signal between tissue layers to orchestrate cell shape change and cellular rearrangement through their role in actin cytoskeletal reorganization. For example, Pitx2 may promote or inhibit the transcription of ECM proteins like laminin, GPC3, and Tenascin and/or growth factors like Hgf and Fgf.

Many of the ECM proteins can interact with Wnt, Integrin, or growth factor receptors to mediate signaling and downstream activation of Rho/Rac effectors to influence F-actin. It is unlikely that Pitx2c acts through the canonical Wnt signaling pathway during liver development as we saw no differences in proliferation or differentiation. Future research should focus on describing the expression patterns and requirement of each of these candidate genes during LR liver development.

112

5) Ramifications for human health

As many as 1 in 10,000 human infants are born with defects in LR asymmetry, including defects in liver laterality (Lin et al., 2000). Many individuals with heterotaxia also have the liver positioned at the midline and abnormalities within the biliary system

(Brueckner, 2007). Extrahepatic biliary atresia is a condition in which the extrahepatic bile duct that connects the liver, gall bladder, and intestines, is blocked, occluded, or absent leading to fibrosis and cirrhosis of the liver and biliary tree (Mathur et al., 2014). One form of EHBA is associated with heterotaxia as individuals with polysplenia, asplenia, cardiac defects, annular pancreas, and intestinal malrotation are more likely to also present with

EHBA (Mathur et al., 2014). Heterotaxic mice also develop symptoms associated with

EHBA as neonates become jaundiced and rarely live past one week due to discontinuities of the biliary system and blocked bile flow (Mazziotti et al., 1999; Shimadera et al., 2007).

Therefore, knowledge of the mechanism by which the hepatobiliary system develops asymmetrically is vital to understanding the etiology of EHBA and other heterotaxia associated defects. Identifying downstream targets of Pitx2c during left-right liver development will be valuable as future therapeutic targets. Interestingly, a glycoprotein,

GPC1, has already been identified as a biliary atresia susceptibility gene, further lending credence to the possibility of GPC3 as a downstream target of Pitx2c during LR liver development (Cui et al., 2013).

113

Table 3- 1) Genes with Pitx2 binding sites in the promoter

Gene Function Citation Daam2 Formin; Wnt/PCP pathway; Eng et al., 2010; Welsh et al., Downstream of Pitx2 during gut looping 2014 Gelsolin Actin binding protein, possibly an Eng et al., 2010 effector of the Wnt/PCP pathway GPC3 Glypican; ECM protein; implicated in Eng et al., 2010; Kurpios et biliary atresia al., 2008 Tenascin-C Non-structural ECM protein; signals via Eng et al., 2010; Imanaka- integrins and syndecans; cell adhesion Yoshisa and Aoki, 2014 and motility; heart formation Laminin, ECM protein in the basement membrane; Eng et al., 2010; Hochgreb gamma1 zebrafish gut looping and Hagele et al., 2013; Yin et al., 2010 Timp2 Tissue inhibitor of metalloprotease, Eng et al., 2010; Hochgreb zebrafish gut looping and Hagele et al., 2013; Yin et al., 2010 HGF Hepatocyte growth factor, branching of Eng et al., 2010; Ikari et al., salivary gland; necessary for liver 2003; Schmidt et al., 1995 development CxCl12 Chemokine, asymmetric vasculogenesis Eng et al., 2010; Franco et al., in the DM, liver vascular development, 2009; Katsumoto and Kume, pancreas induction 2013; Mahadevan et al., 2014

Table 3- 2) Direct protein-protein binding partners of Pitx2

Gene Function Citation Nuclear - Wnt signaling effector; gene expression; Basu and Roy, 2012; Sharp catenin proliferation; differentiation; tooth et al., 2014; Vadlamudi et development; cancer al., 2005 Lef-1 Wnt signaling effector; gene expression; Pillai et al., 2015; Sharp et proliferation; tooth development; al., 2014; Vadlamudi et al., differentiation; cancer 2005 mir-200 microRNA; dental epithelial cell Sharp et al., 2014 differentiation

114

Figure 3- 1) Possible downstream targets of Pitx2c There are Pitx2c binding sites in the promoter region of several genes including Cxcl12, Dkk, Wnt, GPC3, Tenascin-C, Laminin, and HGF. Inhibition or promotion of the expression of any of these genes by Pitx2c in the mesoderm (mesenchyme) could influence endodermal (epithelial) morphogenesis in the developing liver diverticulum.

115

References

Alam, D.A., Sala, F.G., Baptista, S., Galzote, R., Danopoulos, S., Tiozzo, C., Gage, P., Grikscheit, T., Warburton, D., Frey, M.R., and Bellusci, S. (2012). FGF9-Pitx2-FGF10 signaling controls cecal formation in mice. Dev. Biol. 369, 340-348.

Auman, H.J., Coleman, H., Riley, H.E., Olale, F., Tsai, H., and Yelon, D. (2007). Functional modulation of cardiac form through confined cell shape changes. PLoS Biol. 5, e53.

Bakkers, J., Verhoeven, M.C., and Abdelilah-Seyfried, S. (2009). Shaping the zebrafish heart: from left-right axis specification to epithelial tissue morphogenesis. Dev Biol. 330, 213-220.

Basu, M., and Roy, S.S. (2013). Wnt/-catenin pathway is regulated by Pitx2 homeodomain protein and thus contributes to proliferation of human ovarian adenocarcinoma cell, SKOV- 3. Journal of Biol. Chem. 288, 4355-4367.

Basu, M., Mukhopadhyay, S., Chatterjee, U., and Roy, S.S. (2014). FGF16 promotes invasive behavior of SKOV-3 ovarian cancer cells through activation of MAPK signaling pathway. J. Biol. Chem. 289, 1415-1418.

Brueckner, M. (2007). Heterotaxia, congenital heart disease, and primary ciliary dyskinesia. Circulation 115, 2793-2795.

Capurro, M., Martin, T., Shi, W., and Filmus, J. (2014). Glypican-3 binds to Frizzled and plays a direct role in the stimulation of canonical Wnt signaling. J. Cell Sci. 127, 1565-1575.

Clevers, H., and Nusse, R. (2012). Wnt/-Catenin signaling and disease. Cell. 149, 1192- 1205.

Cojoc, M., Peitzsch, C., Trautmann, F., Polishchuk, L., Telegeev, G.D., and Dubrovska, A. (2013). Emerging targets in cancer management: role of CXCL12/CXCR4 axis. OncoTargets and Therapy. 6, 1347-1361.

Cui, S., Leyva-Vega, M., Tsai, E.A., Eayclaire, S.F., Glessner, J.T., Hakonarson, H., Devoto, M., Haber, B.A., Spinner, B.B., and Matthews, R.P. (2013). Evidence from

116 human and zebrafish that GPC1 is a biliary atresia susceptibility gene. Gastroenterology. 144, 1107-1115.

Dale, R.M., Sisson, B.E., and Topczewski, J. (2009). The emerging role of Wnt/PCP signaling in organ formation. Zebrafish 6, 9-14.

Davis, A., Amin, N.M., Johnson, C., Bagley, K., Ghashghaei, T., and Nascone-Yoder, N. (2017) Stomach curvature is generated by left-right asymmetric gut morphogenesis. Development.

Eng, D., Campbell, A., Hilton, T., Leid, M., Gross, M.K., and Kioussi, C. (2010). Prediction of regulatory networks in mouse abdominal wall. Gene. 469, 1-8.

Franco, D., Rueda, P., Lendinez, E., Arenzana-Seisdedos, F., and Caruz, A. (2009). Developmental expression profile of the CXCL12 isoform: insights into its tissue-specific role. The Anatomical Record. 292, 891-901.

Frantz, C., Stewart, K.M., and Weaver, V.M. (2010). The extracellular matrix at a glance. J. Cell Sci. 123, 4195-4200.

Friand, V., Haddad, O., Papy-Garcia, D., Hlawaty, H., Vassy, R., Hamma-Kourbali, Y., Perret, G., Courty, J., Baleux, F., Oudar, O., Gattegno, L., Sutton, A., and Charnauz, N. (2009). Glycosaminoglycan mimetics inhibit SDF-1/CXCL12-mediated migration and invasion of human hepatoma cells. Glycobiology. 19, 1511-1524.

Grosse, A.S., Pressprich, M.F., Curley, L.B., Hamilton, K.L., Margolis, B., Hildebrand, J.D., and Gumucio, D.L. (2011). Cell dynamics in fetal intestinal epithelium: implication for intestinal growth and morphogenesis. Development. 138, 4423-4432.

Handa, K., Matsubara, K., Fukumitsu, K., Guzman-Lepe, J., Watson, A., and Soto- Gutierrez, A. (2014). Assembly of human organs from stem cells to study liver disease. American Journal of Pathology. 184, 348-357.

117

Ikari, T., Hiraki, A., Seki, K., Sugiura, T., Matsumoto, K., and Shirasuna, K. (2003). Involvement of hepatocyte growth factor in branching morphogenesis of murine salivary gland. Dev. Dyn. 228, 173-184.

Imanaka-Yoshida, K., and Aoki, H. (2014). Tenascin-C and mechanotransduction in the development and diseases of cardiovascular system. Frontiers in Physiology. 5, 1-12.

Katsumoto, K., and Kume, S. (2013). The role of CXCL12-CXCR4 signaling pathway in pancreatic development. Theranostics. 3, 11-17.

Kioussi, C., Briata, P., Baek, S.H., Rose, D.W., Hamblet, N.S., and Herman, T. (2002). Identification of a Wnt/Dvl/beta-catenin Pitx2 pathway mediating cell-type-specific proliferation during development. Cell. 111, 673-685.

Kosodo, Y. (2012). Interkinetic nuclear migration: beyond a hallmark of neurogenesis. Cell Mol. Life Sci. 69, 2727-2738.

Lecuit, T., and Lenne, P. (2007). Cell surface mechanics and the control of cell shape, tissue patterns and morphogenesis. Nature Reviews Mol. Cell Biol. 8, 633-644.

Lee, W., Chakraborty, P.K., and Thevenod, F. (2013). Pituitary homeobox 2 (Pitx2) protects renal cancer cell lines against doxorubicin toxicity by transcriptional activation of the multidrug transporter ABCB1. International Journal of Cancer. 113, 556-567.

Lin, A.E., Ticho, B.S., Houde, K., Westgate, M.N., and Holmes, L.B. (2000). Heterotaxy: associated conditions and hospital-based prevalence in newborns. Genet. Med. 2, 157-172.

Ma, H.Y., Xu, J., Eng, D., Gross, M.K., and Kioussi, C. (2013). Pitx2 mediated cardiac outflow tract remodeling. Dev. Dyn. 242, 456-468.

Mathur, P., Gupta, R., Soni, V., Ahmed, R., and Goyal, R.B. (2014). Biliary atresia associated with polysplenia syndrome, dextrocardia, situs inversus totalis, and malrotation of intestines. J. Neonatal Surg. 3, 9.

118

Nakaya, M., Habas, R., Biris, K., Dunty Jr., W.C., Kato, Y., He, X., and Yamaguchi, T.P. Identification and comparative expression analyses of Daam genes in mouse and Xenopus. Gene Expr. Patterns. 5, 97-105.

Parikh, R.A., Wang, P., Beumer, J.H., Chu, E., and Appleman, L.J. (2014). The potential roles of hepatocyte growth factor (HGF) - MET pathway inhibitors in cancer treatment. OncoTargets and Therapy. 7, 969-983.

Pillai, S.G., Dasgupta, N., Siddappa, C.M., Watson, M.A., Fleming, T., Trinkaus, K., and Aft, R. (2015). Paired-like homeodomain transcription factor 2 expression by breast cancer bone marrow disseminated tumor cells is associated with early recurrent disease development. Breast Cancer Res. Treat. 153, 507-517.

Pilot, F., and Lecuit, T. (2005). Compartmentalized morphogenesis in epithelia: from cell to tissue shape. Dev. Dyn. 232, 685-694.

Rozario, T., and DeSimone, D.W. (2010). Extracellular matrix in development and morphogenesis: a dynamic view. Dev. Biol. 341, 126-140.

Sawyer, J.M., Harrell, J.R., Shemer, G., Sullivan-Brown, J., Roh-Johnson, M., and Goldstein, B. (2010). Apical constriction: a cell shape change that can drive morphogenesis. Dev. Biol. 341, 5-19.

Schmidt, C., Bladt, F., Goedecke, S., Brinkmann, V., Zschiesche, W., Sharpe, M., Gherardi, E., and Birchneier, C. (1995). Scatter factor/ hepatocyte growth factor is essential for liver development. Nature. 373, 699-702.

119

Sharp, T., Wang, J., Li, X., Cao, H., Gao, S., Moreno, M., and Amendt, B.A. (2014). A pituitary homeobox 2 (Pitx2): microRNA-200a-3p: -catenin pathway converts mesenchymal cells to amelogenin-expressing dental epithelial cells. J. Biol. Chem. 289, 27327-27341.

Shi, G., Sohn, K., Choi, T., Choi, D., Lee, S., Ou, B., Kim, S., Lee, Y., Yoon, T., Kim, S., Lee, Y., Seo, Y., Lee, J., and Kim, C. (2010). Expression of paired-like homeodomain transcription factor 2c (Pitx2c) in epidermal keratinocytes. Exp. Cell Res. 316, 3263-3271.

Shibuta, K., Mori, M., Shimoda, K., Inoue, H., Mitra, P., and Barnard, G.F. (2002). Regional expression of CXCL12/CXCR4 in liver and hepatocellular carcinoma and cell- cycle variation during in vitro differentiation. Jpn. J. Cancer Res. 93, 789-797.

Shimadera, S., Iwai, N., Deguchi, E., Kimura, O., Fumino, S., and Yokoyama, T. (2007). The inv mouse as an experimental model of biliary atresia. J. Pediatr. Surg. 42, 1555-1560.

Si-Tayeb, K., Lemaigre, F.P., and Duncan, S.A. (2010). Organogenesis and Development of the Liver. Developmental Cell. 18, 175-189.

Spiering, D., and Hodgson, L. (2011). Dynamics of Rho-family small GTPases in actin regulation and motility. Cell Adh. Migr. 5, 170-180.

Suh, H.N., and Han, H.J. (2010). Laminin regulates mouse embryonic stem cell migration involvement of Epac1/Rap1 and Rac1/cdc42. Am. J. Physiol. Cell. Physiol. 298, 1159-1169.

Vadlamudi, U., Espinoza, H.M., Ganga, M., Martin, D.M., Liu, X., Engelhardt, J.F., and Amendt, B.A. (2005). Pitx2, -catenin, and LEF-1 interact to synergistically regulate the LEF-1 promoter. J. of Cell Science. 118, 1129-1137.

Varner, V.D., and Nelson, C.M. (2014). Cellular and physical mechanisms of branching morphogenesis. Development. 141, 2750-2759.

Warburton, D., El-Hashash, A., Carraro, G., Tiozzo, C., Sala, F., Rogers, O., De Langhe, S., Kemp, P.J., Riccardi, D., Torday, J., Bellusci, S., Shi, W., Lubkin, S.R., and Jesudason, E. (2010). Lung Organogenesis. Curr. Top. Dev. Biol. 90, 73-158.

Wei, Q., and Adelstein, R.S. (2002). Pitx2a expression alters actin-myosin cytoskeleton and migration of HeLa cells through Rho GTPase signaling. Mol. Biol of the Cell. 13, 683-697.

120

Yang, Y. (2012). Wnt signaling in development and disease. Cell Biosci. 2, 14

Zeng, G., Awan, F., Otruba, W., Muller, P., Apte, U., Tan, X., Gandhi, C., Demetris, A.J., and Monga, S.P.S. (2006). Wnt’er in liver: expression of Wnt and Frizzled genes in mouse. Hepatology. 45, 195-204.

Zorn, A.M., and Wells, J.M. (2009). Vertebrate endoderm development and organ formation. Annu Rev Cell Dev. Biol. 25, 221-51.

121

CHAPTER 4: Conclusions and Future Directions

Many internal organs of the body exhibit LR asymmetry in morphology including the liver. We found that left-limited expression of Pitx2c orchestrates LR differences in cell shape and epithelial remodeling required for liver laterality. Pitx2 is a known downstream target of the LR asymmetry pathway and is required for cell shape changes and cell rearrangements during midgut looping and stomach curvature. Our study is the first to show that cellular level morphological asymmetries downstream of Pitx2c are required for LR liver development. Our findings reveal a conserved role for Pitx2c in orchestrating laterality in multiple organ systems while also uncovering surprising differences in the downstream action of Pitx2c based on the tissue layer of expression (whether mesodermal or endodermal). In this manner, Pitx2c directs the asymmetrical development of organs with varying geometric configurations, from looping or curving tubes to asymmetric branching.

We hope that future work will expand on downstream targets of Pitx2c during liver morphogenesis as well as link these finding to other unexplored asymmetric lobated organs like the lungs and pancreas.

Uncovering downstream targets of Pitx2c during LR liver development will be vital to increasing our understanding of the mechanisms required during embryogenesis. Our laboratory is currently investigating downstream targets of Pitx2c during stomach and heart development using RNA sequencing of the left and right halves of organs dissected from the large Lepidobatrachus laevis frog embryo. Unfortunately, liver morphology in this species is relatively symmetrical, perhaps due to a posterior shift in the domain of Pitx2c expression and therefore cannot be used for identifying targets in the liver. Another possibility is to use

122 laser capture microdissection of multiple X. laevis cross sections for either RNA sequencing or microarray gene expression profiling. These experiments will provide likely downstream targets of Pitx2c during liver development. Researchers have identified possible Pitx2c candidate genes in other LR organs including the dorsal mesentery of the midgut. Identifying the expression pattern of genes like GPC3, Daam1/2, and N-cadherin within the developing liver and in embryos with manipulated Pitx2c expression will give us clues as to their involvement in LR liver development (Welsh et al., 2014). Other genes like HGF, Tenascin, and Laminin with Pitx2 binding sites in their promoter regions should be investigated for asymmetric expression (Eng et al., 2010). Additionally, cytoskeletal components like actin microfilaments and microtubules as well as motor proteins like Myosin II play roles in orchestrating cell shape changes and epithelial remodeling in many other asymmetric organs and during liver laterality in zebrafish (Cayuso et al., 2016; Kurpios et al., 2008). Therefore the distribution and localization of these proteins should be analyzed in liver diverticulum cross sections using immunohistochemistry with and without Pitx2c manipulation.

We found LR differences in epithelial character during liver diverticulum development as the right side increases surface area by remaining a pseudostratified epithelium while the left side became a multiple-layered stratified epithelium. The mechanism by which these epithelial differences occur is still unknown. Using BrdU labeling experiments, we can determine if differences in interkinetic nuclear migration or the reincorporation of post-mitotic cells contributes to these differences or if cell shape changes alone can account for remodeling events. By culturing live cross sections of the liver diverticulum and tracking labeled cells, we could also determine how cells are remodeling in

123 real time and how misexpression of Pitx2c affects these cellular movements and cell shape changes.

The anatomical morphology of the liver is strikingly different from other LR asymmetric organs studied like the tubular stomach and midgut. The liver is a multi-lobated organ with branching morphology similar to the lungs. In humans, the right side of the liver is larger than the left and is composed of more lobes; four lobes on the right and two on the left (Abdel-Misih and Bloomston, 2010; Gray, 1918). Interestingly, the right lung is also larger than the left and has three lobes while the left side only has two (Gray, 1918). Pitx2 is required for the development of lung laterality but the mechanism by which this asymmetry is established is unknown (Campione et al., 1999, Kitamura et al., 1999). It will be interesting to determine whether Pitx2c is expressed in mesoderm, endoderm, or both within the developing lung bud. I would hypothesize that Pitx2c is expressed in just the mesoderm surrounding the left side of the lung during development and that cellular level asymmetric morphogenesis is similar to our observations in the liver diverticulum.

Pitx2 is a conserved protein required for the development of asymmetric organ systems in a variety of animals including mouse, chicken, Xenopus, and even echinoderms

(Blum et al., 2014; Duboc et al., 2005; Kitamura et al., 1999). The requirement of left-sided

Pitx2c expression during liver development should be investigated in other model systems like the mouse. Hepatobiliary morphology is strikingly diverse within the animal kingdom as the size and number of lobes is species dependent. Horses have fairly symmetrical liver morphology and do not develop gall bladders. As mentioned previously, the liver of the

Lepidobatrachus laevis tadpole appears more symmetrical as well and may provide a more

124 amenable model system to study naturally divergent mechanisms of liver development.

Interestingly, the dogfish shark, shows asymmetries in liver morphology, however, the left lobe of the liver is larger and more elongate than the right. It would be interesting to determine if any differences exist in the expression or requirement of Pitx2c during L. laevis,

X. laevis, horse, and dogfish liver development that could account for these discrepancies in morphology. This information may provide clues as to the evolutionary significance of liver laterality.

Heterotaxia of the liver is associated with extrahepatic biliary atresia in humans and leads to substantial morbidity and hardship (Gottschalk et al., 2016; Zhu et al., 2005).

Knowledge pertaining to the development of LR asymmetry in the liver is vital to our understanding of the etiology of heterotaxia linked defects like EHBA. Our study focuses on

LR asymmetries in liver diverticulum development, however, future work should determine how Pitx2c and liver laterality influences the formation of the gall bladder and its associated biliary ducts and networks. Additionally, any downstream targets of Pitx2c during LR development of the hepatobiliary system could provide future avenues for developing therapeutic drugs and treatments.

125

References

Abdel-Misih, S.R.Z., and Bloomston, M. (2010). Liver Anatomy. Surg Clin North Am. 90, 643-653.

Blum, M., Schweickert, A., Vick, P., Wright, C.V.E., and Danilchik, M.V. (2014). Symmetry breakage in the vertebrate embryo: When does it happen and how does it work? Dev. Biol. 393, 109-123.

Duboc, V., Rottinger, E., Lapraz, F., Besnardeau, L., and Lepage, T. (2005). Left-right asymmetry in the sea urchin embryo is regulated by nodal signaling on the right side. Dev. Cell. 9, 147-58.

Eng, D., Campbell, A., Hilton, T., Leid, M., Gross, M.K., and Kioussi, C. (2010). Prediction of regulatory networks in mouse abdominal wall. Gene. 469, 1-8.

Gray, H. (1918). Anatomy of the Human Body. Philadelphia: Lea & Febiger, Bartleby.com, 2000. www.bartleby.com/107/.

126

Zhu, L., Belmont, J.W., and Ware, S.M. (2005). Genetics of human heterotaxias. Eur. J. Hum. Genet. 14, 17-25.

127

APPENDICES

128

Appendix A: Budgett’s frog (Lepidobatrachus laevis): a new amphibian embryo for developmental biology

Nirav M. Amin1, Mandy Womble1, Cris Ledon-Rettig2, Margaret Hull1, Amanda Dickinson3,

Nanette Nascone-Yoder1,*

1Department of Molecular Biomedical Sciences 1060 William Moore Drive College of Veterinary Medicine North Carolina State University Raleigh, NC, 27607 USA

2Department of Biology Indiana University 915 E. Third St. Bloomington, IN 47405

3Biology Department Virginia Commonwealth University 1000 W. Cary St. Richmond, VA 23284

*Author for correspondence (email: [email protected]), phone: (919) 513-8284

Published in Dev. Biol. 2015 Sep 15: 405 (2): 291-303. doi: 10.1016/j.ydbio.2015.06.007

129

Abstract

The large size and rapid development of amphibian embryos has facilitated ground- breaking discoveries in developmental biology. Here, we describe the embryogenesis of the

Budgett’s frog (Lepidobatrachus laevis), an unusual species with eggs that are over twice the diameter of laboratory Xenopus, and embryos that can tolerate higher temperatures to develop into a tadpole four times more rapidly. In addition to detailing their early development, we demonstrate that, like Xenopus, these embryos are amenable to explant culture assays and can express exogenous transcripts in a tissue-specific manner. Moreover, the steep developmental trajectory and large scale of Lepidobatrachus make it exceptionally well-suited for morphogenesis research. For example, the developing organs of the Budgett’s frog are massive compared to those of most model species, and are composed of larger individual cells, thereby affording increased subcellular resolution of early vertebrate organogenesis. Furthermore, we found that complete limb regeneration, which typically requires months to achieve in most vertebrate models, occurs in a matter of days in the

Budgett’s tadpole, which substantially accelerates the pace of experimentation. Thus, the unusual combination of the greater size and speed of the Budgett’s frog model provides inimitable advantages for developmental studies—and a novel inroad to address the mechanisms of spatiotemporal scaling during evolution.

130

Introduction

Amphibians have historically been ideal model organisms for experimental embryology because their relatively large, rapidly-developing embryos are abundant and highly accessible to microsurgical manipulations. Indeed, classical experiments on frog and salamander embryos profoundly influenced our current understanding of fundamental events in vertebrate development (Spemann and Mangold, 1924). In the twentieth century, the frog

Xenopus laevis ascended as the amphibian of choice for modern embryologists, largely because of its responsiveness to year-round, hormone-induced ovulation in the laboratory

(Callery, 2006). Xenopus explant assays transformed our conception of multi-potency and the cell behaviors that drive morphogenesis (Logan and Mohun, 1993; Saint-Jeannet et al., 1994;

Sater and Jacobson, 1989; Wilson et al., 1989), while the capacity to microinject single blastomeres with molecular reagents continues to serve as a powerful system for tissue- specific analyses of gene function and the elucidation of regulatory networks (Amaya et al.,

1991; Hopwood and Gurdon, 1990; Moody, 1987a; Moody, 1987b; Smith et al., 1993;

Tandon et al., 2012; Woodland and Jones, 1987). In more recent years, genome resources have become available for X. laevis and its genetically tractable cousin, X. tropicalis

(Hellsten et al., 2010). Consequently, even more sophisticated technologies continue to be added to the amphibian toolbox, including transgenic approaches (Hamlet et al., 2006; Kroll and Amaya, 1996; Ogino et al., 2006; Yergeau et al., 2009), targeted mutagenesis/genome editing via zinc finger nucleases, TALENs, and CRISPRs, and transcriptomic and proteomic profiling (Amin et al., 2014; Blitz et al., 2013; Guo et al., 2014; Lei et al., 2012; Liu et al.,

131

2014; Nakajima and Yaoita, 2013; Nakayama et al., 2013; Rao et al., 2014; Sakane et al.,

2014; Sun et al., 2014; Suzuki et al., 2013; Wuhr et al., 2014; Young et al., 2011)

In addition to Xenopus, and a few other species whose embryology has been described by evolutionary developmental biologists (del Pino et al., 2004; del Pino et al.,

2007; Moya et al., 2007; Perez et al., 2009; Romero-Carvajal et al., 2009), there are about

6,500 other recorded species of frogs and toads with tremendous diversity, but there is almost no information on their development (AmphibiaWeb, 2014). Ironically, as one of the most highly derived genera, Xenopus is arguably not representative of most anurans (Hall, 1992).

Given that hundreds of frogs are now threatened with extinction (Stuart et al., 2004), it is imperative that the biodiversity of amphibian embryogenesis be more fully assessed, not only to inform conservation efforts, but also because many species may be uniquely suited for illuminating critical issues at the interface of development, ecology and evolution (Calboli et al., 2011). For example, the amazing capacity of urodele amphibians to regrow multiple adult organs and tissue types has long been utilized as a system in which to illuminate the mechanisms of regeneration (Brockes and Kumar, 2002). Likewise, studying the distinct ontogeny of direct-developing frogs (e.g., Eleutherodactylus) has provided insight into the evolutionary origins of the amniotic egg during the rise of terrestrial vertebrates (Elinson and

Beckham, 2002). Finally, the exceptional developmental plasticity of Spadefoot toad tadpoles (e.g., Spea) has served as a striking example of the dynamic interplay between the environment and the genome in the manifestation of phenotype (Ledon-Rettig and Pfennig,

2011). In the context of technological advances that now facilitate “omic”-era approaches and enable genome editing in such species (Fei et al., 2014; Flowers et al., 2014; Khattak et

132 al., 2014; Sobkow et al., 2006; Whited et al., 2012), the time is ripe to redouble efforts to leverage the unique biology of additional frogs.

One intriguing candidate species is Lepidobatrachus laevis (known in the pet trade as

“Budgett’s frog”), a robust aquatic frog that lives in the Chaco region of South America

(Budgett, 1899). L. laevis adults have enormous mouths and are aggressive and cannibalistic predators (Hanken, 1993; Fabrezi and Lobo, 2009; Ruibal and Thomas, 1988). The larvae of

L. laevis have been studied because they progress extremely rapidly through metamorphosis, and exhibit an unusual feeding strategy for anuran tadpoles: obligate carnivory (Bloom et al.,

2013; Fabrezi, 2006; Parker, 1931; Ruibal and Thomas, 1988; Ziermann et al. 2013; Hanken

1993). L. laevis larvae consume live prey, including other tadpoles, whole; the craniofacial and digestive tract specializations associated with this megalophagic (and often cannibalistic) feeding ecology have made L. laevis larvae captivating subjects for evolutionary biologists

(Bloom et al., 2013; Carroll et al., 1991; Fabrezi, 2006; Hanken, 1993; Parker, 1931; Ruibal and Thomas, 1988; Ziermann et al., 2013), but the pre-feeding stages of L. laevis development have not yet been formally described.

Here, we document key stages in the early pre-feeding embryogenesis of L. laevis and demonstrate, through the application of standard methodologies, that they also possess the experimental amenability that has made Xenopus embryos so valuable. We provide evidence that the more expedient oocyte size and thermal tolerance of L. laevis provides practical advantages over existing vertebrate embryo models, including enhanced resolution of early organogenesis, and an accelerated time frame for regeneration. We posit that the unusual combination of “extreme” features in L. laevis could provide a novel inroad to address the

133 cellular and molecular mechanisms that influence the pace and scale of morphogenesis during evolution.

Materials & Methods

Lepidobatrachus laevis embryo collection

Adult L. laevis frogs were obtained through the pet trade (Backwater Reptiles), and housed in de-chlorinated tap water at 28°C in individual tanks (to avoid cannibalism within the adult colony). Embryos were produced by natural matings between hormonally-induced male and female frogs; breeding pairs were size-matched to minimize potential aggression.

Briefly, ovulation was induced by injection of 50 g luteinizing hormone (LHRH; Sigma

L7134) into the dorsal lymph sac of the female frog. Two hours later, 30 g LHRH was injected into the dorsal lymph sac of the male. The male was then placed immediately into a

10 gallon mating tank, followed by the larger female (to discourage cannibalism), and the tank was covered with a dark towel. Amplexus was typically observed within one to two hours of placing the male and female together, and ovulation usually commenced within four hours of injecting the female with LHRH. In tanks with amplexing pairs, the fertilized eggs were collected periodically over a 4 hour period.

Fertilized embryos were carefully sorted and rinsed in 10% Holtfreter’s solution (5.9 mM NaCl, 67 M KCl, 76 m CaCl2, 240 M NaHCO3), then transferred to 10 cm petri plates containing fresh 10% Holtfreter’s at 28°C. To avoid overcrowding and promote uniform development, embryos were cultured at a density of <30 embryos per 100 mm plate.

Embryos were staged according to standard amphibian staging criteria (Gosner, 1960).

134

To monitor embryonic development at different temperatures, fertilized X. laevis and

L. laevis embryos were collected at the 2-cell stage and sorted into dishes of 10% Holtfreter’s solution, which were incubated at five different temperatures (16°C, 23°C, 28°C, 32°C,

37°C). For this analysis, 20 X. laevis and 10 L. laevis embryos were scored for viability and growth rate at each temperature. Time-lapse imaging of X. laevis and L. laevis embryos was performed on a Zeiss SteREO Lumar.V12 with AxioVision and compiled into movies using

ImageJ software, version 1.45S (NIH).

Animal cap assays

L. laevis animal caps were dissected at Gosner stage 8-8.5 (Gosner, 1960) using sharpened Watchmaker’s forceps, and cultured at 28°C in 2 mL of 2 ng/ml human activin A

(R&D Systems) in 0.75X MMR (75 mM NaCl, 1.5 mM KCl, 0.75 mM MgSO4, 1.5 mM

CaCl2, 3.75 mM HEPES, 75 M EDTA, pH 7.8) for 14 hours in agarose coated wells of a

24-well plate. Untreated caps were used as controls. For comparison, X. laevis caps were also dissected and cultured in the presence or absence of 2 ng/ml activin, as previously described

(Dush et al., 2011). Caps were fixed in 4% paraformaldehyde (PFA) and imaged at 14 hours post dissection.

Microinjection

L. laevis embryos from natural matings were de-jellied manually, or incubated for 5-7 minutes in 2% cysteine hydrochloride (pH 7.8-8.1), and microinjected at the indicated stages.

Injections were performed in 3% Ficoll dissolved in 10% Holtfreter’s solution. The

135 mMessage transcription kit (Ambion) was used to generate capped mRNA for eGFP (120 pg/nl), which was combined with 2 ng/nl fluorescent dextran-(Alexa555), and back-loaded into pulled, calibrated glass pipettes (Sive et al., 1998); 2 nl of this mixture was injected at the stages described. Embryos were allowed to recover in 3% Ficoll for 20 minutes, and then transferred to individual wells of a 12-well plate containing 10% Holtfreter’s solution for longer term incubation. Fluorescence was imaged at 48 hpf using a Zeiss SteREO

Lumar.V12 with AxioVision.

Immunostaining

Immunohistochemical staining was performed as previously described for Xenopus

(Dush, 2011). Whole embryos were fixed with eight consecutive washes of Dent’s Fixative

(80% Methanol, 20% DMSO) and stored overnight at -20°C. Embryos were rinsed three times with PBST before transfer to sucrose/gelatin (15% sucrose/25% cold water fish gelatin) and stored overnight at room temperature. Embryos were embedded in OCT (Tissue-Tek), sectioned at 10 μm using a Leica CM 1850 cryostat and mounted on coated slides (Fisher

Superfrost). Sections were post-fixed for two minutes in 4% PFA [100 mM Hepes (pH 7.4),

100 mM NaCl, 4% paraformaldehyde] and blocked for 30 minutes as previously described

(Reed, 2009). Antibodies were applied overnight at 4°C in an antibody amplifier humidity chamber (Prohisto). Dilutions of primary antibodies used are as follows: anti--catenin

(Santa Cruz, sc-7199, 1:100), anti--tubulin (abcam, ab270714, 1:1000), anti--tubulin

(Sigma, T9026, 1:1000), and anti-F-actin (Cytoskeleton, AAN01, 1:200). Slides were then washed three times in PBST for 5 minutes, incubated for 3 hours in blocking buffer

136 containing Alexa 488-conjugated goat anti-mouse IgG (Invitrogen, A11029; 1:2000) and/or

Alexa 546-conjugated goat anti-rabbit IgG (Invitrogen, A11035; 1:2000). To stain nuclei, sections were washed twice in PBST, twice with PBS and incubated for 5 minutes in PBS containing TO-PRO-3 (Invitrogen, T3605, 1:1000). Stained slides were washed twice with

PBS and mounted with coverslips and ProLong Gold Antifade Reagent (Invitrogen) before imaging on a Leica (Model TCS-SPE) confocal microscope. Maximum projections of z- stacks are shown.

Limb Regeneration

Tadpoles (48 hpf) were maintained in large tanks and fed on siblings ad libitum.

Individual tadpoles were anesthetized by immersion in 0.05% tricaine methanesulfonate

(MS-222). Once anesthetized, the right limb was amputated just distal to the geniculate joint in a straight line with sharp iridectomy scissors (Dent, 1962; Goode, 1967). The tadpole was monitored for hemostasis before anesthetic recovery. Operated tadpoles were then cultured individually and fed Xenopus tadpoles. Regenerating limbs were imaged on a Zeiss SteREO

Lumar.V12 with AxioVision every 24 hours for at least three weeks.

Results

Lepidobatrachus laevis developmental staging

Below, we include an account of normal L. laevis development, with a focus on the critical phases of early ontogeny (cleavage, gastrulation, neurulation and organogenesis), as compared with Xenopus laevis. We have chosen to describe L. laevis normal development at

137

28°C, a temperature that falls within the overlap of the published ranges of thermal tolerance of embryos, larvae and adults of this species (Carroll, 1996). As X. laevis embryos do not survive at this temperature (Khokha et al., 2002), Figure 1 compares the schedule of L. laevis development at 28°C with that of X. laevis embryos raised at 23°C, as described in the widely-used normal table of X. laevis development (Nieuwkoop and Faber, 1994); however, it should be noted that L. laevis and X. laevis develop at the same rate when raised at mutually permissive temperatures (e.g., 23°C; see Figure 5).

Cleavage

The L. laevis egg (Figure 2A) is relatively large (~2.6 mm) compared to that of X. laevis (~1.2mm). Nonetheless, the early L. laevis embryo appears to undergo holoblastic cleavage (albeit with larger vegetal than animal blastomeres), and requires only 15-20 minutes between divisions. The orientation of cleavage planes is analogous to X. laevis

(Figure 2B-E) and largely synchronous among different individuals through the 32-cell stage, i.e., Gosner (GS) stages 1-7, after which the orientation of these divisions is less uniform. L. laevis embryos reach mid-cleavage stage within 4 hours post fertilization (hpf, Figure 2F) and most reach late blastula stages by 5-6 hpf (Figure 2G-H). Thus, despite the increased size of the egg in L. laevis (see Figure 2I), cleavage can still occur rapidly.

Gastrulation

Gastrulation commences in L. laevis with the appearance of an arc of pigmentation demarcating the involuting dorsal lip at GS10 (Figure 2J). The arc then expands

138 circumferentially to form a complete circle delimiting the yolk plug and future blastopore at

GS11 (Figure 2K). These events are reminiscent of initial gastrulation in Xenopus species.

However, whereas in Xenopus the entire vegetal endoderm is gradually subsumed into the interior of the embryo, the endoderm mass of L. laevis embryos remains slightly protuberant during gastrulation (Figure 2L). Interestingly, despite the substantially larger diameter and potential physical hindrance of this protruding yolk plug, blastopore closure ensues rapidly and is complete by 10 hpf (see Figure 3A).

Neurulation and Organogenesis

During neurulation, L. laevis forms a broad, keyhole-shaped neural plate (Figure 3A).

Despite the larger scale, subsequent formation of neural folds (Figure 3B) and closure of the neural tube (Figure 3C) takes only ~6hrs (GS13-GS16), and migratory streams of neural crest in the prospective pharyngeal arches become obvious even before the neural tube is closed, as early as ~12hpf (G14; see arrows, Figure 3B). These detailed features of neural tube development are highly discernable in L. laevis (Supplemental Movie 1). Upon neural tube closure (~16hpf), the somites become visible (see Figure 3C). The embryo immediately begins elongating along the anterior-posterior axis (Figure 3D-E) and “hatches” out of the surrounding thick jelly layers. At this stage, the embryo begins to exhibit a motor response to external stimuli.

The formation of the major craniofacial structures, including the cement gland, eyes, nasal pit and gill buds, begins within a few hours of neural tube closure (Figures 3D-H, 4).

Initially, the pharyngeal arches appear as protruding pouches at GS17-18 (18-20 hpf; Figure

139

4A-B), which form external gill buds a few hours later (Figure 4D-E), and then quickly undergo branching morphogenesis to form elaborate gills by 28 hpf (Figures 3F-H, 4G-K).

Along with the pharyngeal arches, the stomodeum becomes obvious as a dorsoventrally elongated invagination at the midline of the face (Figure 4). By 28 hpf, the buccopharyngeal membrane has already perforated (see Figure 4H-I) to allow the embryonic mouth to be connected to the foregut. By GS20, the nasal pits have formed (Figure 4D), and by GS21 (27 hpf), the optic cup has expanded into a horseshoe shape with an evident lens (Figure 4G). We note that the L. laevis craniofacial structures are substantially larger and more prominent than in X. laevis. Indeed, individual cells in the face are readily apparent under only a dissecting microscope (e.g., see Figure 4C, F, I, L).

Along with craniofacial morphogenesis, other organogenesis events also commence shortly after neurulation in L. laevis. The pronephros is evident immediately following neural tube closure (Figures 3D-F). Heartbeat commences before the end of the first day (~21 hpf;

GS19), with looping and septation complete by ~24hpf, and functional blood circulation visible in the gills and fins by 27-28 hpf (GS20-22; not shown). Morphogenesis of the digestive tract is also evident before the end of the first day (Figure 3H), with overt gut looping and pre-feeding intestinal lengthening concluded by ~36-40 hpf [not shown, but see

Bloom et al., 2013]. L. laevis tadpoles have been observed to begin feeding on their siblings as early as 48 hpf, indicating that the gut is fully functional by two days post fertilization at

28°C. Thus, despite the larger size, organogenesis proceeds at a rapid rate in the L. laevis embryo.

140

L. laevis embryos can tolerate a wide range of temperatures

The ability to accelerate or decelerate the developmental trajectory of laboratory

Xenopus embryos by shifting them to higher or lower temperatures, respectively, has enabled experimental manipulation of any stage of development at convenient times of the day.

However, at temperatures outside of a species-specific optimal range, accelerated or decelerated embryos display developmental defects and decreased viability (Khokha et al.,

2002). Although L. laevis embryos and larvae have been reported to enjoy a broad range of thermal tolerance (Carroll, 1996), the morbidity of embryos grown at different temperatures in the laboratory has not been explicitly described.

To determine the optimal temperature range for normal morphological development of L. laevis, we raised L. laevis and X. laevis embryos in parallel at temperatures ranging from 16°C to 37°C. At the lowest temperature tested (16°C), X. laevis embryos developed normally, while all the L. laevis embryos failed to complete gastrulation. However, at 23°C, the embryos of both species developed with normal morphology and, despite their size difference, at a similar pace (Figure 5; Supplemental Movie 2). This suggests that the lower temperature limit for L. laevis development is between 16°C and 23°C.

At 28°C, both species cleaved at a similar rate. However, the majority of the X. laevis embryos (55%; n=20) failed to gastrulate and the remainder developed severe ventral edemas by GS17, indicating that the upper temperature limit of X. laevis development is below 28°C, consistent with previous results (Khokha et al, 2002). In contrast, all L. laevis embryos developed normally at 28°C, as expected. Moreover, L. laevis embryos continued to thrive at temperatures that cause all X. laevis embryos to die before completing gastrulation (32°C and

141

37°C; Figure 5). Strikingly, L. laevis develops at a dramatically accelerated pace at 37°C, reaching feeding tadpole stages in less than a day—about four times as fast as X. laevis raised at 23°C—with no obvious defects in development. Overall, these results demonstrate that, like Xenopus species, L. laevis is amenable to temperature-mediated manipulation of developmental rate for experimenter convenience.

L. laevis enables lineage tracing in live embryos at high resolution and accuracy

Xenopus blastulae often exhibit patterns of cleavage and pigmentation that enable specific lineages to be targeted for microinjection. For example, future ventral blastomeres can often be identified by their darker pigmentation and slightly larger size in comparison to the lighter and slightly smaller dorsal blastomeres (Moody, 1987a; see Figure 2D-E).

Lepidobatrachus blastulae exhibit similar dorsal-ventral asymmetries and apparent holoblastic cleavage. However, in other frog species with large eggs, it has been found that germ layer positions, and thus blastomere fate maps, may be altered (i.e., shifted along the animal-vegetal axis), or that the early cleavage divisions remain incomplete (Elinson and

Beckham, 2002; Moya et al., 2007). If such features exist in L. laevis, this would limit the utility of this species for lineage-specific assays.

To determine whether specific tissues can be reliably targeted by microinjection in L. laevis, we microinjected fluorescent dextran into one blastomere of a L. laevis embryo at the

2-cell stage. In Xenopus species, the first cleavage generally corresponds to the embryonic midline; thus, injecting one cell at the 2-cell stage results in mostly unilateral localization of the fluorescent lineage tracer to either the left or right side of the embryo (Ramsdell et al.,

142

2006). Likewise, in L. laevis, this injection resulted in robust fluorescence limited predominantly to one side of the animal (Figure 6A-B). Thus, despite their larger scale, the first cleavage plane also corresponds generally with the axial midline of L. laevis.

To ascertain whether fate maps of later stage X. laevis blastomeres (e.g., at the 16-32- cell stage) could also be used to target specific tissues in L. laevis, we microinjected fluorescent dextran into analogous later stage blastomeres in L. laevis. These injections yielded a similar distribution of the fluorescent label in L. laevis as previously reported in

Xenopus, with high reproducibility (Figure 6D-F; 75% targeting the foregut, n=8). Moreover, we found that L. laevis blastomeres are easily injectable up to at least the 128-cell stage

(Figure 6G-I), potentially allowing lineage tracing of later stage fates and, thus, finer targeting of more discrete tissues in the larger embryo. Importantly, co-injection of synthetic capped mRNA encoding eGFP resulted in strong green fluorescence in the progeny of the injected blastomeres (Figure 6C). These results show that L. laevis embryos are not only amenable to fate mapping studies with lineage tracers, but are also able to express exogenous mRNA for highly targeted gain of function studies.

L. laevis animal caps respond to external stimuli similarly to Xenopus

A key technical advantage of amphibian embryos is the ability to culture explants of embryonic tissues in vitro (Logan and Mohun, 1993; Saint-Jeannet et al., 1994; Sater and

Jacobson, 1989; Sater and Jacobson, 1990; Wilson et al., 1989; Woodland and Jones, 1987).

X. laevis has long been exploited for the “animal cap” assay, in which a disc of naïve ectoderm tissue from the animal pole of the GS8-9 embryo is induced to form an alternate

143 fate (e.g., mesoderm) by culturing it in the presence of biologically active molecules, such as the TGF- ligand, activin. An easily observable readout of the induction of mesoderm is the elongation of the isolated tissue (which normally forms a ball of ciliated epidermis in vitro), reflecting the convergent extension rearrangements that are executed by axial/paraxial mesoderm during gastrulation (compare Figures 7A and C; Brieher and Gumbiner, 1994;

Dush et al., 2011; Smith et al., 1990). However, considering the more sizeable dimensions of the ectoderm in L. laevis, and the potential for accelerated execution of its morphogenetic processes, it is unclear how Budgett’s frog animal caps might respond to in vitro explant culture.

To test this response, we dissected animal caps from GS8-9 embryos and cultured them in the presence or absence of 2ng/ml activin. Although animal caps from the L. laevis embryo are equivalent in diameter to an entire X. laevis blastula (but smaller caps may be cultured, e.g., rightmost cap in Figure 7B), they nonetheless quickly heal into spherical shapes that eventually differentiate into ciliated epidermis (Figure 7B), similar to untreated X. laevis caps. Moreover, when exposed to activin, these explants rapidly undergo obvious convergent extension (Figure 7D). Thus, as in X. laevis, L. laevis tissues are highly amenable to explant assays.

A high resolution view of organ development in L. laevis

Because L. laevis embryos are considerably bigger than X. laevis, they provide outstanding resolution of the emergence of gross morphology, such as neural folds and craniofacial features. However, they also possess relatively enormous internal organs. For

144 example, at ~0.6 mm in length, the early Lepidobatrachus heart tube is almost as long as an entire neurula stage X. laevis embryo (Figure 8A). Likewise, the looped heart of a L. laevis tadpole is substantially larger than a comparable stage X. laevis heart (Figure 8B). These observations suggest that the increased scale of L. laevis organs could provide greater cellular resolution of organogenesis events than X. laevis.

To assess this possibility, we characterized tissue architecture during L. laevis heart morphogenesis by applying a panel of antibodies known to detect Xenopus proteins, including markers of cell-cell adhesion (-catenin) and the cytoskeleton (/-tubulin, actin)

(Figure 8C-J). We note that, with the increased scale of the larger embryo, a cross section of only the heart of L. laevis is approximately the same size as an entire Xenopus embryo (100x magnification; compare Figure 8C and D). Consequently, the distinction between the myocardial and endocardial tissue layers, including individual cell shapes, is more evident in

L. laevis heart sections at low magnification (compare Figure 8C and D).

Interestingly, comparing the dimensions of individual cells in L. laevis and X. laevis hearts at a similar stage of cardiogenesis revealed that differentiating cells in the L. laevis organ are actually about 50% larger (1.46X, p<0.001 for myocardium; 1.47X, p<0.0001 for endocardium) than those that comprise a comparable stage X. laevis heart. Fortuitously, this increase in cell size enables discernment of subcellular organization in the endocardial versus myocardial layers at even early heart tube stages (Figure 8H-J). For example, standard confocal microscopy at only moderately high (400X) magnification can be used to visualize the orientation of microtubule architecture (Figure 8G-H), the position of centrioles (Figure

8I-J), the apical enrichment of adherens junction components (Figure 8G-H) and the

145 polarization and alignment of bundles of actin filaments (Figure 8I-J). Thus, the inherently greater cell size of L. laevis embryonic organs yields exceptional resolution of very early stages of vertebrate organogenesis.

Limb Regeneration in L. laevis occurs rapidly

Given the rapid development attainable in L. laevis, we reasoned that the process of regeneration might also be accelerated. However, not all frogs show regenerative capacity

(Scadding, 1977). To test if L. laevis would be an appropriate frog model for accelerated regeneration studies, we amputated the hind limb of L. laevis tadpoles at a stage of limb development comparable to that typically used in X. laevis (8 dpf; Figure 9A; Beck, 2012).

By 24 hours, a blastema was visible (Figure 9B), and a fully regenerated limb (Figure 9C) was observed less than 10 days post-amputation (80%; n=5). Notably, by 21 days post amputation, the regenerated L. laevis limb was indistinguishable from the unoperated limb in both pattern and size (data not shown)—this can take months to achieve in other species, such as axolotl (Goode, 1967; Tank et al., 1976; Young et al., 1983).

In Xenopus and other frogs, regeneration capacity diminishes as tadpoles approach metamorphosis (Dent, 1962). L. laevis also possesses this plasticity, as amputations performed a few days later (12 dpf) result in partial regeneration, 2.33±0.58 digits; n=3).

Moreover, amputations performed at 14 dpf result in only a single spike (n=2; data not shown). This fortuitous plasticity allows investigation of the differences between regenerative and non-regenerative limbs (Yokoyama et al., 2011). Thus, L. laevis can serve as an accelerated model of all aspects of tadpole limb regeneration.

146

Discussion

Here, we describe the normal embryogenesis and experimental amenability of the

Budgett’s Frog (Lepidobatrachus laevis). Although we previously showed that L. laevis embryos are receptive to RNA in situ hybridization and pharmacological manipulation of signaling pathways (Bloom et al., 2013), their utility for classical experimental manipulation was untested. In this study, we assayed the responsiveness of the L. laevis embryo to microinjection, explant culture, immunohistochemical staining and regeneration, as proof of principle of its potential to serve as a complementary amphibian system. Moreover, the unusual combination of large size and rapid embryogenesis in this species make it a powerful developmental model in its own right, particularly when an increase in the resolution or speed of morphogenesis is desired.

Size matters: Benefits of larger amphibian embryos

Most amphibian eggs undergo holoblastic cleavage; however, eggs greater than ~2 mm in diameter can exhibit altered cleavage patterns, such as partially meroblastic divisions

(Collazo and Keller, 2010; del Pino and Loor-Vela, 1990; Elinson and Beckham, 2002).

Likewise, while most small eggs completely internalize the vegetal yolk mass during gastrulation/epiboly, in some larger (3.5mm) eggs (e.g., E. coqui), the yolk mass cannot be completely subsumed by epiboly and must be secondarily covered after gastrulation by the ventral-ward expansion of tissue from the body wall (Elinson and Beckham, 2002). Our observations indicate that, despite their relatively large size, L. laevis embryos are still able to undergo the ancestral mode of holoblastic cleavage and completely internalize the yolk mass

147 by epiboly. Thus, at ~2.6 mm in diameter, the L. laevis embryo appears to have achieved maximal enlargement while still retaining an early developmental program that is comparable to ancestral anurans.

From a pragmatic perspective, the increased scale of Lepidobatrachus embryos enables facile injection of single blastomeres up to at least the 128-cell stage and poses the prospect of high resolution fate mapping or extremely localized gain-of-function strategies.

The morphology of developing structures, such as craniofacial features, is inherently magnified, allowing easier visualization and tracking of the fine details of morphogenesis.

Structures such as the pharyngeal arches, stomodeum, gill buds, and embryonic mouth are larger and more prominent than in X. laevis, which could enhance studies of craniofacial development. Indeed, under a simple dissecting microscope, individual cells in the face of L. laevis are apparent; this resolution could facilitate cell fate and migration studies. In addition, tissues of interest may be more accurately isolated in the larger embryo, opening the possibility of fine-scale explant culture or transplantation experiments. Finally, the substantial (~50%) enlargement of the individual cells themselves affords views of the cellular architecture of early organs that, to our knowledge, are not otherwise attainable without advanced microscopy or special imaging modalities. Fortuitously, many antibodies known to detect Xenopus proteins also tend to be efficacious in L. laevis. Combined with the ability to express exogenous fluorescently-tagged proteins, the amplified scale of L. laevis cells and tissues has the potential to yield spectacular resolution of the cell biology underlying morphogenesis.

148

Perhaps an application in which the distinctive size of L. laevis will be particularly profitable is as a source of embryonic tissues for transcriptomic and/or proteomic profiling of highly discrete developmental events. Xenopus embryos have been shown to offer distinct advantages for proteomic analyses because of the relatively large amounts of material available (Amin et al., 2014; Rao et al., 2014; Sun et al., 2014; Wuhr et al., 2014). It is reasonable to assume that the even larger size of L. laevis could enable isolation of sufficient protein from sub-regions of organ anlage or key zones of morphogenetic activity. Although the L. laevis genome still awaits sequencing, genome-independent proteomic strategies have recently been shown to be feasible for non-model species (Wuhr et al., 2014). Since L. laevis is a diploid organism (Faivovich et al., 2014), sequencing and assembly of its genomic sequence should be less cumbersome than for the pseudotetraploid X. laevis. Moreover, a draft transcriptome assembly from embryonic organs contains greater than 26,000 predicted

L. laevis proteins [including over 10,000 full-length ORFs with significant similarity to human proteins (BLAST e-value of less than 10-5; N.M.A., N.N.-Y.-unpublished data)]; thus, what we learn in this large-egged cannibalistic frog has great potential to provide insight into bio-medically relevant biology.

The need for speed: rapid development, rapid results

Despite the sizeable mass of the L. laevis embryo compared to X. laevis, its cleavage and development to tadpole proceeds at a comparable rate at the same temperature. This is highly unexpected because, in most amphibians, larger egg size is accompanied by substantially slower development. For example, although frog embryos comparable in size to

149

L. laevis (e.g., G. riobambae at ~3 mm, or the Dendrobatids at 2-3.5 mm) have been reported to require 14 days or longer to complete gastrulation (del Pino et al., 2007), this feat may be achieved in as few as ~9 hours in L. laevis raised at 28°C. In addition, the ability to dramatically accelerate L. laevis development by culturing at temperatures as high as 37°C will facilitate the study of later stage events that take days or weeks to complete in other vertebrate models. For example, digestive tract morphogenesis takes ~3.5-4 dpf in X. laevis

(raised at 23°C), ~3 days in zebrafish, ~10 days in chicken, and >2 weeks in the mouse, but can be achieved in less than 24 hpf in L. laevis raised at 37°C. Likewise, the ability to culture

L. laevis at lower temperatures to decelerate development will allow one to maximize the size advantage of the species for dissections or lineage tracing at precise developmental time points.

The steep developmental trajectory of L. laevis extends beyond embryogenesis.

Currently, a major limitation of limb regeneration experiments is the extended time required to achieve full regeneration (Azevedo et al., 2011). However, we have shown that, in L. laevis, amputation may be initiated as early as 7-8 days after fertilization, with limb regeneration evident a little over a week (~10 days) post-amputation (at 28°C). Thus, a complete regeneration experiment, from fertilization to regrowth (~45 days in Xenopus) requires only ~2.5 weeks in L. laevis—and is likely to be further accelerated by higher rearing temperatures. The dramatically compressed timetable for organogenesis and regeneration at higher temperatures not only enables more experiments to be completed in less time, but may also facilitate high resolution imaging of events that typically occur over weeks or months in other species, a prohibitively long duration for standard time-lapse.

150

Lastly, we note that, under ideal conditions of nutrition and temperature, L. laevis metamorphosis is complete in only a few weeks. Froglets exhibit adult size and evidence of sexual behavior in as few as 4 months, and yield F1 offspring in as few as 8 months (NNY,

NA, unpublished observations); thus, expedient multi-generational studies involving mutant and/or transgenic lines may be a realistic possibility for this species.

Future Prospects: A model of biological scaling

Because of their impressive size and expeditious development, L. laevis embryos constitute a novel vertebrate system in which to investigate the poorly understood mechanisms of biological scaling, specifically, the impact of increased cell size and/or accelerated developmental rate on basic cellular processes and embryonic patterns. For example, how are the larger blastomeres in L. laevis able to properly distribute chromosomes, cytoplasmic determinants and organelles prior to cytokinesis in a time frame comparable to a much smaller embryo? How are intra- and extracellular signaling events regulated in order to maintain pattern invariance in the context of enlarged cells, expanded morphogenetic fields, and a hastened rate of organogenesis? Are the biomechanical forces and material properties that drive morphogenetic cell movements modified in larger embryos? How do different size embryos modulate cellular and molecular events to control the pace of cell division during morphogenesis? The answer to this latter question has implications for not only evolution, but also for the advancement of pathological events such as tumor progression. By serving as an extreme point of comparison to other anurans, the larger and faster Budgett’s frog embryo may help us to understand, and ultimately predict, the

151 phenotypic consequences of varying the fundamental physical parameters of metazoan development.

Acknowledgements

We would like to thank Dr. Michael Dush for suggestions on the manuscript, Dr. Michael

Levy for his guidance in the husbandry of the animals used in this work, and Dr. Michael

Levin for advice on regeneration experiments. This work was supported by NIH (RO1

DK085300, R21 OD017963) awards to N.N-Y.

152

Figure A- 1 ) The schedule of Lepidobatrachus laevis embryonic development (versus Xenopus laevis) Developmental trajectories from fertilization (Gosner stage 1; GS1) to early tadpole (GS22) were plotted for L. laevis (blue diamonds, 28°C) and X. laevis (black circles, 23°C). Data points for L. laevis were calculated from the average time to achieve the developmental milestones characteristic of each stage in at least three separate clutches of embryos from at least two different adult breeding pairs. Developmental timing of Xenopus laevis was plotted from published reports (Nieuwkoop and Faber, 1994). For clarity, stages are clustered into cleavage/blastula (pink), gastrula (green), neurula (yellow) and organogenesis (blue) phases. The dashed line at 30 hpf indicates a break in the x-axis, to accommodate the substantial difference (~50 hours) in the developmental schedule of the two species (at their respective optimal temperatures—note that both organisms develop at the same rate when raised at mutually permissive temperatures; see Figure 5). Gosner (GS) and Nieuwkoop and Faber (NF) stages are indicated.

153

Figure A- 2) Early cleavage and gastrulation patterns in L. laevis are similar to Xenopus Representative images of L. laevis fertilization (A; GS1); 2-cell (B; GS3); 4-cell (C; GS4); 8- cell (D; GS5); 16-cell (E; GS6); mid-blastula (F; GS8); late blastula (G; GS9); and early gastrula (H; GS10). “D” and “V” indicate the prospective dorsal and ventral sides of the embryo, respectively. (I) A side-by-side comparison of a L. laevis 2-cell stage embryo with a X. laevis gastrula demonstrates the substantial size difference between the two species. J) The dorsal lip (DL) is obvious as a pigmented depression (arrow) in the early gastrula (GS10). K) The dorsal lip (arrow) expands circumferentially (arrowheads). L) The yolk plug is still slightly protuberant in the midgastrula (GS 11). A-I are animal views; J and K are latero- vegetal views; L is a vegetal view. Scale bar = 1 mm.

154

Figure A- 3) Neurulation and organogenesis in L. laevis The neural plate (A; GS12.5), neural folds (yellow arrowheads, B; GS14) and migrating streams of neural crest (arrows, B; GS14) are highly prominent in dorsal views of early neurulae. C) Neural tube closure is complete and individual somites (S) are evident by GS16. D-F) The cement gland (CG) and pronephros (P) are visible at the earliest tail bud stages (GS17-19). G-H) Later tail bud stages (GS20-22) show rapid tail elongation, the development of gills and craniofacial features (see also Figure 4), and early gut morphogenesis. NP= neural plate; H=heart; L=liver; G=gills; FG=foregut; MG=midgut; HG=hindgut. Scale bar = 1 mm.

155

Figure A- 4) Craniofacial morphogenesis in L. laevis The embryonic head is shown in lateral (A, D, G, J; anterior to the left) and frontal views (B, E, H, K). Arrows indicate the location of the stomodeum (stom), or mouth. Higher magnification views (C, F, I, L) detail the gradual maturation of the mouth, including the perforating buccopharyngeal membrane (bm). The face and mouth widen dramatically as development proceeds, and the cement gland (cg), which initially forms a horseshoe shape (B), becomes bifurcated as the face widens (K). Gosner Stages (GS) are as indicated. Scale bars in A-B, D-E, G-H and J-K = 500μm; scale bars in C, F, I, L = 250μm. Other abbreviations: gb=gill buds; n=nasal pits; op=optic cup; ot=otic vesicle, pa=pharyngeal arches; so= somites.

156

157

Figure A- 5) Thermal tolerance of L. laevis and X. laevis embryos L. laevis (solid lines, n=10) and X. laevis (dotted lines, n=20) were assayed for growth from 2-cell stage (GS3) to tadpole (GS22) at 37°C (purple), 32°C (green), 28°C (black), 23°C (orange), 16°C (blue). Asterisks (*) indicate time points at which at least 50% of embryos fail to gastrulate, indicating decreased viability. Gosner (GS) and Nieuwkoop and Faber (NF) stages are indicated.

158

Figure A- 6) L. laevis embryos are amenable to microinjection of exogenous reagents for fate-mapping and expression of synthetic mRNA Embryos were injected at the 2-cell stage (A-C), 8-cell stage (D-F) or 128-cell stage (G-I) with red fluorescent dextran and/or capped eGFP mRNA. By 48 hours post fertilization (hpf), injection into a single blastomere at the 2-cell stage results in predominantly unilateral labeling (B) and expression of eGFP (C, green), while injection into a single blastomere at the 8-cell stage results in fluorescence limited to the developing gut (E). Injection of dextran into a single blastomere at the 128-cell stage labels a limited region of only 4 cells by the mid-blastula stage (H). Merge of bright field and fluorescent images in (F, I). Scale bars = 1 mm.

159

Figure A- 7) L. laevis explants are amenable to animal cap explant culture Untreated animal caps from X. laevis (A) or L. laevis (B) heal into balls of ciliated epidermis. In contrast, treatment of freshly microdissected caps with activin results in significant elongation of dissected tissue in both X. laevis (C) and L. laevis (D). The smaller right-most cap in B and D was trimmed to a reduced size. Scale bars = 1 mm.

160

Figure A- 8) Heart morphogenesis occurs at a larger scale in L. laevis The scale of L. laevis cardiogenesis is illustrated by comparing a dissected L. laevis heart tube (GS18) with a whole X. laevis neurula embryo (A), and by comparing mature hearts isolated at an equivalent stage from each species (B; GS22). Immunofluorescence analysis of tissue architecture was performed on transverse sections through the heart of GS21 (C-F) and GS20 (G-J) embryos. C-F) Localization of beta-catenin (cat; green) in X. laevis and actin (actin; green) in L. laevis reveals the cellular architecture of the heart. G-H) Localization of the microtubule marker alpha-tubulin (tub; green) and beta-catenin (cat; red) in the L. laevis heart tube. I-J) Localization of the centrosome marker gamma-tubulin (tub; green), and filamentous actin (F-actin; red), in a neighboring section through the L. laevis heart tube. Nuclei are counterstained with TO-PRO-3. White boxes in C, D, G, I indicate regions of the heart magnified in E, F, H, J, respectively. c, conus; v, ventricle; endo, endocardial layer; myo, myocardium. Scale bars = 100 m.

161

162

Figure A- 9) Limb regeneration occurs at a rapid pace in L. laevis Amputation was conducted on 8-day tadpoles at the indicated level (red dotted line; A). A blastema was identified within 24 hours of amputation (arrow; B). Full regeneration was observed in the operated limb by 10 days post amputation (C).

163

References

Amaya,E., Musci,T.J., and Kirschner,M.W. (1991). Expression of a dominant negative mutant of the FGF receptor disrupts mesoderm formation in Xenopus embryos. Cell 66, 257- 270.

Amin,N.M., Greco,T.M., Kuchenbrod,L.M., Rigney,M.M., Chung,M.I., Wallingford,J.B., Cristea,I.M., and Conlon,F.L. (2014). Proteomic profiling of cardiac tissue by isolation of nuclei tagged in specific cell types (INTACT). Development 141, 962- 73.

AmphibiaWeb . Information on amphibian biology and conservation. Berkeley, California: AmphibiaWeb. 12-15-2014. 12-15-2014. Ref Type: Electronic Citation

Azevedo,A.S., Grotek,B., Jacinto,A., Weidinger,G., and Saude,L. (2011). The regenerative capacity of the zebrafish caudal fin is not affected by repeated amputations. PLoS One. 6, e22820.

Beck,C.W. (2012). Studying regeneration in Xenopus. Methods Mol. Biol. 917, 525-39.

Blitz,I.L., Biesinger,J., Xie,X., and Cho,K.W. (2013). Biallelic genome modification in F(0) Xenopus tropicalis embryos using the CRISPR/Cas system. Genesis 51, 827-834.

Bloom,S., Ledon-Rettig,C., Infante,C., Everly,A., Hanken,J., and Nascone-Yoder,N. (2013). Developmental origins of a novel gut morphology in frogs. Evolution & Development 15, 213-223.

Brieher,W.M. and Gumbiner,B.M. (1994). Regulation of C-cadherin function during activin induced morphogenesis of Xenopus animal caps. J Cell Biol. 126, 519-527.

Brockes,J.P. and Kumar,A. (2002). Plasticity and reprogramming of differentiated cells in amphibian regeneration. Nat. Rev. Mol. Cell Biol. 3, 566-574.

Budgett,J.S. (1899). Notes on the batrachians of the Paraguayan Chaco, with observations upon their breeding habits and development, especially with regard to Phyllomedusa hypochondrialis, Cope. Also a description of a new genus. QJ Microsc. Sci. 305-333.

Calboli,F.C., Fisher,M.C., Garner,T.W., and Jehle,R. (2011). The need for jumpstarting amphibian genome projects. Trends Ecol. Evol. 26, 378-379.

Callery,E.M. (2006). There's more than one frog in the pond: a survey of the Amphibia and their contributions to developmental biology. Semin. Cell Dev. Biol. 17, 80-92.

Carroll,E.J. (1996). Thermal tolerance and heat shock protein synthesis during development in the anuran Lepidobatrachus laevis. Dev Growth Differ. 38, 9-14.

164

Carroll,E.J., Seneviratne,A.M., and Ruibal,R. (1991). Gastric Pepsin in an Anuran Larva. Dev Growth Differ 33, 499-507.

Collazo,A. and Keller,R. (2010). Early development of Ensatina eschscholtzii: an amphibian with a large, yolky egg. Evodevo. 1, 6-1. del Pino,E.M., Avila,M.E., Perez,O.D., Benitez,M.S., Alarcon,I., Noboa,V., and Moya,I.M. (2004). Development of the dendrobatid frog Colostethus machalilla. Int J Dev Biol. 48, 663-670. del Pino,E.M. and Loor-Vela,S. (1990). The pattern of early cleavage of the marsupial frog Gastrotheca riobambae. Development. 110, 781-789. del Pino,E.M., Venegas-Ferrin,M., Romero-Carvajal,A., Montenegro-Larrea,P., Saenz- Ponce,N., Moya,I.M., Alarcon,I., Sudou,N., Yamamoto,S., and Taira,M. (2007). A comparative analysis of frog early development. Proc Natl Acad Sci U S A. 104, 11882- 11888.

Dent,J.N. (1962). Limb regeneration in larvae and metamorphosing individuals of the South African clawed toad. J Morphol 110, 61-77.

Dush,M.K., McIver,A.L., Parr,M.A., Young,D.D., Fisher,J., Newman,D.R., Sannes,P.L., Hauck,M.L., Deiters,A., and Nascone-Yoder,N. (2011). Heterotaxin: a TGF-beta signaling inhibitor identified in a multi-phenotype profiling screen in Xenopus embryos. Chem Biol 18, 252-263.

Elinson,R.P. and Beckham,Y. (2002). Development in frogs with large eggs and the origin of amniotes. Zoology (Jena). 105, 105-117.

Fabrezi,M. (2006). Morphological evolution of Ceratophryinae (Anura, Neobatrachia). J Zool Sys Evol Res. 44, 153-166.

Faivovich,J., Nicoli,L., Blotto,B.L., Pereyra,M.O., Baldo,D., Barrionuevo,J.S., Fabrezi,M., Wild,E.R., and Haddad,C.F.B. (2014). Big, Bad, and Beautiful: Phylogenetic Relationships of the Horned Frogs (Anura: Ceratophryidae). South Am J Herpetol. 9, 207- 227.

Fei,J.F., Schuez,M., Tazaki,A., Taniguchi,Y., Roensch,K., and Tanaka,E.M. (2014). CRISPR-mediated genomic deletion of Sox2 in the axolotl shows a requirement in spinal cord neural stem cell amplification during tail regeneration. Stem Cell Reports. 3, 444-459.

Flowers,G.P., Timberlake,A.T., McLean,K.C., Monaghan,J.R., and Crews,C.M. (2014). Highly efficient targeted mutagenesis in axolotl using Cas9 RNA-guided nuclease. Development. 141, 2165-2171.

165

Goode,R.P. (1967). The regeneration of limbs in adult anurans. J Embryol Exp Morphol. 18, 259-267.

Gosner,K.L. (1960). A simplified table for staging anuran embryos and larvae. Herpetologica 1960.

Guo,X., Zhang,T., Hu,Z., Zhang,Y., Shi,Z., Wang,Q., Cui,Y., Wang,F., Zhao,H., and Chen,Y. (2014). Efficient RNA/Cas9-mediated genome editing in Xenopus tropicalis. Development 141, 707-714.

Hall,B.K. (1992). Evolutionary Developmental Biology. Chapman & Hall.

Hamlet,M.R., Yergeau,D.A., Kuliyev,E., Takeda,M., Taira,M., Kawakami,K., and Mead,P.E. (2006). Tol2 transposon-mediated transgenesis in Xenopus tropicalis. Genesis 44, 438-445.

Hanken,J. (1993). Model Systems Versus Outgroups - Alternative Approaches to the Study of Head Development and Evolution. Amer Zool. 33, 448-456.

Hellsten,U., Harland,R.M., Gilchrist,M.J., Hendrix,D., Jurka,J., Kapitonov,V., Ovcharenko,I., Putnam,N.H., Shu,S., Taher,L. et al. (2010). The genome of the Western clawed frog Xenopus tropicalis. Science 328, 633-636.

Hopwood,N.D. and Gurdon,J.B. (1990). Activation of muscle genes without myogenesis by ectopic expression of MyoD in frog embryo cells. Nature 347, 197-200.

Khattak,S., Murawala,P., Andreas,H., Kappert,V., Schuez,M., Sandoval-Guzman,T., Crawford,K., and Tanaka,E.M. (2014). Optimized axolotl (Ambystoma mexicanum) husbandry, breeding, metamorphosis, transgenesis and tamoxifen-mediated recombination. Nat Protoc. 9, 529-540.

Khokha,M.K., Chung,C., Bustamante,E.L., Gaw,L.W., Trott,K.A., Yeh,J., Lim,N., Lin,J.C., Taverner,N., Amaya,E. et al. (2002). Techniques and probes for the study of Xenopus tropicalis development. Dev Dyn. 225, 499-510.

Kroll,K.L. and Amaya,E. (1996). Transgenic Xenopus embryos from sperm nuclear transplantations reveal FGF signaling requirements during gastrulation. Development 122, 3173-3183.

Ledon-Rettig,C.C. and Pfennig,D.W. (2011). Emerging model systems in eco-evo-devo: the environmentally responsive spadefoot toad. Evol Dev. 13, 391-400.

Lei,Y., Guo,X., Liu,Y., Cao,Y., Deng,Y., Chen,X., Cheng,C.H., Dawid,I.B., Chen,Y., and Zhao,H. (2012). Efficient targeted gene disruption in Xenopus embryos using

166 engineered transcription activator-like effector nucleases (TALENs). Proc Natl Acad Sci U S A. 109, 17484-17489.

Liu,Y., Luo,D., Lei,Y., Hu,W., Zhao,H., and Cheng,C.H. (2014). A highly effective TALEN-mediated approach for targeted gene disruption in Xenopus tropicalis and zebrafish. Methods 14, 00041-00043.

Logan,M. and Mohun,T. (1993). Induction of cardiac muscle differentiation in isolated animal pole explants of Xenopus laevis embryos. Development 118, 865-875.

Moody,S.A. (1987a). Fates of the blastomeres of the 16-cell stage Xenopus embryo. Dev Biol. 119, 560-578.

Moody,S.A. (1987b). Fates of the blastomeres of the 32-cell-stage Xenopus embryo. Dev Biol. 122, 300-319.

Moya,I.M., Alarcon,I., and del Pino,E.M. (2007). Gastrulation of Gastrotheca riobambae in comparison with other frogs. Dev Biol. 304, 467-478.

Nakajima,K. and Yaoita,Y. (2013). Comparison of TALEN scaffolds in Xenopus tropicalis. Biol Open. 2, 1364-1370.

Nakayama,T., Fish,M.B., Fisher,M., Oomen-Hajagos,J., Thomsen,G.H., and Grainger,R.M. (2013). Simple and efficient CRISPR/Cas9-mediated targeted mutagenesis in Xenopus tropicalis. Genesis 51, 835-843.

Nieuwkoop,P.D. and Faber,J. (1994). Normal Table of Xenopus laevis (Daudin): A Systematical & Chronological Survey of the Development from the Fertilized Egg till the End of Metamorphosis. Amsterdam.

Ogino,H., McConnell,W.B., and Grainger,R.M. (2006). High-throughput transgenesis in Xenopus using I-SceI meganuclease. Nat Protoc. 1, 1703-1710.

Parker,H.W. (1931). Reports of an Expedition to Brazil and Paraguay in 1926-7, supported by the Trustees of the Percy Sladen Memorial Fund and the Executive Committee of the Carnegie Trust for Scotland. Amphibia and Reptilia. J Proc Linn Soc. Zoology. 285-289.

Perez,O.D., Lai,N.B., Buckley,D., del Pino,E.M., and Wake,M.H. (2009). The morphology of prehatching embryos of Caecilia orientalis (Amphibia: Gymnophiona: Caeciliidae). J Morphol. 270, 1492-1502.

Ramsdell,A.F., Bernanke,J.M., and Trusk,T.C. (2006). Left-right lineage analysis of the embryonic Xenopus heart reveals a novel framework linking congenital cardiac defects and laterality disease. Development. 133, 1399-1410.

167

Rao,N., Song,F., Jhamb,D., Wang,M., Milner,D.J., Price,N.M., Belecky-Adams,T.L., Palakal,M.J., Cameron,J.A., Li,B.,Chen,X., and Stocum,D.L. (2014). Proteomic analysis of fibroblastema formation in regenerating hind limbs of Xenopus laevis froglets and comparison to axolotl. BMC Dev Biol. 14-32.

Romero-Carvajal,A., Saenz-Ponce,N., Venegas-Ferrin,M., Almeida-Reinoso,D., Lee,C., Bond,J., Ryan,M.J., Wallingford,J.B., and del Pino,E.M. (2009). Embryogenesis and laboratory maintenance of the foam-nesting tungara frogs, genus Engystomops (=Physalaemus). Dev Dyn. 238, 1444-1454.

Ruibal,R. and Thomas,E. (1988). The obligate carnivorous larvae of the frog Lepidobatrachus laevis (Leptodactylidae). Copeia 3, 591-604.

Saint-Jeannet,J.P., Karavanov,A.A., and Dawid,I.B. (1994). Expression of mesoderm markers in Xenopus laevis Keller explants. Int J Dev Biol. 38, 605-611.

Sakane,Y., Sakuma,T., Kashiwagi,K., Kashiwagi,A., Yamamoto,T., and Suzuki,K.T. (2014). Targeted mutagenesis of multiple and paralogous genes in Xenopus laevis using two pairs of transcription activator-like effector nucleases. Dev Growth Differ. 56, 108-114.

Sater,A.K. and Jacobson,A.G. (1989). The specification of heart mesoderm occurs during gastrulation in Xenopus laevis. Development. 105, 821-830.

Sater,A.K. and Jacobson,A.G. (1990). The restriction of the heart morphogenetic field in Xenopus laevis. Dev Biol. 140, 328-336.

Scadding,S.R. (1977). Phylogenic Distribution of Limb Regeneration Capacity in Adult Amphibia. J Exp Zool. 202, 57-67.

Sive,H.L., Grainger,R.M., and Harland,R.M. (1998). Early Development of Xenopus laevis. Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press.

Smith,J.C., Symes,K., Hynes,R.O., and DeSimone,D. (1990). Mesoderm induction and the control of gastrulation in Xenopus laevis: the roles of fibronectin and integrins. Development. 108, 229-238.

Smith,W.C., Knecht,A.K., Wu,M., and Harland,R.M. (1993). Secreted noggin protein mimics the Spemann organizer in dorsalizing Xenopus mesoderm. Nature. 361, 547-549.

Sobkow,L., Epperlein,H.H., Herklotz,S., Straube,W.L., and Tanaka,E.M. (2006). A germline GFP transgenic axolotl and its use to track cell fate: dual origin of the fin mesenchyme during development and the fate of blood cells during regeneration. Dev Biol. 290, 386-397.

168

Spemann,H. and Mangold,H. (1924). Induction of embryonic primordia by implantation of organizers from a different species. Roux's Arch Entw Mech. 100, 599-638.

Stuart,S.N., Chanson,J.S., Cox,N.A., Young,B.E., Rodrigues,A.S., Fischman,D.L., and Waller,R.W. (2004). Status and trends of amphibian declines and extinctions worldwide. Science. 306, 1783-1786.

Sun,L., Bertke,M.M., Champion,M.M., Zhu,G., Huber,P.W., and Dovichi,N.J. (2014). Quantitative proteomics of Xenopus laevis embryos: expression kinetics of nearly 4000 proteins during early development. Sci Rep. 4, 4365.

Suzuki,K.T., Isoyama,Y., Kashiwagi,K., Sakuma,T., Ochiai,H., Sakamoto,N., Furuno,N., Kashiwagi,A., and Yamamoto,T. (2013). High efficiency TALENs enable F0 functional analysis by targeted gene disruption in Xenopus laevis embryos. Biol Open. 2, 448-452.

Tandon,P., Showell,C., Christine,K., and Conlon,F.L. (2012). Morpholino injection in Xenopus. Methods Mol Biol. 843, 29-46.

Tank,P.W., Carlson,B.M., and Connelly,T.G. (1976). A staging system for forelimb regeneration in the axolotl, Ambystoma mexicanum. J Morphol. 150, 117-128.

Whited,J.L., Lehoczky,J.A., and Tabin,C.J. (2012). Inducible genetic system for the axolotl. Proc Natl Acad Sci U S A. 109, 13662-13667.

Wilson,P.A., Oster,G., and Keller,R. (1989). Cell rearrangement and segmentation in Xenopus: direct observation of cultured explants. Development. 105, 155-166.

Woodland,H.R. and Jones,E.A. (1987). The development of an assay to detect mRNAs that affect early development. Development. 101, 925-930.

Wuhr,M., Freeman,R.M., Jr., Presler,M., Horb,M.E., Peshkin,L., Gygi,S.P., and Kirschner,M.W. (2014). Deep proteomics of the Xenopus laevis egg using an mRNA- derived reference database. Curr Biol. 24, 1467-1475.

Yergeau,D.A., Johnson Hamlet,M.R., Kuliyev,E., Zhu,H., Doherty,J.R., Archer,T.D., Subhawong,A.P., Valentine,M.B., Kelley,C.M., and Mead,P.E. (2009). Transgenesis in Xenopus using the Sleeping Beauty transposon system. Dev Dyn. 238, 1727-1743.

Yokoyama,H., Maruoka,T., Ochi,H., Aruga,A., Ohgo,S., Ogino,H., and Tamura,K. (2011). Different requirement for Wnt/beta-catenin signaling in limb regeneration of larval and adult Xenopus. PLoS One. 6, e21721.

Young,H.E., Bailey,C.F., and Dalley,B.K. (1983). Gross morphological analysis of limb regeneration in postmetamorphic adult Ambystoma. Anat Rec. 206, 295-306.

169

Young,J.J., Cherone,J.M., Doyon,Y., Ankoudinova,I., Faraji,F.M., Lee,A.H., Ngo,C., Guschin,D.Y., Paschon,D.E., Miller,J.C. et al. (2011). Efficient targeted gene disruption in the soma and germ line of the frog Xenopus tropicalis using engineered zinc-finger nucleases. Proc Natl Acad Sci U S A. 108, 7052-7057.

Ziermann,J.M., Infante,C., Hanken,J., and Olsson,L. (2013). Morphology of the cranial skeleton and musculature in the obligate carnivorous tadpole of Lepidobatrachus laevis (Anura: Ceratophryidae). Acta Zool. 94, 101-112.

170

Appendix B: Frogs as integrative models for understanding digestive organ development and evolution

Mandy Womble1, Melissa Pickett1 and Nanette Nascone-Yoder*

Department of Molecular Biomedical Sciences North Carolina State University Raleigh, NC 27676

* author for correspondence [email protected], 919-513-8284

1 these authors contributed equally to this work

Published in Semin. Cell Dev. Biol. 2016 Mar; 51:92-105. doi: 10.1016/j.semcdb.2016.02.001.

171

Abstract

The digestive system comprises numerous cells, tissues and organs that are essential for the proper assimilation of nutrients and energy. Many aspects of digestive organ function are highly conserved among vertebrates, yet the final anatomical configuration of the gut varies widely between species, especially those with different diets. Improved understanding of the complex molecular and cellular events that orchestrate digestive organ development is pertinent to many areas of biology and medicine, including the regeneration or replacement of diseased organs, the etiology of digestive organ birth defects, and the evolution of specialized features of digestive anatomy. In this review, we highlight specific examples of how investigations using Xenopus laevis frog embryos have revealed insight into the molecular and cellular dynamics of digestive organ patterning and morphogenesis that would have been difficult to obtain in other animal models. Additionally, we discuss recent studies of gut development in non-model frog species with unique feeding strategies, such as

Lepidobatrachus laevis and Eleutherodactylous coqui, which are beginning to provide glimpses of the evolutionary mechanisms that may generate morphological variation in the digestive tract. The unparalleled experimental versatility of frog embryos make them excellent, integrative models for studying digestive organ development across multiple disciplines.

172

1. Introduction

The anatomical and physiological complexity of the vertebrate digestive system develops from a simple primitive gut tube (PGT). This PGT undergoes intricate patterning and differentiation events to enable the epithelial lining of the tube to assume the absorptive and secretory functions required of a gastrointestinal (GI) tract, while discrete segments bud off of the original structure to form accessory organs, including the pancreas and liver.

Concomitantly, the tube lengthens and rotates, as it transforms from a short, occluded cylinder to a long, hollow conduit arranged in a three dimensional configuration of loops and coils.

Elucidating the mechanisms of digestive organ development has broad implications for many areas of biology and medicine. Some of the most common human birth defects affect the digestive tract, yet the genetic and/or environmental factors that contribute to the etiology of these malformations remain to be discovered. In addition, diseases of the digestive system affect millions worldwide, generating substantial demand for therapeutic interventions; full knowledge of the developmental events that pattern and shape the PGT is likely to be vital for successful regeneration or engineering of human digestive tissues.

Finally, although many features of digestive anatomy are highly conserved among vertebrates, the length, compartmentalization and topological orientation of the GI tract can vary tremendously among and between species, especially those with different diets, yet the evolutionary origins of this ecologically-relevant variation are largely unknown.

173

1.1. The advantages of the frog embryo.

Amphibians have long been used as model organisms for studying embryonic development, and have played instrumental roles in unraveling the intricate events that guide germ layer formation, gastrulation and neurulation [1-3]. Beyond early development, frog embryos also boast several advantages for the study of organ specification and morphogenesis [4]. Unlike amniotic embryos that are confined to a uterus or shell during development, frog embryos are externally fertilized and can be easily cultured in vitro, making them amenable to a wide variety of experimental manipulations. For example, the rate of development of frog embryos can be accelerated or slowed by adjusting temperature, facilitating convenient analyses of any stage of organogenesis [5]. Moreover, precise fate maps have been generated for the early blastomeres (32-cell stage), allowing loss- and/or gain-of-function (LOF/GOF) reagents and lineage tracers to be targeted to specific organs by standard microinjection technology, enabling gene function to be queried in a tissue-specific manner [6, 7]. Furthermore, because frog embryos are relatively large and harbor an innate, intracellular yolk supply, tissue explants can be dissected, recombined and transplanted, or cultured in simple saline, at almost any stage of development, facilitating expedient, inexpensive specification and trans-differentiation studies [8-14]. Finally, the frog embryo’s accessibility to chemical agonists/antagonists allows the role of specific signaling pathways to be interrogated during critical windows of organogenesis (i.e., subsequent to earlier developmental events that may also depend on such pathways). In fact, thanks to large clutch sizes, frog embryos provide a powerful platform for high-throughput “chemical genetic” or toxin screening using organ morphology as a phenotypic readout [15-18]. This experimental

174 amenability makes the frog embryo an ideal model in which to interrogate the mechanisms of organ development.

1.2. More than one frog in the water.

Amphibian models (mainly urodeles) have been employed in developmental biology research for over a century, but the convenience of in vitro fertilization methods made

Xenopus species the most popular frogs in the laboratory [19]. Nonetheless, many non-model frog species are equally amenable to experimentation as Xenopus. Comparative “evo-devo” studies utilizing frogs with different reproductive strategies and/or developmental rates [20] are beginning to provide fascinating insight into the molecular and cellular mechanisms that shape different embryos, while species that fill unique ecological niches or possess intriguing specializations are shedding light on the developmental origins of novel morphologies [21].

In this review, we provide a broad perspective on the ways in which Xenopus and emerging frog models have yielded new insight into digestive organ patterning, morphogenesis, and evolution.

2. What can the frog tell us about foregut organ specification?

The developing digestive tract may be divided into foregut (esophagus, stomach, duodenum, liver, pancreas, gall bladder) and midgut/hindgut (intestine) domains. The foregut-derived organs play critical roles in processes such as digestion, glucose homeostasis, and detoxification. Therefore, congenital defects or disease in these organs (e.g., diabetes,

175 pancreatitis, fatty liver disease, biliary atresia, gall stones and gastric/pancreatic cancer) are the cause of substantial morbidity and mortality worldwide [22-25]. To ameliorate such afflictions, translational researchers seek to develop regenerative therapies and engineer replacement tissues in vitro. Progress in these areas has been profoundly influenced by models of the normal process of foregut organ specification and morphogenesis in the embryo [26, 27].

In all vertebrates, the PGT is comprised of an inner endoderm layer, which differentiates into the epithelial lining of the GI tract, surrounded by an outer layer of mesoderm, which will give rise to the visceral muscle and connective tissue. Early in gut development, reciprocal signaling between the endoderm and mesoderm layers gradually distinguishes anterior foregut and posterior hindgut domains [28]. In addition to digestive tissues, numerous structures with diverse physiological functions are derived from the anterior region of the PGT (including organs of the respiratory and endocrine systems), all of which must undergo morphogenesis in close proximity. The expression of foregut organ- specific genes must therefore be tightly coordinated in time and space to allow individual organs to differentiate appropriately. Moreover, many of the signaling factors involved in foregut organogenesis are re-deployed during different stages of development, and/or evoke contradictory responses depending on their concentration [27].

Studying such spatially intricate and seemingly paradoxical signaling dynamics is not trivial in mouse models due to the functional redundancy of the factors involved, and the challenges of achieving tissue-specific or conditional perturbation of gene function to overcome early lethality and pleiotropy. In contrast, the experimental amenability of the

176

Xenopus embryo—which enables the pattern, timing and dosage of gene expression to be manipulated in a tissue-specific manner—has provided key insights into the spatiotemporal signaling dynamics that specify region and organ identities in the PGT. We highlight a few salient examples in sections 2.1-2.4 below.

2.1. Complex control of Wnt signaling is required for foregut specification.

Wnt signaling pathways are highly conserved and involved in many fundamental developmental events, including body axis patterning, cell fate specification, cell proliferation, and cell migration. Multiple Wnt ligands stimulate canonical (Wnt/-catenin) and/or non-canonical (e.g., Wnt/JNK) pathways [29, 30]. Early in Xenopus development, the establishment of the dorso-anterior axis of the embryo is accompanied by high levels of nuclear -catenin in the anterior endoderm, a readout of canonical Wnt signaling. However, soon afterwards, this same tissue exhibits low levels of nuclear-catenin, suggesting that

Wnt signaling must become restricted from the prospective foregut during gut patterning

[31]. This idea was confirmed by experiments in which the Wnt/-catenin pathway was ectopically activated in the prospective foregut of the Xenopus embryo, resulting in ablation of liver- and pancreas-specific gene expression; in the converse experiment, inhibition of

-catenin signaling in the prospective hindgut region expanded liver and pancreas domains at the expense of intestinal tissue [31].

The expression of Wnt antagonists in the anterior endoderm also suggests that Wnt signaling is actively suppressed in this region. In Xenopus, foregut-specific knockdown of

177

Sfrp5, a secreted antagonist that sequesters Wnt ligands in the extracellular space to prevent their binding to Wnt receptors, reduced liver and pancreas gene expression [32]. In contrast, ectopic Sfrp5 activity, achieved via targeted injection of synthetic mRNA, expanded the foregut region, inducing massive liver and pancreatic buds[32]. Co-immunoprecipitation assays confirm that Sfrp5 binds and antagonizes Wnt 11 and 5, demonstrating a direct inhibition of the posteriorizing Wnt pathway in the anterior foregut endoderm [32].

Interestingly, Sfrp molecules have been shown to have biphasic potential, i.e., acting to inhibit Wnt ligands at high concentrations, but improving diffusion and signaling at low concentrations [33, 34]. The ability to manipulate the effective concentrations of LOF/GOF reagents targeted to specific tissues in Xenopus revealed that this biphasic functionality of

Sfrp5 is also deployed during foregut specification—high levels of the Wnt inhibitor Sfrp5 decreased levels of the liver-specific transcription factor, hhex, but moderate levels increased hhex expression [35]. These results suggest that, although Wnt/-catenin signaling must be suppressed in the prospective foregut region of the PGT, relative to the midgut/hindgut region, low levels of Wnt/-catenin signaling actually potentiate foregut organ development.

Indeed, foregut-targeted knockdown of the Wnt receptor Fzd7 elicits foregut organ hypoplasia [35]. Interestingly, in addition to canonical Wnt/-catenin signaling, non- canonical Wnt/Jnk mediated cellular morphogenetic pathways were also implicated in this process; Fzd7 knockdown caused the endoderm cells of the developing foregut to be enlarged and loosely adherent with reduced C-cadherin, -integrin, cortical -catenin and F- actin levels and disorganized microtubules [35].

178

Importantly, the frog model of Wnt-mediated foregut specification is nicely corroborated by results obtained in mammalian studies. For example, transgenic mice with foregut-specific Wnt overexpression exhibit pancreas agenesis [36] and knock out [37] or downregulation [38] of mouse Sfrps leads to hypoplastic stomach development. Thus, tissue- targeted LOF/GOF assays in frog embryos can provide detailed mechanistic insights into the complex spatiotemporal roles of Wnt signaling in foregut specification (see Figure 1) that are directly relevant to higher vertebrates but would have been difficult to ascertain in such models.

2.2. Concentration and time-dependent FGF signals segregate foregut organs.

Fibroblast Growth Factors (FGFs) are required for multiple developmental processes including mesoderm induction, limb bud development, neural patterning, myogenesis, and organ morphogenesis [39, 40]. During gastrulation, FGF signaling specifies posterior fates in the PGT but, shortly thereafter, FGF secreted from the anterior lateral plate and cardiac mesoderm is required to specify anterior foregut organs (pancreas, liver and lung) in the ventral endoderm [27]. In in vitro studies, the induction of different organs by FGF appears to be concentration-dependent, suggesting that FGF signaling must be tightly regulated during foregut specification. However, it is not known whether the segregation of organ fates in vivo is determined by proximity to, or duration of contact with, the neighboring FGF- secreting mesoderm.

The experimental amenability of Xenopus, which permits straightforward explant culture of isolated endoderm and mesoderm layers of the PGT [41], has facilitated a deeper

179 appreciation of the in vivo context-dependent regulation of FGF dosage during foregut organ specification. Consistent with higher vertebrates, gut-targeted hyper-activation of FGF signaling in Xenopus embryos results in the expansion of liver gene expression and repression of pancreas genes, while inhibition of FGF signaling causes a loss of liver gene expression and expansion of pancreatic markers [42]. Mechanistic insight was obtained from ex vivo cultures in which removal of the lateral plate mesoderm from explants of ventral endoderm at successive stages of development showed that the pancreas and liver require different time periods of interaction with the neighboring FGF-expressing mesoderm; the liver requires a more prolonged period of incubation to form correctly [42]. Chemical inhibitors of both the PI3K and MEK branches of the FGF pathway are capable of eliciting a reduction in liver gene expression, suggesting that both branches are likely required [42].

However, in the absence of mesoderm, addition of FGF to cultured foregut endoderm explants was not sufficient to induce liver, suggesting that other signaling pathways must also be involved in ventral foregut organ specification.

This example illustrates how the unique ability to isolate and culture Xenopus explants in different tissue-layer and chemical reagent combinations can enable elegant analyses of the concentration and time-dependent nature of conserved growth factor signaling in specifying vertebrate foregut organs (see Figure 1).

180

2.3. Early retinoic acid signaling is required for dorsal pancreas specification.

The pancreas is formed from three different progenitor populations, one in the dorsal region of the foregut and two (left and right) in the ventral domain. Endocrine cells are initially specified in the dorsal pancreas to make hormones such as Insulin, while the ventral pancreas produces mainly exocrine cells and digestive enzymes. As development proceeds, the dorsal and ventral buds fuse together to form one organ [4, 43-45], with endocrine and exocrine cells distributed throughout the fused structure. At somite stages, the development of the dorsal (endocrine) pancreas is regulated by signaling molecules secreted from neighboring tissues (e.g., the notochord), including TGF-β and FGF signals, which repress sonic hedgehog (shh) expression in the dorsal foregut endoderm, a prerequisite for pancreatic fate [46].

The ability to culture early embryonic tissues from Xenopus embryos provided some of the first evidence of the key role of another signaling molecule, retinoic acid (RA), in vertebrate pancreas specification. RA is a small, diffusible lipophilic molecule (synthesized from a vitamin A precursor) that acts as a morphogen to exert concentration-dependent effects on embryonic patterning. RA signals through Retinoic Acid Receptors (RARs), converting them from transcriptional repressors to activators [47]. Utilizing the ability to culture tissue explants from early Xenopus embryos, Moriya et al showed that treatment of naïve ectoderm tissue from blastulae with a combination of both Activin and RA induces the differentiation of morphological and functional pancreatic tissue [48]. In a related study, slightly later explants from early gastrulae were also induced to form pancreas after exposure

181 to RA [49]. In both cases, the RA-treated explants expressed endocrine hormones, including

Insulin, and developed pancreas-like tissue architecture.

The above studies indicate that RA influences pancreas specification very early, during or soon after gastrulation. To determine whether RA signaling is also required at this time for endogenous pancreas specification, Xenopus embryos were exposed to RA, chemical

RAR inhibitors or injected with mRNA encoding dominant-negative mutant versions of

RARs. In all cases, inhibition of RA signaling ablated both exocrine and endocrine gene expression in the dorsal pancreas, while the ventral pancreas was unaffected [50, 51]. This result correlated with ectopic shh expression in the dorsal pancreatic endoderm [50], suggesting that RA signaling contributes to the exclusion of Shh in the dorsal pancreas field.

In the reciprocal experiment, exposing Xenopus gastrulae to exogenous RA resulted in enlargement of the pancreas domain. This perturbation expanded the dorsal endocrine population at the expense of the exocrine population, as indicated by a dose-dependent decrease in exocrine cell markers and an increase in insulin expression [50].

Consistent with the role of RA in the frog, mice deficient in an enzyme required to synthesize RA (Retinaldehyde dehydrogenase; RALDH) exhibit dorsal pancreas hypoplasia, but retain pancreas markers in the ventral endoderm [52, 53]. Thus, Xenopus explant and pharmacological assays have facilitated clarification of the critical roles of a specific signaling pathway (RA) in pancreas specification and differentiation that are highly relevant to mammalian systems (see Figure 1).

182

2.4. Genome-wide microarray screens identify new foregut genes.

While sections 2.1-2.3 highlight a few of the key signaling pathways known to be involved in foregut specification and patterning, there is still a paucity of knowledge of all the molecular players and effectors involved in integrating and implementing these signals.

Fortunately, microarray analyses of Xenopus embryos, explants and developing organs are now being used to identify, on a genome-wide scale, hundreds of new factors likely to play important roles in foregut patterning and organogenesis in all vertebrates. These examples illustrate the power of frog embryos as a platform for unbiased gene discovery.

2.4.1. Microarray analyses of chemically-treated Xenopus embryos.

The ability to culture frog embryos in the presence of compounds that modulate specific signaling pathways facilitates straightforward profiling of genes downstream of these pathways. For example, recent microarray profiling of Xenopus embryos exposed to an RA inhibitor, identified Ndrg1α as a new RA-responsive factor [54]. Ndrg1α has diverse functions in development and tumorigenesis, but has not previously been associated with digestive organ development [55]. Interestingly, Ndrg1was found to repress the Wnt/β- catenin pathway allowing specification of foregut progenitor cells [54]. This study thereby revealed novel cross-talk between RA and Wnt signaling in foregut development. Given the central role of RA and Wnt signaling in foregut specification, it will be important to ascertain the role of Ndrg1α in the development of the digestive organs of higher vertebrates.

183

2.4.2. Microarray analyses of Xenopus explants.

It is well established that reciprocal signaling between the mesoderm and endoderm layers of the vertebrate PGT is crucial for its regional patterning [41]. To identify new molecules involved in mesoderm-endoderm signaling, Kenny et al cultured isolated endoderm explants versus endoderm/mesoderm explants from the Xenopus PGT and conducted microarray analyses to interrogate resultant differences in gene expression [56].

One endoderm gene upregulated in response to mesoderm was the Sfrp-related protein

Sizzled (Szl) [56]. Szl was found to be required for foregut organ specification downstream of mesodermal BMP signaling, a highly conserved developmental pathway critical for axial patterning and the development of multiple organs [57] . Like Wnt, BMP also plays dynamic roles in foregut organ specification as BMP signaling initially promotes hindgut development and inhibits foregut fates [58], but is later required to specify foregut lineages [59, 60].

Interestingly, Szl maintains BMP signaling by regulating Fibronectin deposition between the endoderm and mesoderm layers of the PGT. In this example, the experimental amenability of Xenopus revealed a novel extracellular feedback mechanism that mediates reciprocal

Wnt/BMP crosstalk between the endoderm and mesoderm during foregut patterning.

2.4.3. Microarray analyses of Xenopus embryonic organs.

It is relatively easy to isolate individual prospective organs within the large and accessible PGT of the frog embryo. This approach has been exploited to discover a trove of new factors and cellular processes involved in the morphogenesis of the pancreas. For example, microarray profiling of ptf1a-positive pancreatic endoderm isolated from Xenopus

184 embryos revealed putative target genes of this important pancreas-specific transcription factor [61]. The genes identified in this analysis contribute to a surprising variety of cellular functions with, as yet, unexplored roles in pancreas morphogenesis, such as intracellular vesicle docking and fusion, metabolism, cell adhesion, and extracellular matrix stabilization

[61]. In another study, Jarikji et al used microarray technology to identify genes differentially expressed between isolated dorsal and ventral pancreatic buds of the Xenopus embryo [43].

This study identified Tetraspanin (Tn4sf3), a transmembrane scaffolding protein that is upregulated in ventral pancreatic tissue and, intriguingly, required for dorsal and ventral pancreatic fusion [43].

The above examples suggest that investigation of the molecules and pathways identified by microarray (or RNAseq) analyses in frog embryos could yield profound new insight into the regulatory networks and cellular processes required for the specification and morphogenesis of vertebrate foregut organs. Additional studies are necessary to determine the degree to which the new genes identified in these and other frog studies are conserved in higher vertebrates.

3. What can the frog tell us about intestinal lengthening?

In contrast to the multiple organs specified in the foregut region of the PGT, the posterior (midgut/hindgut) zone is destined to become intestine. This segment of the PGT must undergo dramatic morphogenetic changes, including lumen formation, extensive elongation and counterclockwise rotation while, concomitantly, the visceral mesoderm and

185 epithelial lining of the tract undergo lineage restriction and cellular differentiation. These concurrent events shape and integrate multiple levels of biological organization, from cellular architecture to intricate three-dimensional anatomy. The mechanisms underlying the complex morphogenesis of the intestine are just beginning to be understood.

Intestinal development is pertinent to a variety of human afflictions. Congenital anomalies of intestinal morphogenesis include narrowing (stenosis) or occlusion (atresia) of the GI tract [62-66], deficits in the normal length of the intestine (congenital short bowel syndrome [67-73]), and intestinal rotation and fixation abnormalities, which occur in as many as 1 in 500 infants [67, 69, 74-79]. While malrotation itself is not always symptomatic, it predisposes affected individuals to volvulus, a life-threatening strangulation of the gut tube

[74]. In addition to birth defects, inflammatory bowel diseases (e.g., Crohn’s, ulcerative colitis) are increasingly common chronic conditions in both pediatric and adult populations; progressive complications from these disorders often require surgical resection of the damaged regions of the gut, leading to a shortened GI tract and attendant nutritional issues

[80-82]. Understanding the events that control normal intestinal development is therefore critical, not only for preventing the causes of common birth defects, but also for devising strategies to restore normal gut length in congenital or acquired short bowel syndromes.

The concentrically coiled anatomy of the pre-metamorphic Xenopus tadpole intestine is relatively simple compared to the visceral anatomy of higher vertebrates, yet it undergoes analogous elongation and rotation events, which occur over the course of only a few days [5,

83]. Because of its internal location, the gut can be challenging to visualize in amniotes, but tadpoles are transparent throughout organogenesis, allowing the cells of the PGT to be

186 labeled and tracked during morphogenesis. Combined with the ability to target LOF/GOF reagents to the gut by microinjection, such studies have led to a deeper understanding of the cellular and molecular events that drive intestine development.

3.1. Endoderm cell rearrangements drive gut lengthening.

Despite early differences in the initial formation of the PGT in amphibian and amniotic embryos [84-86], there are remarkable similarities in the process of gut elongation.

In both frogs and mammals, the early gut tube narrows and elongates coincident with an apparent remodeling of the endoderm (i.e., future epithelial) layer, suggesting that a cell rearrangement process may be involved in vertebrate gut elongation [87, 88]. In the frog, very little cell division is observed during the early stages of gut elongation, supporting the idea that (at least initially), this event is driven almost exclusively by cell rearrangement [89,

90]. In the Xenopus embryo, small groups of gut cells can be easily labeled with vital dyes, and their behavior visualized during gut elongation. Such studies revealed that the most central endoderm cells of the PGT radially intercalate during gut morphogenesis [85]. As the gut lengthens, the number of endoderm cell layers is reduced from 4-5 cells deep to a single epithelial layer, suggesting that the intercalation of the central cells and concomitant thinning of the epithelium provides the increased surface area necessary to generate length [85, 90].

Moreover, clusters of labeled endoderm cells become aligned in longitudinal tracts along the

A-P axis of the lengthening intestine [85, 90]. This indicates that radial intercalation is biased to preferentially occur between A-P neighbors, or is closely coordinated with a convergent extension process, increasing intestinal length, rather than girth [85, 90, 91] (Figure 2).

187

In amphibian embryos, the PGT begins as a solid cylinder full of concentrically stratified endoderm cells, but in amniotes, the endoderm lining of the PGT is already a single-layer, albeit pseudostratified, epithelium[92]; thus, multilayer radial intercalation per se is not likely to drive gut elongation, but cell rearrangements are likely still involved. It has been hypothesized that the elongation of the mammalian gut tube results from the reorganization of the pseudostratified endoderm layer into a columnar epithelium, but how this results in anisotropic tissue lengthening is not known [92]. Nonetheless, short gut phenotypes in both frogs and mice are associated with disorganization and stratification of the intestinal epithelium [37, 88, 92, 93], and the genes and pathways found to direct endoderm rearrangement in frogs (by regulating cell shape/polarity/adhesion—discussed in

Sections 3.2-3.4 below) also control gut elongation and epithelial morphogenesis in mammalian models. Therefore, despite differences in endoderm tissue architecture in the initial PGT, the processes of gut elongation appear to be conserved at many levels, making the frog embryo a highly relevant model for investigating intestine lengthening.

3.2. Shroom3 mediates endoderm cell shape changes

The PDZ-containing protein Shroom3 is required for directing cell shape changes, including apical constriction and apicobasal cell elongation during neural tube closure [94-

96]. In Xenopus, intestine-targeted microinjection of mRNA encoding a dominant-negative mutant form of Shroom3 (DN-Shroom3) results in severely shortened intestinal tracts, demonstrating a requirement for this protein during gut development [97]. At the cellular level, DN-Shroom3 expressing cells display reduced apical constriction, or remain rounded

188 and do not intercalate, resulting in a stratified disorganized epithelium; thus, Shroom3 must normally direct cell shape changes in the endoderm that are necessary for both epithelial morphogenesis and gut elongation [97]. Interestingly, Pitx transcription factors, which are also required for intestinal elongation [98, 99], were found to directly regulate shroom3 expression, suggesting that Shroom3 directs intestinal morphogenesis downstream of Pitx factors [97]. These results underscore the relationship between endoderm morphogenesis and gut elongation. Notably, Shroom3 activity has also been correlated with endodermal shape changes and epithelial architecture in the mouse intestine [92, 100].

3.3. Non-canonical Wnt/PCP signaling controls intestine lengthening and endoderm rearrangements

In addition to foregut specification, Wnt signaling is also essential for intestinal morphogenesis as knock-out/down of Wnt signaling components results in short, malformed intestinal tracts in all species studied [37, 88, 90, 93, 101]. However, while canonical Wnt/- catenin signaling is necessary for cell specification and maintenance of stem cell niches in the intestine, the elongation of the gut tube depends primarily on the non-canonical Planar

Cell Polarity (Wnt/PCP) pathway, which involves distinct downstream effectors such as JNK and Rho family GTPases [102]. Isolating the mechanisms by which Wnt/PCP signaling coordinates cell movements in the gut is non-trivial in murine models due to the redundancy of Wnt signaling components, and the need for stage- and tissue-specific promoters and/or combinations of mutant alleles [37]. However, the use of small molecules and gut-specific

189 targeting of LOF reagents in Xenopus has contributed a mechanistic understanding of the molecular players and cellular mechanisms involved in gut elongation.

For example, both pharmacological inhibition of RhoA and gut-targeted microinjection of mRNA encoding a dominant negative mutant version of RhoA (DN-RhoA) implicate Rho activity in Xenopus gut elongation, as both perturbations abrogate intestinal elongation [90]. As they prepare to undergo intercalation, endoderm cells normally become polarized and radially oriented, progressing from the most basally located cells toward the center of the PGT [90]. However, DN-RhoA-expressing cells remain round in shape, unpolarized and do not intercalate. Rho-inhibited cells also exhibit aberrant Myosin II organization and increased expression of the adherens junction protein, E-cadherin, suggesting that, without Rho activity, endoderm cells are unable to remodel adhesive contacts and, as a result, cannot rearrange or intercalate [90]. Chemical inhibitors of downstream effectors of Rho kinase (ROCK) and Myosin II phenocopy Rho-deficient intestinal malformations at both the gross and cellular level, suggesting that actomyosin contractility regulated by the Rho/ROCK/Myosin II branch of the Wnt/PCP network is required for the endoderm cell rearrangements that generate gut length [90]. Interestingly, embryos with less severe gut elongation phenotypes display abnormal intestinal coiling, revealing a potential mechanistic link between the processes of gut lengthening and rotation.

Non-canonical Wnt/PCP signaling is also mediated by activation of Jun N-terminal kinase (JNK) [30, 103]. Chemical inhibition or gut-targeted knockdown of JNK activity results in shortened intestines, similar to perturbation of Rho/ROCK/Myosin II activity.

However, the JNK-deficient phenotype differs at the cellular level, as endoderm cell

190 adhesion is lost in guts lacking JNK activity, in contrast to the increased adhesion observed in ROCK-deficient guts [101]. Abrogation of microtubule polymerization phenocopies loss of JNK activity, and suggests that this arm of the Wnt/PCP signaling cascade mediates cell rearrangement by promoting microtubule polymerization, and maintaining cell adhesion

[101]. The amenability of Xenopus explants to ex vivo cell assays further confirmed that adhesive remodeling is likely to be involved in gut elongation. PGT cells were isolated and dissociated, and their ability to reaggregate was used as an assay for changes in cell adhesion that may be caused by perturbing different branches of the Wnt/PCP cascade. The results confirmed that Rho kinase promotes decreased cell-cell adhesion, while the JNK pathway increases adhesion, suggesting that these two arms of the Wnt/PCP signaling network act in complementary ways to regulate intestinal cell intercalation [101]. Thus, the accessibility of the frog embryo was instrumental in clarifying Wnt/PCP mediated cellular and molecular dynamics underlying the cell rearrangements that lengthen the gut.

3.4. Hedgehog signals mediate reciprocal mesoderm-endoderm signaling

Hedgehogs (Hhs) are secreted proteins that elicit concentration dependent responses via multi-pass transmembrane receptors [104]. Hh signaling plays crucial patterning and morphogenetic roles in ectoderm, mesoderm and endoderm-derived tissues throughout development [104]. In mouse models, loss of Hh signaling results in shortened, malrotated gastrointestinal tracts, with disrupted architecture in all three tissue layers [105, 106].

Xenopus have been instrumental in refining our understanding of how Hh-mediated communication between tissue layers functions in gut development. Hhs are expressed in the

191 endoderm layer of the gut tube but, surprisingly, microinjection of mRNA encoding a constitutively active version of the Smoothened receptor (to induce excessive Hh signaling) does not affect gut development when targeted to the endoderm layer [107]. In contrast, ectopic Hh signaling severely disrupts gut elongation and coiling when targeted to the mesoderm layer, indicating that Hh ligands from the endoderm act by binding to receptors in the mesoderm [107]. Of significance, this study also showed that Hh-mediated signaling within the mesoderm is essential for the endoderm epithelial rearrangements that elongate the intestine, revealing insight into the molecular nature of the reciprocal signaling known to be required between the layers of the developing gut [107].

Xenopus experiments also shed light on the role of downstream components of Hh signaling in gut morphogenesis. The transcription factor foxf1 is upregulated in the gut mesoderm in response to Hh signaling [108, 109]. However, knockout of foxf1 is lethal in mice prior to gastrointestinal elongation and looping, precluding use of the murine model for understanding the function of FoxF1 in intestinal development [110]. In contrast, FoxF1 can be directly knocked down in Xenopus using targeted microinjection of morpholino oligonucleotides. Morpholino-mediated loss of FoxF1 activity resulted in disruption of mesodermal differentiation and severely reduced the elongation and rotation of the Xenopus intestine, confirming that FoxF1 is essential for normal gastrointestinal morphogenesis [111].

Subsequent to this work, foxf2 knockouts and compound foxf1+/-/foxf2+/- mice were generated, which survive to birth [112]. Similar to the observations in Xenopus, the intestines of these mice were deformed and showed severe disruption in mesoderm-derived tissues

[112]. These examples underscore how the frog embryo can be used to discern the tissue-

192 specific functions of highly conserved genes in directing crucial aspects of gut morphogenesis.

4. Insights from chemical screening.

One of the greatest advantages of externally-fertilized, aquatic embryos, like those of the frog, is that they can be exposed to exogenous chemicals to reveal potential roles for the cellular target of the compound in development. For studies of organ morphogenesis, these are particularly useful reagents because they allow earlier events that might be dependent on the same signaling pathway, to proceed unperturbed. The effects of chemical reagents targeting RA, Wnt, and FGF signaling pathways have been described above, but a few studies using additional small molecules are also worth highlighting (section 4.1). Although the potential mechanisms of action of these compounds in disrupting gut specification or morphogenesis is not yet well understood, these reports nonetheless implicate interesting pathways and processes in digestive organogenesis. Finally, it is important to mention that numerous chemical toxicants, many of which have unknown mechanisms of action, also elicit gut phenotypes (section 4.2); these chemicals could provide interesting avenues for future research on both normal and abnormal gut development.

4.1.1 Calcineurin implicates Wnt/Ca++ signaling in gut elongation and rotation.

Calcineurin is a calcium/calmodulin-dependent serine/threonine phosphatase which is a component of the non-canonical Wnt/Calcium signaling pathway [113]. Exposure of

Xenopus tadpoles to the Calcineurin inhibitors cyclosporine A, FK506, or FK520 for six hour

193 windows beginning at Nieuwkoop and Faber (NF) stage 18, 29/30, 37/38, or 41 resulted in shortened gut tubes often displaying a reversed coiling direction [114]. Given the roles of non-canonical Wnt signaling in gut morphogenesis, it seems possible that Calcineurin regulates endoderm cell properties, such as adhesive or cytoskeletal dynamics, and/or cell polarity. Injection of these inhibitors into dorsal blastomeres at the four cell stage similarly disrupted gut coiling, supporting the idea that Calcineurin plays a specific role in gut development [114]. However, other dorsally derived organs (heart, liver, etc.) were also affected by this injection, and further experimentation is required to determine the importance of Calcineurin in gut morphogenesis per se. It will be interesting to determine how Wnt/Calcium signaling is integrated with canonical and/or non-canonical Wnt signaling in this context, using more specific reagents to target Calcineurin activity within the developing intestine.

4.1.2. Lysyl Oxidase and a role for the extracellular matrix in gut elongation.

Lysyl oxidase (Lox) is a copper-dependent enzyme that catalyzes cross-linkage of

Collagen and Elastin in the extracellular matrix (ECM). lox knockout mice die at birth and have a number of deformities including cleft palate, spinal, cardiovascular and respiratory defects, indicating that Lox is required for normal development in a number of systems [115-

117]. In Xenopus, exposure to β-aminoproprionitrile (β-APN), a specific inhibitor of the Lox catalytic domain, from NF 6-45 affects many developmental processes, including proper cross-linking of connective tissue fibers in the notochord and somites [118]. Of particular interest, β-APN exposure also results in short, straight gut tubes, suggesting that Lox may

194 also function in gut morphogenesis, but its role in intestinal development has not been specifically investigated. In mice, Lox regulates ECM organization in muscle connective tissue, suggesting that Lox could be required for proper ECM assembly between the mesoderm and endoderm layers of the PGT [119]. Additional studies that investigate the role of Lox specifically within the gut are necessary to evaluate the potential importance of this protein in intestinal development.

4.1.3. mTOR signaling is implicated in gut elongation

The mTORs are serine/threonine kinases which form complexes with FKBP12 and

Raptor [120-122]. In Xenopus, inhibition of mTOR with rapamycin treatment from NF 2 - 45 results in shorter, fatter gut tubes as compared with controls [123]. Other organs form relatively normally, suggesting that mTORs may be specifically required in gut elongation

[123]. Consistent with this idea, injection of a dominant-negative Rheb (an upstream positive regulator of mTOR) at the 2 cell stage also decreases intestinal elongation and coiling [123].

As zebrafish exposed to rapamycin also have GI defects, this pathway may play a conserved role in vertebrate intestinal organogenesis [124].

Interestingly, Sirtuin deacetylases (Sirts) are also implicated in regulation of the TOR pathway [125]. Inhibition of Sirt-1 with the specific inhibitor, Ex-527, from the 2 cell stage on also disrupts intestinal elongation and coiling, although Sirt-1 deficient tadpoles exhibit more general defects than those treated with rapamycin, including decreased lengthening along the anterior-posterior body axis, edema, and abnormal eye development [126]. As Sirt-

1 inactivates p53, and p53 activity is essential for normal embryonic development through

195 regulation of TGF-β signaling [127, 128], Sirt-1 could be involved in regulating multiple pathways during gut development. It is important to note that none of these experiments were conducted in a way that specifically evaluates the function(s) of Sirt-1/mTOR in GI morphogenesis, and it is unclear exactly when these reagents may be acting to impact gut development. Thus, additional studies that specifically address the functions of Sirts and

TORs in the gut are necessary to determine how this pathway may contribute to intestinal morphogenesis.

4.2 Numerous toxicants perturb gut morphogenesis

The etiology of intestinal malrotation is largely unknown but it is believed to have a multifactorial origin, implicating both genetic and environmental causes. Frog embryos have proven to be excellent models for screening toxicants that impact development, and many studies have implicated exogenous chemicals, including insecticides, nanoparticles and explosives, in digestive tract malformations (Table 1). Unfortunately, few of these reports attempt to identify the underlying molecular or cellular developmental mechanism(s) disrupted by toxicant exposure. Future research on the mechanism(s) of action of these compounds in Xenopus could provide invaluable insight into the pathways required for gut development and potentially reveal environmental factors that contribute to the relatively high incidence of intestinal malrotation and other gut defects in the human population.

196

Table B- 1) Anthropogenic toxicants found to disrupt gut development in Xenopus laevis

Chemical Class Chemicals tested Uses References Azoles Triadimefon, n-butyl Fungicide [129, 130] isocaynate, carbendazim

Bipyridyliums Paraquat Herbicide [131, 132] Carbamates Carbaryl Insecticide [133] Carboxylic Acids Valproic acid, pentanoic acid, Various: [134] butyric acid, 2-ethylhexanoic Plasticizer, acid lubricant Chlorophenoxy Acids 2,4-D Herbicide [129] Estrogen 17β - estradiol Drug [135] Nanoparticles CuO, ZnO, polystyrene Various: [136-138] semiconductors; drugs, skin care Nitroaromatic TNT, 2ADNT, 4ADNT Explosive [139] compounds Organochlorines Chlorothalonil, DDT, DDD Insecticide [139, 140] Organophosphates Malathion, Malaoxon, Insecticide [141-144] Parathion, Paraoxon, Dicrotophos, Monocrotophos, Chlorpyrifos, Diazinon Phenols Bisphenol A, nonylphenol Various: [135] Plastics, resins, adjuvant Phosphonoglycines Glyphosate Herbicide [129] Triazines Atrazine Herbicide [129, 145, 146] Polymer Mixtures Tire Debris Organic Extract Tire product [147] Chemical Mixtures Corexit 9500 Dispersant [148]

5. From Frogs to Humans.

Because the molecules, pathways and processes important for endoderm specification and digestive morphogenesis are conserved across vertebrates, amphibian studies are highly relevant to human gut development. Indeed, frog studies have already informed translational

197 research strategies. For example, knowledge of the hierarchical relationship of factors such as

Wnt, Fgf, RA, Bmp, and Hh in Xenopus endoderm specification was instrumental in the successful development of protocols to direct human pluripotent stem cells to digestive organ fates and generate digestive “organoids” from human stem cells [26]. Likewise, one of the most important genes in pancreatic development, pdx1, was discovered in the Xenopus model

[149]. Activation of this pancreatic master regulator is now used in human transdifferentiation protocols to elicit endocrine pancreas fates from extra-pancreatic tissues, such as liver cells, a critical step towards successfully reprogramming adult cells as a cell replacement therapy for diabetes [150].

Genes for human congenital GI malformations are just beginning to be identified

[151]. To complement these efforts, frog embryos not only provide excellent models for the discovery of genes critical for normal digestive organ morphogenesis (see section 2.4 above), but facilitate rapid in vivo validation of candidate birth defect gene function. For example, the importance of FoxF1 in gut morphogenesis was first demonstrated in Xenopus, as discussed above (section 3.4; [111]). Mutations in this gene have recently been detected in human patients with similar malformations, including intestinal malrotation and congenital short bowel [152-154]. Moreover, trisomy of chromosome 16, which contains the human foxf1 gene, is also associated with intestinal maladies [153] . Finally, mutations in zic3, a transcription factor involved in directing organ laterality in animal models [155], have recently been detected in humans with congenital GI defects [152]. In Xenopus, overexpression of zic3, or injection of a mutant zic3 mRNA that acts in a dominant-negative manner, disrupt the direction of intestinal looping, providing in vivo confirmation of the

198 suspected role of this molecule in organogenesis [155]. These examples illustrate the immense potential and relevance of the Xenopus model for human biomedical research.

6. What can frogs tell us about digestive organ evolution?

6.1. The diversity of the tadpole gut.

The morphology of the digestive tract determines an organism’s ability to assimilate the energy necessary to grow, survive and reproduce--and thus has a profound effect on fitness. For example, the dimensions of the gut tube itself, including its diameter, length and compartmentalization, impact the capacity to digest different food resources. Although there is remarkable disparity between these parameters in the GI tracts of different vertebrates, the underlying mechanism by which different topologies of the digestive tract evolve is unknown.

Frogs inhabit most of the planet, including every continent except Antarctica. This success is facilitated by a wide array of reproductive strategies which allows them to breed in diverse environments, including terrestrial niches [156]. Because the tadpoles of many terrestrial breeding frogs are derived from eggs laid in environments with limited water and food supplies (e.g., bromeliads, leaves), these species may exhibit unusual larval feeding strategies. For example, in contrast to typical herbivorous tadpoles, terrestrial tadpoles may be carnivorous and feed on unfertilized eggs, invertebrates or even other tadpoles.

Alternatively, they may delay or omit the feeding stage entirely by becoming more dependent on maternal yolk stores, as observed in direct-developing species [157]. Not surprisingly,

199 these novel feeding (or non-feeding) strategies are complemented by specialized larval gut morphologies. Evo-devo investigations of gut development in two emerging frog models,

Lepidobatrachus laevis and Eleutherodactylous coqui, are providing novel insight into the potential sources of variation that lead to diverse gut morphologies during evolution.

6.2. The evolution of a carnivorous foregut in Lepidobatrachus.

The Budgett’s frog, Lepidobatrachus laevis, lives in the semi-arid regions of South

America [158]. As adults, Lepidobatrachus are aggressive, and often cannibalistic, predators, while their tadpole larvae are obligate carnivores that routinely consume other tadpoles, including siblings [159-161]. Unlike tadpoles that have a long, un-compartmentalized tract adapted for their nutrient-poor herbivorous diet, Lepidobatrachus tadpoles have a large, distendable stomach compartment [161, 162]. Analysis of foregut morphogenesis in

Lepidobatrachus has revealed that the development of this unusual anatomy is preceded by a disparity in the proportion of the PGT that is ascribed to foregut versus hindgut, as compared to that observed in Xenopus (which is used as a point of comparison to represent the ancestral state [163]). This ultimately results in dramatic differences in stomach morphogenesis and the final anatomical orientation of the gastroduodenal (GD) loop (Figure 3).

To identify potential signaling pathways that may have been involved in the evolution of this novel carnivore morphology, a small molecule screen was conducted in Xenopus embryos [163]. Compounds targeting known morphogenetic pathways were screened for the ability to transform the more typical herbivore GD loop found in Xenopus to resemble that

200 found in the carnivorous Lepidobatrachus tadpole. Remarkably, five compounds produced this change, two of which inhibit RA signaling [163]. RA plays an early role in dorsal pancreas specification (described in section 2.3 above), but RALDH expression also persists throughout the development of the stomach and duodenum, and perturbations of RA signaling in tail bud stage Xenopus embryos implicate RA patterning in GD looping [164].

Thus decreased RA signaling in the Lepidobatrachus lineage may have led to the unusual carnivore foregut morphology, an idea supported by the formation of a smaller stomach and shallower GD loop (anatomically similar to an ancestral tadpole like Xenopus) in

Lepidobatrachus embryos exposed to exogenous RA [163]. This study demonstrated that subtle changes in the levels of a specific foregut signaling factor can lead to anatomical variants that closely mimic extant interspecific variation.

6.3. The evolution of nutritional endoderm in a direct-developing frog.

The epithelial lining of the vertebrate digestive tract arises from the endoderm germ layer of the early embryo. In the frog, the endoderm is derived from the cellularization of the yolky vegetal pole of the egg. In typical tadpoles, all of the endoderm-derived cells rearrange in the gut tube to become the gut epithelium and contribute to intestinal lengthening (see section 3.1 above). In most species, the inherent yolk in the embryonic cells supports development only through the initial tadpole stages, after which the animal needs a functional digestive system to acquire the energy necessary to continue to grow and, ultimately, metamorphose, when the long gut is remodeled into the shorter gut of an adult frog. In contrast, endotrophic (non-feeding, yolk-dependent) species, such as direct developers, often

201 delay or completely skip the formation of a long, coiled gut, since they do not need to feed

[157]. Instead, these embryos directly form a short adult-like gut by the time they become a froglet.

The mechanisms by which the processes of yolk utilization and gut development are altered in direct-developing species may provide insight into the origin of novel gut morphologies and feeding strategies during evolution. Indeed, in contrast to species like

Xenopus that produce feeding tadpoles, in the direct-developing embryo of

Eleutherodactylous coqui, much of the yolky endoderm does not contribute to the final epithelial lining of the gut tube [165, 166]. Instead, it becomes nutritional endoderm (NE) that is utilized solely as a source of energy—the yolk platelets in these cells are metabolized, extruded and, eventually, eliminated as waste from the body. Some of the yolky vegetal cells are specifically set aside for this function during blastula stages, as indicated by the existence of a population of endoderm with reduced signaling/responsiveness in such species [167].

This alternate fate of the NE is likely related to the modified germ layer patterning often seen in larger eggs [168]. The use of endoderm to provide energy to sustain growth, rather than form a longer gut, has interesting implications for the ancient origins of definitive vs. extraembryonic endoderm, as well as the acellular yolk sac of higher vertebrates [165].

7. Conclusion/ Future directions.

Amphibian embryos have a rich history as developmental models and have been instrumental in understanding fundamental embryological events, such as gastrulation and neurulation. Here, we argue that the experimental versatility of frog embryos—e.g., the

202 ability to isolate tissue explants, target LOF/GOF reagents, and/or use pharmacological agents to investigate the underlying mechanisms of development—also makes them ideal models in which to examine many facets of digestive organogenesis. Their amenability to these experimental manipulations has enhanced our understanding of the spatiotemporal dynamics of conserved signaling pathways, such as Wnt, FGF, BMP, RA and Hh, in foregut specification and intestinal elongation. In addition, microarray profiling and small molecule/toxicant screening in frog models have revealed novel proteins and pathways likely to play critical roles in normal and abnormal gut morphogenesis. Such information enhances our understanding of gut patterning and morphogenesis in all vertebrates, including humans, making the frog a powerful model for translational embryology.

While many signaling pathways are conserved in gut organogenesis, differences do exist in the size and shapes of digestive organs among species, which is exemplified by the gastrointestinal tracts of tadpoles with different feeding strategies. Studies designed to elucidate differences in gut development in frog species with unique feeding ecologies are beginning to provide intriguing insight into the variety of molecular and cellular mechanisms underlying morphological evolution. Such knowledge has the potential to illuminate specific environmental or ecological parameters that affect gut development and, therefore, impact survival and fitness. Since frog species are continuing to disappear at an alarming rate, this line of evo-devo research may provide critical information for conservation efforts [169].

Rapid advances in de novo transcriptome assembly, proteomics, and genome editing

(CRISPR-Cas) continue to make functional genetic studies even more accessible for

Xenopus, and nearly any frog species [170-175]. New techniques continue to arise for

203 refining tissue-specific gene manipulation at late stages of organogenesis in the frog, including lipofection [176] and electroporation [177]. Utilizing these cutting-edge technologies to investigate amphibian digestive organ development—integrating aspects of organ specification, morphogenesis, toxicology, and/or evolution—has the potential to advance multiple scientific disciplines.

204

Figure B- 1) Wnt, RA, and FGF pattern the foregut The primitive gut tube is regionalized along both A-P and D-V axes. During gastrulation, RA (red) is required for dorsal pancreas (DP; purple) specification, likely by inhibiting Shh expression. Slightly later, the foregut is distinguished from the hindgut by a gradient of Wnt signaling (orange). High posterior Wnt specifies the hindgut domain, while low anterior Wnt (yellow; limited by Sfrp5) signals through the Fzd7 receptor to promote foregut fates and initiate cellular morphogenesis. Finally, a gradient of FGF signaling from the neighboring cardiac/lateral plate mesoderm segregates ventral foregut organs; prolonged, higher levels of FGF are needed to specify liver (green) versus ventral pancreas (VP; blue).

205

Figure B- 2) Intestine lengthening involves Hedghog- and Wnt/PCP-mediated endoderm cell polarization, rearrangement and epithelial differentiation Initially, the endoderm cells of the prospective intestine are rounded, unpolarized and disorganized. Signaling via Hedgehogs (HH; from the endoderm) induces foxF1 expression in the surrounding mesoderm layer of the gut tube. This facilitates reciprocal signaling from the now differentiating visceral mesoderm, which regulates the rate of epithelial differentiation in the underlying endoderm. Concomitant non-canonical Wnt signals (presumably from the mesoderm) are required for the endoderm cells to become polarized, starting with the outermost (most basal) layers and progressing towards the center of the gut tube. Both actomyosin contractile forces, regulated by ROCK, and microtubule organization, regulated by JNK, are required to dynamically remodel adhesive contacts between the polarized endoderm cells. This enables productive radial intercalation of the most central cells into the outermost layer, resulting in tissue lengthening and the morphogenesis of a single layer intestinal epithelium.

206

207

Figure B- 3) Altered RA signaling may have led to a novel foregut morphology A) In the hypothetical ancestral anuran (represented by Xenopus), the herbivorous tadpole requires only a rudimentary stomach. The foregut domain of the primitive gut tube is small relative to the hindgut domain, causing the gastroduodenal (GD) loop to form in a relatively anterior position and acquire an acute curvature during later foregut morphogenesis. B) In contrast, in the carnivorous Lepidobatrachus tadpole, which requires a capacious stomach, the ratio of foregut to hindgut is greater, and the GD loop forms in a more posterior position. Consequently, the larger carnivore stomach becomes more transversely oriented. This anatomical change may have been dependent on a decrease in RA signaling during foregut development in the Lepidobatrachus lineage, since inhibiting RA in Xenopus (representative of the ancestral condition) transforms the GD loop to resemble that observed in Lepidobatrachus. Conversely, exposing Lepidobatrachus embryos to excess RA elicits a more typical foregut configuration.

208

Figure B- 4) Endoderm morphogenesis in ancestral versus direct-developing frog species A) In ancestral frogs that produce feeding (exotrophic) tadpoles, all of the yolky vegetal endoderm cells (yellow) in the primitive gut tube (PGT) are used to generate the lining of the tadpole gut. As these cells radially rearrange (see Figure 2) and differentiate into the final digestive epithelium (orange/red), a central lumen is formed and the intestine is lengthened to form a long, coiled tract. The extensive gut is eventually remodeled to a shorter adult tract during metamorphosis (not shown). B) In contrast, in the direct-developing (endotrophic) frog embryo, a subset of the vegetal endoderm cells are fated to become nutritional endoderm (NE; pink), a cell type that does not rearrange nor contribute to the gut epithelium. Instead, these cells are gradually depleted of their yolk, extruded and eliminated as waste. Consequently, the PGT does not generate a long tract, and the developing froglet directly forms a short adult-length intestine.

209

210

References

[1] Keller R. Cell migration during gastrulation. Current opinion in cell biology 2005;17:533- 41.

[2] Keller R, Davidson LA, Shook DR. How we are shaped: the biomechanics of gastrulation. Differentiation 2003;71:171-205.

[3] Spemann H, Mangold H. Induction of embryonic primordia by implantation of organizers from a different species. 1923. Int J Dev Biol 2001;45:13-38.

[4] Blitz IL, Andelfinger G, Horb ME. Germ layers to organs: using Xenopus to study "later" development. Seminars in cell & developmental biology 2006;17:133-45.

[5] Nieuwkoop PaF, J. Normal Table of Xenopus Laevis (Daudin). New York, NY: Garland Publishing Inc.; 1994.

[6] Moody SA. Fates of the blastomeres of the 32-cell-stage Xenopus embryo. Developmental biology 1987;122:300-19.

[7] Moody SA, Kline MJ. Segregation of fate during cleavage of frog (Xenopus laevis) blastomeres. Anat Embryol (Berl) 1990;182:347-62.

[8] Logan M, Mohun T. Induction of cardiac muscle differentiation in isolated animal pole explants of Xenopus laevis embryos. Development 1993;118:865-75.

[9] Milet C, Monsoro-Burq AH. Dissection of Xenopus laevis neural crest for in vitro explant culture or in vivo transplantation. J Vis Exp 2014.

[10] Saint-Jeannet JP, Karavanov AA, Dawid IB. Expression of mesoderm markers in Xenopus laevis Keller explants. Int J Dev Biol 1994;38:605-11.

[11] Sater AK, Jacobson AG. The specification of heart mesoderm occurs during gastrulation in Xenopus laevis. Development 1989;105:821-30.

[12] Sater AK, Jacobson AG. The restriction of the heart morphogenetic field in Xenopus laevis. Developmental biology 1990;140:328-36.

[13] Wilson PA, Oster G, Keller R. Cell rearrangement and segmentation in Xenopus: direct observation of cultured explants. Development 1989;105:155-66.

211

[14] Smith JC, Symes K, Hynes RO, DeSimone D. Mesoderm induction and the control of gastrulation in Xenopus laevis: the roles of fibronectin and integrins. Development 1990;108:229-38 . [15] Dush MK, McIver AL, Parr MA, Young DD, Fisher J, Newman DR, et al. Heterotaxin: a TGF-beta signaling inhibitor identified in a multi-phenotype profiling screen in Xenopus embryos. Chem Biol 2011;18:252-63.

[16] Tomlinson ML, Field RA, Wheeler GN. Xenopus as a model organism in developmental chemical genetic screens. Mol Biosyst 2005;1:223-8.

[17] Tomlinson ML, Guan P, Morris RJ, Fidock MD, Rejzek M, Garcia-Morales C, et al. A chemical genomic approach identifies matrix metalloproteinases as playing an essential and specific role in Xenopus melanophore migration. Chem Biol 2009;16:93-104.

[18] Wheeler GN, Brandli AW. Simple vertebrate models for chemical genetics and drug discovery screens: lessons from zebrafish and Xenopus. Developmental dynamics : an official publication of the American Association of Anatomists 2009;238:1287-308.

[19] Callery EM. There's more than one frog in the pond: A survey of the Amphibia and their contributions to developmental biology. Seminars in cell & developmental biology 2006;17:80-92.

[20] Benitez MS, Del Pino EM. Expression of Brachyury during development of the dendrobatid frog Colostethus machalilla. Developmental dynamics : an official publication of the American Association of Anatomists 2002;225:592-6.

[21] Elinson RP, del Pino EM. Developmental diversity of amphibians. Wiley Interdiscip Rev Dev Biol 2012;1:345-69.

[22] Cano DA, Hebrok M, Zenker M. Pancreatic development and disease. Gastroenterology 2007;132:745-62.

[23] Desmet VJ. Cystic diseases of the liver. From embryology to malformations. Gastroenterol Clin Biol 2005;29:858-60.

[24] Hezel AF, Kimmelman AC, Stanger BZ, Bardeesy N, Depinho RA. Genetics and biology of pancreatic ductal adenocarcinoma. Genes Dev 2006;20:1218-49.

[25] Nair RJ, Lawler L, Miller MR. Chronic pancreatitis. Am Fam Physician 2007;76:1679- 88.

[26] Wells JM, Spence JR. How to make an intestine. Development 2014;141:752-60.

212

[27] Zorn AM, Wells JM. Vertebrate endoderm development and organ formation. Annu Rev Cell Dev Biol 2009;25:221-51.

[28] Arterbery AS, Bogue CW. Endodermal and mesenchymal cross talk: a crossroad for the maturation of foregut organs. Pediatr Res 2014;75:120-6.

[29] Nusse R. Wnt signaling in disease and in development. Cell Res 2005;15:28-32. [30] van Amerongen R, Nusse R. Towards an integrated view of Wnt signaling in development. Development 2009;136:3205-14.

[31] McLin VA, Rankin SA, Zorn AM. Repression of Wnt/beta-catenin signaling in the anterior endoderm is essential for liver and pancreas development. Development 2007;134:2207-17.

[32] Li Y, Rankin SA, Sinner D, Kenny AP, Krieg PA, Zorn AM. Sfrp5 coordinates foregut specification and morphogenesis by antagonizing both canonical and noncanonical Wnt11 signaling. Genes Dev 2008;22:3050-63.

[33] Mii Y, Taira M. Secreted Frizzled-related proteins enhance the diffusion of Wnt ligands and expand their signalling range. Development 2009;136:4083-8.

[34] Mii Y, Taira M. Secreted Wnt "inhibitors" are not just inhibitors: regulation of extracellular Wnt by secreted Frizzled-related proteins. Dev Growth Differ 2011;53:911-23.

[35] Zhang Z, Rankin SA, Zorn AM. Different thresholds of Wnt-Frizzled 7 signaling coordinate proliferation, morphogenesis and fate of endoderm progenitor cells. Developmental biology 2013;378:1-12.

[36] Heller RS, Dichmann DS, Jensen J, Miller C, Wong G, Madsen OD, et al. Expression patterns of Wnts, Frizzleds, sFRPs, and misexpression in transgenic mice suggesting a role for Wnts in pancreas and foregut pattern formation. Developmental dynamics : an official publication of the American Association of Anatomists 2002;225:260-70.

[37] Matsuyama M, Aizawa S, Shimono A. Sfrp controls apicobasal polarity and oriented cell division in developing gut epithelium. PLoS genetics 2009;5:e1000427.

[38] Kim BM, Buchner G, Miletich I, Sharpe PT, Shivdasani RA. The stomach mesenchymal transcription factor Barx1 specifies gastric epithelial identity through inhibition of transient Wnt signaling. Dev Cell 2005;8:611-22.

[39] Dorey K, Amaya E. FGF signalling: diverse roles during early vertebrate embryogenesis. Development 2010;137:3731-42.

213

[40] Pownall ME, Isaacs HV. FGF Signalling in Vertebrate Development. San Rafael (CA)2010.

[41] Horb ME, Slack JM. Endoderm specification and differentiation in Xenopus embryos. Developmental biology 2001;236:330-43.

[42] Shifley ET, Kenny AP, Rankin SA, Zorn AM. Prolonged FGF signaling is necessary for lung and liver induction in Xenopus. BMC Dev Biol 2012;12:27.

[43] Jarikji Z, Horb LD, Shariff F, Mandato CA, Cho KW, Horb ME. The tetraspanin Tm4sf3 is localized to the ventral pancreas and regulates fusion of the dorsal and ventral pancreatic buds. Development 2009;136:1791-800.

[44] Kelly OG, Melton DA. Development of the pancreas in Xenopus laevis. Developmental dynamics : an official publication of the American Association of Anatomists 2000;218:615- 27.

[45] Pearl EJ, Bilogan CK, Mukhi S, Brown DD, Horb ME. Xenopus pancreas development. Developmental dynamics : an official publication of the American Association of Anatomists 2009;238:1271-86.

[46] Hebrok M. Hedgehog signaling in pancreas development. Mech Dev 2003;120:45-57. [47] Niederreither K, Dolle P. Retinoic acid in development: towards an integrated view. Nat Rev Genet 2008;9:541-53.

[48] Moriya N, Komazaki S, Takahashi S, Yokota C, Asashima M. In vitro pancreas formation from Xenopus ectoderm treated with activin and retinoic acid. Dev Growth Differ 2000;42:593-602.

[49] Moriya N, Komazaki S, Asashima M. In vitro organogenesis of pancreas in Xenopus laevis dorsal lips treated with retinoic acid. Dev Growth Differ 2000;42:175-85.

[50] Chen Y, Pan FC, Brandes N, Afelik S, Solter M, Pieler T. Retinoic acid signaling is essential for pancreas development and promotes endocrine at the expense of exocrine cell differentiation in Xenopus. Developmental biology 2004;271:144-60.

[51] Stafford D, Hornbruch A, Mueller PR, Prince VE. A conserved role for retinoid signaling in vertebrate pancreas development. Dev Genes Evol 2004;214:432-41.

[52] Martin M, Gallego-Llamas J, Ribes V, Kedinger M, Niederreither K, Chambon P, et al. Dorsal pancreas agenesis in retinoic acid-deficient Raldh2 mutant mice. Developmental biology 2005;284:399-411.

214

[53] Ostrom M, Loffler KA, Edfalk S, Selander L, Dahl U, Ricordi C, et al. Retinoic acid promotes the generation of pancreatic endocrine progenitor cells and their further differentiation into beta-cells. PLoS One 2008;3:e2841.

[54] Zhang T, Guo X, Chen Y. Retinoic acid-activated Ndrg1a represses Wnt/beta-catenin signaling to allow Xenopus pancreas, oesophagus, stomach, and duodenum specification. PLoS One 2013;8:e65058.

[55] Melotte V, Qu X, Ongenaert M, van Criekinge W, de Bruine AP, Baldwin HS, et al. The N-myc downstream regulated gene (NDRG) family: diverse functions, multiple applications. FASEB J 2010;24:4153-66.

[56] Kenny AP, Rankin SA, Allbee AW, Prewitt AR, Zhang Z, Tabangin ME, et al. Sizzled- tolloid interactions maintain foregut progenitors by regulating fibronectin-dependent BMP signaling. Dev Cell 2012;23:292-304.

[57] Hogan BL. Bone morphogenetic proteins: multifunctional regulators of vertebrate development. Genes Dev 1996;10:1580-94.

[58] Tiso N, Filippi A, Pauls S, Bortolussi M, Argenton F. BMP signalling regulates anteroposterior endoderm patterning in zebrafish. Mech Dev 2002;118:29-37.

[59] Chung WS, Shin CH, Stainier DY. Bmp2 signaling regulates the hepatic versus pancreatic fate decision. Dev Cell 2008;15:738-48.

[60] Wills A, Dickinson K, Khokha M, Baker JC. Bmp signaling is necessary and sufficient for ventrolateral endoderm specification in Xenopus. Developmental dynamics : an official publication of the American Association of Anatomists 2008;237:2177-86.

[61] Bilogan CK, Horb ME. Microarray analysis of Xenopus endoderm expressing Ptf1a. Genesis 2012;50:853-70.

[62] Boyden EA, Cope JG, Bill AH, Jr. Anatomy and embryology of congenital intrinsic obstruction of the duodenum. Am J Surg 1967;114:190-202.

[63] Carpenter HM. Pathogenesis of congenital jejunal atresia. Arch Pathol 1962;73:390-6.

[64] Dalla Vecchia LK, Grosfeld JL, West KW, Rescorla FJ, Scherer LR, Engum SA. Intestinal atresia and stenosis: a 25-year experience with 277 cases. Arch Surg 1998;133:490- 6; discussion 6-7.

[65] DeLorimier AA, Fonkalsrud EW, Hays DM. Congenital atresia and stenosis of the jejunum and ileum. Surgery 1969;65:819-27.

215

[66] Seashore JH, Collins FS, Markowitz RI, Seashore MR. Familial apple peel jejunal atresia: surgical, genetic, and radiographic aspects. Pediatrics 1987;80:540-4.

[67] Chu SM, Luo CC, Chou YH, Yen JB. Congenital short bowel syndrome with malrotation. Chang Gung Med J 2004;27:548-50.

[68] Hasosah M, Lemberg DA, Skarsgard E, Schreiber R. Congenital short bowel syndrome: a case report and review of the literature. Can J Gastroenterol 2008;22:71-4.

[69] Kern IB, Leece A, Bohane T. Congenital short gut, malrotation, and dysmotility of the small bowel. J Pediatr Gastroenterol Nutr 1990;11:411-5.

[70] van der Werf CS, Halim D, Verheij JB, Alves MM, Hofstra RM. Congenital Short Bowel Syndrome: from clinical and genetic diagnosis to the molecular mechanisms involved in intestinal elongation. Biochim Biophys Acta 2015;1852:2352-61.

[71] Palle L, Reddy B. Case report: Congenital short bowel syndrome. Indian J Radiol Imaging 2010;20:227-9.

[72] Sabharwal G, Strouse PJ, Islam S, Zoubi N. Congenital short-gut syndrome. Pediatr Radiol 2004;34:424-7.

[73] Schalamon J, Schober PH, Gallippi P, Matthyssens L, Hollwarth ME. Congenital short- bowel; a case study and review of the literature. Eur J Pediatr Surg 1999;9:248-50.

[74] Aslanabadi S, Ghalehgolab-Behbahan A, Jamshidi M, Veisi P, Zarrintan S. Intestinal malrotations: a review and report of thirty cases. Folia Morphol (Warsz) 2007;66:277-82.

[75] Filston HC, Kirks DR. Malrotation - the ubiquitous anomaly. J Pediatr Surg 1981;16:614-20.

[76] Ford EG, Senac MO, Jr., Srikanth MS, Weitzman JJ. Malrotation of the intestine in children. Ann Surg 1992;215:172-8.

[77] Kamal IM. Defusing the intra-abdominal ticking bomb: intestinal malrotation in children. CMAJ 2000;162:1315-7.

[78] Stewart DR, Colodny AL, Daggett WC. Malrotation of the bowel in infants and children: a 15 year review. Surgery 1976;79:716-20.

[79] Torres AM, Ziegler MM. Malrotation of the intestine. World J Surg 1993;17:326-31.

216

[80] O'Sullivan M, O'Morain C. Nutrition in inflammatory bowel disease. Best Pract Res Clin Gastroenterol 2006;20:561-73.

[81] Vernier-Massouille G, Balde M, Salleron J, Turck D, Dupas JL, Mouterde O, et al. Natural history of pediatric Crohn's disease: a population-based cohort study. Gastroenterology 2008;135:1106-13.

[82] Cosnes J, Gower-Rousseau C, Seksik P, Cortot A. Epidemiology and natural history of inflammatory bowel diseases. Gastroenterology 2011;140:1785-94.

[83] Chalmers AD, Slack JM. Development of the gut in Xenopus laevis. Developmental dynamics : an official publication of the American Association of Anatomists 1998;212:509- 21.

[84] Cervantes S. Cellular and molecular mechanisms of intestinal elongation in mammals: the long and short of it. Histol Histopathol 2013;28:427-36.

[85] Chalmers AD, Slack JM. The Xenopus tadpole gut: fate maps and morphogenetic movements. Development 2000;127:381-92.

[86] Roberts DJ. Molecular mechanisms of development of the gastrointestinal tract. Developmental dynamics : an official publication of the American Association of Anatomists 2000;219:109-20.

[87] Matsumoto A, Hashimoto K, Yoshioka T, Otani H. Occlusion and subsequent re- canalization in early duodenal development of human embryos: integrated organogenesis and histogenesis through a possible epithelial-mesenchymal interaction. Anat Embryol (Berl) 2002;205:53-65.

[88] Yamada M, Udagawa J, Matsumoto A, Hashimoto R, Hatta T, Nishita M, et al. Ror2 is required for midgut elongation during mouse development. Developmental dynamics : an official publication of the American Association of Anatomists 2010;239:941-53.

[89] Muller JK, Prather DR, Nascone-Yoder NM. Left-right asymmetric morphogenesis in the Xenopus digestive system. Developmental dynamics : an official publication of the American Association of Anatomists 2003;228:672-82.

[90] Reed RA, Womble MA, Dush MK, Tull RR, Bloom SK, Morckel AR, et al. Morphogenesis of the primitive gut tube is generated by Rho/ROCK/myosin II-mediated endoderm rearrangements. Developmental dynamics : an official publication of the American Association of Anatomists 2009;238:3111-25.

217

[91] Keller R, Davidson L, Edlund A, Elul T, Ezin M, Shook D, et al. Mechanisms of convergence and extension by cell intercalation. Philos Trans R Soc Lond B Biol Sci 2000;355:897-922.

[92] Grosse AS, Pressprich MF, Curley LB, Hamilton KL, Margolis B, Hildebrand JD, et al. Cell dynamics in fetal intestinal epithelium: implications for intestinal growth and morphogenesis. Development 2011;138:4423-32.

[93] Cervantes S, Yamaguchi TP, Hebrok M. Wnt5a is essential for intestinal elongation in mice. Developmental biology 2009;326:285-94.

[94] Haigo SL, Hildebrand JD, Harland RM, Wallingford JB. Shroom induces apical constriction and is required for hingepoint formation during neural tube closure. Curr Biol 2003;13:2125-37.

[95] Hildebrand JD, Soriano P. Shroom, a PDZ domain-containing actin-binding protein, is required for neural tube morphogenesis in mice. Cell 1999;99:485-97.

[96] Lee C, Scherr HM, Wallingford JB. Shroom family proteins regulate gamma-tubulin distribution and microtubule architecture during epithelial cell shape change. Development 2007;134:1431-41.

[97] Chung MI, Nascone-Yoder NM, Grover SA, Drysdale TA, Wallingford JB. Direct activation of Shroom3 transcription by Pitx proteins drives epithelial morphogenesis in the developing gut. Development 2010;137:1339-49.

[98] Campione M, Steinbeisser H, Schweickert A, Deissler K, van Bebber F, Lowe LA, et al. The homeobox gene Pitx2: mediator of asymmetric left-right signaling in vertebrate heart and gut looping. Development 1999;126:1225-34.

[99] Al Alam D, Sala FG, Baptista S, Galzote R, Danopoulos S, Tiozzo C, et al. FGF9-Pitx2- FGF10 signaling controls cecal formation in mice. Developmental biology 2012;369:340-8. [100] Plageman TF, Jr., Zacharias AL, Gage PJ, Lang RA. Shroom3 and a Pitx2-N-cadherin pathway function cooperatively to generate asymmetric cell shape changes during gut morphogenesis. Developmental biology 2011;357:227-34.

[101] Dush MK, Nascone-Yoder NM. Jun N-terminal kinase maintains tissue integrity during cell rearrangement in the gut. Development 2013;140:1457-66.

[102] Chien AJ, Conrad WH, Moon RT. A Wnt survival guide: from flies to human disease. J Invest Dermatol 2009;129:1614-27.

218

[103] Boutros M, Paricio N, Strutt DI, Mlodzik M. Dishevelled activates JNK and discriminates between JNK pathways in planar polarity and wingless signaling. Cell 1998;94:109-18.

[104] Briscoe J, Therond PP. The mechanisms of Hedgehog signalling and its roles in development and disease. Nat Rev Mol Cell Biol 2013;14:416-29.

[105] Ramalho-Santos M, Melton DA, McMahon AP. Hedgehog signals regulate multiple aspects of gastrointestinal development. Development 2000;127:2763-72.

[106] Sukegawa A, Narita T, Kameda T, Saitoh K, Nohno T, Iba H, et al. The concentric structure of the developing gut is regulated by Sonic hedgehog derived from endodermal epithelium. Development 2000;127:1971-80.

[107] Zhang J, Rosenthal A, de Sauvage FJ, Shivdasani RA. Downregulation of Hedgehog signaling is required for organogenesis of the small intestine in Xenopus. Developmental biology 2001;229:188-202.

[108] Mahlapuu M, Enerback S, Carlsson P. Haploinsufficiency of the forkhead gene Foxf1, a target for sonic hedgehog signaling, causes lung and foregut malformations. Development 2001;128:2397-406.

[109] Madison BB, McKenna LB, Dolson D, Epstein DJ, Kaestner KH. FoxF1 and FoxL1 link hedgehog signaling and the control of epithelial proliferation in the developing stomach and intestine. J Biol Chem 2009;284:5936-44.

[110] Mahlapuu M, Ormestad M, Enerback S, Carlsson P. The forkhead transcription factor Foxf1 is required for differentiation of extra-embryonic and lateral plate mesoderm. Development 2001;128:155-66.

[111] Tseng HT, Shah R, Jamrich M. Function and regulation of FoxF1 during Xenopus gut development. Development 2004;131:3637-47.

[112] Ormestad M, Astorga J, Landgren H, Wang T, Johansson BR, Miura N, et al. Foxf1 and Foxf2 control murine gut development by limiting mesenchymal Wnt signaling and promoting extracellular matrix production. Development 2006;133:833-43.

[113] Kuhl M. The WNT/calcium pathway: biochemical mediators, tools and future requirements. Front Biosci 2004;9:967-74.

[114] Yoshida Y, Kim S, Chiba K, Kawai S, Tachikawa H, Takahashi N. Calcineurin inhibitors block dorsal-side signaling that affect late-stage development of the heart, kidney,

219 liver, gut and somitic tissue during Xenopus embryogenesis. Development, growth & differentiation 2004;46:139-52.

[115] Maki JM, Rasanen J, Tikkanen H, Sormunen R, Makikallio K, Kivirikko KI, et al. Inactivation of the lysyl oxidase gene Lox leads to aortic aneurysms, cardiovascular dysfunction, and perinatal death in mice. Circulation 2002;106:2503-9.

[116] Maki JM, Sormunen R, Lippo S, Kaarteenaho-Wiik R, Soininen R, Myllyharju J. Lysyl oxidase is essential for normal development and function of the respiratory system and for the integrity of elastic and collagen fibers in various tissues. Am J Pathol 2005;167:927-36.

[117] Zhang J, Yang R, Liu Z, Hou C, Zong W, Zhang A, et al. Loss of lysyl oxidase-like 3 causes cleft palate and spinal deformity in mice. Hum Mol Genet 2015;24:6174-85.

[118] Geach TJ, Dale L. Members of the lysyl oxidase family are expressed during the development of the frog Xenopus laevis. Differentiation; research in biological diversity 2005;73:414-24.

[119] Kutchuk L, Laitala A, Soueid-Bomgarten S, Shentzer P, Rosendahl AH, Eilot S, et al. Muscle composition is regulated by a Lox-TGFbeta feedback loop. Development 2015;142:983-93.

[120] Hara K, Maruki Y, Long X, Yoshino K, Oshiro N, Hidayat S, et al. Raptor, a binding partner of target of rapamycin (TOR), mediates TOR action. Cell 2002;110:177-89.

[121] Kim DH, Sarbassov DD, Ali SM, King JE, Latek RR, Erdjument-Bromage H, et al. mTOR interacts with raptor to form a nutrient-sensitive complex that signals to the cell growth machinery. Cell 2002;110:163-75.

[122] Sabers CJ, Martin MM, Brunn GJ, Williams JM, Dumont FJ, Wiederrecht G, et al. Isolation of a protein target of the FKBP12-rapamycin complex in mammalian cells. J Biol Chem 1995;270:815-22.

[123] Moriyama Y, Ohata Y, Mori S, Matsukawa S, Michiue T, Asashima M, et al. Rapamycin treatment causes developmental delay, pigmentation defects, and gastrointestinal malformation on Xenopus embryogenesis. Biochemical and biophysical research communications 2011;404:974-8.

[124] Makky K, Tekiela J, Mayer AN. Target of rapamycin (TOR) signaling controls epithelial morphogenesis in the vertebrate intestine. Developmental biology 2007;303:501- 13.

220

[125] Ghosh HS, McBurney M, Robbins PD. SIRT1 negatively regulates the mammalian target of rapamycin. PLoS One 2010;5:e9199.

[126] Ohata Y, Matsukawa S, Moriyama Y, Michiue T, Morimoto K, Sato Y, et al. Sirtuin inhibitor Ex-527 causes neural tube defects, ventral edema formations, and gastrointestinal malformations in Xenopus laevis embryos. Development, growth & differentiation 2014;56:460-8.

[127] Takebayashi-Suzuki K, Funami J, Tokumori D, Saito A, Watabe T, Miyazono K, et al. Interplay between the tumor suppressor p53 and TGF beta signaling shapes embryonic body axes in Xenopus. Development 2003;130:3929-39.

[128] Wallingford JB, Seufert DW, Virta VC, Vize PD. p53 activity is essential for normal development in Xenopus. Current biology : CB 1997;7:747-57.

[129] Lenkowski JR, Sanchez-Bravo G, McLaughlin KA. Low concentrations of atrazine, glyphosate, 2,4-dichlorophenoxyacetic acid, and triadimefon exposures have diverse effects on Xenopus laevis organ morphogenesis. J Environ Sci (China) 2010;22:1305-8.

[130] Yoon CS, Jin JH, Park JH, Yeo CY, Kim SJ, Hwang YG, et al. Toxic effects of carbendazim and n-butyl isocyanate, metabolites of the fungicide benomyl, on early development in the African clawed frog, Xenopus laevis. Environ Toxicol 2008;23:131-44.

[131] Vismara C, Bacchetta R, Cacciatore B, Vailati G, Fascio U. Paraquat embryotoxicity in the Xenopus laevis cleavage phase. Aquat Toxicol 2001;55:85-93.

[132] Vismara C, Battista VV, Vailati G, Bacchetta R. Paraquat induced embryotoxicity on Xenopus laevis development. Aquat Toxicol 2000;49:171-9.

[133] Bacchetta R, Mantecca P, Andrioletti M, Vismara C, Vailati G. Axial-skeletal defects caused by Carbaryl in Xenopus laevis embryos. The Science of the total environment 2008;392:110-8.

[134] Dawson DA. Joint actions of carboxylic acid binary mixtures on Xenopus embryo development: comparison of joint actions for malformation types. Archives of environmental contamination and toxicology 1994;27:243-9.

[135] Sone K, Hinago M, Kitayama A, Morokuma J, Ueno N, Watanabe H, et al. Effects of 17beta-estradiol, nonylphenol, and bisphenol-A on developing Xenopus laevis embryos. Gen Comp Endocrinol 2004;138:228-36.

221

[136] Bacchetta R, Moschini E, Santo N, Fascio U, Del Giacco L, Freddi S, et al. Evidence and uptake routes for Zinc oxide nanoparticles through the gastrointestinal barrier in Xenopus laevis. Nanotoxicology 2014;8:728-44.

[137] Nations S, Wages M, Canas JE, Maul J, Theodorakis C, Cobb GP. Acute effects of Fe(2)O(3), TiO(2), ZnO and CuO nanomaterials on Xenopus laevis. Chemosphere 2011;83:1053-61.

[138] Tussellino M, Ronca R, Formiggini F, De Marco N, Fusco S, Netti PA, et al. Polystyrene nanoparticles affect Xenopus laevis development. Journal of Nanoparticle Research 2015;17.

[139] Saka M. Developmental toxicity of p,p'-dichlorodiphenyltrichloroethane, 2,4,6- trinitrotoluene, their metabolites, and benzo[a]pyrene in Xenopus laevis embryos. Environmental toxicology and chemistry / SETAC 2004;23:1065-73.

[140] Yu S, Wages MR, Cobb GP, Maul JD. Effects of chlorothalonil on development and growth of amphibian embryos and larvae. Environ Pollut 2013;181:329-34.

[141] Bonfanti P, Colombo A, Orsi F, Nizzetto I, Andrioletti M, Bacchetta R, et al. Comparative teratogenicity of chlorpyrifos and malathion on Xenopus laevis development. Aquat Toxicol 2004;70:189-200.

[142] Modra H, Vrskova D, Macova S, Kohoutkova J, Hajslova J, Haluzova I, et al. Comparison of diazinon toxicity to embryos of Xenopus laevis and Danio rerio; degradation of diazinon in water. Bull Environ Contam Toxicol 2011;86:601-4.

[143] Snawder JE, Chambers JE. Toxic and developmental effects of organophosphorus insecticides in embryos of the South African clawed frog. Journal of environmental science and health Part B, Pesticides, food contaminants, and agricultural wastes 1989;24:205-18.

[144] Snawder JE, Chambers JE. Critical time periods and the effect of tryptophan in malathion-induced developmental defects in Xenopus embryos. Life sciences 1990;46:1635- 42.

[145] Lenkowski JR, McLaughlin KA. Acute atrazine exposure disrupts matrix metalloproteinases and retinoid signaling during organ morphogenesis in Xenopus laevis. J Appl Toxicol 2010;30:582-9.

[146] Lenkowski JR, Reed JM, Deininger L, McLaughlin KA. Perturbation of organogenesis by the herbicide atrazine in the amphibian Xenopus laevis. Environmental health perspectives 2008;116:223-30.

222

[147] Mantecca P, Gualtieri M, Andrioletti M, Bacchetta R, Vismara C, Vailati G, et al. Tire debris organic extract affects Xenopus development. Environ Int 2007;33:642-8.

[148] Smith EE, Carr JA, Wages M, Wang JF, Murali S, Kendall R. Response of larval frogs to Corexit 9500. Toxicol Environ Chem 2012;94:1199-210.

[149] Wright CV, Schnegelsberg P, De Robertis EM. XlHbox 8: a novel Xenopus homeo protein restricted to a narrow band of endoderm. Development 1989;105:787-94.

[150] Horb ME, Shen CN, Tosh D, Slack JM. Experimental conversion of liver to pancreas. Curr Biol 2003;13:105-15.

[151] Dauve V, McLin VA. Recent advances in the molecular and genetic understanding of congenital gastrointestinal malformations. J Pediatr Gastroenterol Nutr 2013;57:4-13.

[152] Hilger AC, Halbritter J, Pennimpede T, van der Ven A, Sarma G, Braun DA, et al. Targeted Resequencing of 29 Candidate Genes and Mouse Expression Studies Implicate ZIC3 and FOXF1 in Human VATER/VACTERL Association. Hum Mutat 2015;36:1150-4.

[153] Martin V, Shaw-Smith C. Review of genetic factors in intestinal malrotation. Pediatric surgery international 2010;26:769-81.

[154] Stankiewicz P, Sen P, Bhatt SS, Storer M, Xia Z, Bejjani BA, et al. Genomic and genic deletions of the FOX gene cluster on 16q24.1 and inactivating mutations of FOXF1 cause alveolar capillary dysplasia and other malformations. Am J Hum Genet 2009;84:780-91.

[155] Kitaguchi T, Nagai T, Nakata K, Aruga J, Mikoshiba K. Zic3 is involved in the left- right specification of the Xenopus embryo. Development 2000;127:4787-95.

[156] McDiarmid RW, Altig R. Tadpoles : the biology of anuran larvae. Chicago: University of Chicago Press; 1999.

[157] Altig RJ, G. Guilds of Anuran Larvae: Relationships among Developmental Modes, Morphologies, and Habitats. Herpetological Monographs 1989;3:81-109.

[158] Budgett JS. Notes on the batrachians of the Paraguayan Chaco, with observations upon their breeding habits and development, especially with regard to Phyllomedusa hypochondrialis, Cope. Also a description of a new genus. QJ Microsc Sci 1899;305.

[159] Fabrezi JS. Morphological evolution of Ceratophyrinae (Anura, Neobatrachia). Journal of Zoological Systematics and Evolutionary Research 2006;44.

223

[160] Hanken J. Model Systems Versus Outgroups - Alternative Approaches to the Study of Head Development and Evolution. American Zoologist 1993;33.

[161] Ruibal RT, E. The obligate carnivorous larvae of the frog Lepidobatrachus laevis (Leptodactylidae). Copeia 1988;3.

[162] Carroll EJS, A.M.; Ruibal, R. Gastric pepsin in an anuran larva. Development Growth & Differentiation 1991;33:499.

[163] Bloom S, Ledon-Rettig C, Infante C, Everly A, Hanken J, Nascone-Yoder N. Developmental origins of a novel gut morphology in frogs. Evolution & development 2013;15:213-23.

[164] Lipscomb K, Schmitt C, Sablyak A, Yoder JA, Nascone-Yoder N. Role for retinoid signaling in left-right asymmetric digestive organ morphogenesis. Developmental dynamics : an official publication of the American Association of Anatomists 2006;235:2266-75.

[165] Buchholz DR, Singamsetty S, Karadge U, Williamson S, Langer CE, Elinson RP. Nutritional endoderm in a direct developing frog: a potential parallel to the evolution of the amniote egg. Developmental dynamics : an official publication of the American Association of Anatomists 2007;236:1259-72.

[166] Karadge U, Elinson RP. Characterization of the nutritional endoderm in the direct developing frog Eleutherodactylus coqui. Dev Genes Evol 2013;223:351-62.

[167] Chatterjee S, Elinson RP. Commitment to nutritional endoderm in Eleutherodactylus coqui involves altered nodal signaling and global transcriptional repression. J Exp Zool B Mol Dev Evol 2014;322:27-44.

[168] Elinson RP, Beckham Y. Development in frogs with large eggs and the origin of amniotes. Zoology (Jena) 2002;105:105-17.

[169] Stuart SN, Chanson JS, Cox NA, Young BE, Rodrigues AS, Fischman DL, et al. Status and trends of amphibian declines and extinctions worldwide. Science 2004;306:1783-6.

[170] Amin NM, Greco TM, Kuchenbrod LM, Rigney MM, Chung MI, Wallingford JB, et al. Proteomic profiling of cardiac tissue by isolation of nuclei tagged in specific cell types (INTACT). Development 2014;141:962-73.

[171] Blitz IL, Biesinger J, Xie X, Cho KW. Biallelic genome modification in F(0) Xenopus tropicalis embryos using the CRISPR/Cas system. Genesis 2013;51:827-34.

224

[172] Guo X, Zhang T, Hu Z, Zhang Y, Shi Z, Wang Q, et al. Efficient RNA/Cas9-mediated genome editing in Xenopus tropicalis. Development 2014;141:707-14.

[173] Wuhr M, Freeman RM, Jr., Presler M, Horb ME, Peshkin L, Gygi SP, et al. Deep proteomics of the Xenopus laevis egg using an mRNA-derived reference database. Curr Biol 2014;24:1467-75.

[174] Rao N, Song F, Jhamb D, Wang M, Milner DJ, Price NM, et al. Proteomic analysis of fibroblastema formation in regenerating hind limbs of Xenopus laevis froglets and comparison to axolotl. BMC Dev Biol 2014;14:32.

[175] Sun L, Bertke MM, Champion MM, Zhu G, Huber PW, Dovichi NJ. Quantitative proteomics of Xenopus laevis embryos: expression kinetics of nearly 4000 proteins during early development. Sci Rep 2014;4:4365.

[176] Ohnuma S, Mann F, Boy S, Perron M, Harris WA. Lipofection strategy for the study of Xenopus retinal development. Methods 2002;28:411-9.

[177] Chernet BT, Levin M. A versatile protocol for mRNA electroporation of Xenopus laevis embryos. Cold Spring Harb Protoc 2012;2012:447-52.

225

Appendix C: Developmental Constraints on Endoderm Morphogenesis Underlie the Evolution of Gut Length

Mandy Womble1, Cris Ledon-Rettig1, Jillian Hattaway1, Heidi Smith2, Michael Ryan2, and Nanette Nascone-Yoder1 1North Carolina State University; 2University of Texas

There is tremendous variation in the length of the digestive tract to accommodate diverse diets. To discern the developmental mechanisms that generate such variation during evolution, we assessed gut morphogenesis in an array of non-model frog species with distinct larval gut morphologies and feeding ecologies. Herbivorous tadpoles, the ancestral state, are typically derived from small eggs and have rapidly elongating guts. As their yolk reserves are quickly depleted, all the yolky endoderm cells that initially fill the gut tube become polarized and rearrange to lengthen the gut and form the epithelial lining. In contrast, tadpoles derived from slightly larger eggs exhibit delayed or decreased gut lengthening. In some species, this delay is a consequence of a larger endoderm mass taking longer to rearrange and complete epithelial morphogenesis, thus extending dependence on yolk reserves. However, in rapidly-developing cannibalistic species, the core of the endoderm never integrates into the gut epithelium; instead, many yolky cells are discarded by apoptosis, resulting in a shorter intestine with a larger lumen, a morphology adaptive for a carnivorous diet. Finally, the largest eggs harbor the greatest yolk reserves and often belong to direct- developing species that form a short adult-like digestive tract. In this context, cell polarization and organization occurs slowly within the bulky mass of yolky endoderm cells.

As in herbivores, the endoderm becomes polarized; however, cell rearrangement contributes

226 to increased epithelial surface area but does not drive tube lengthening. In general, tadpoles derived from larger eggs tend to have delayed gut lengthening or form shorter guts.

Variations in Sonic Hedgehog (Shh) signaling may influence the contribution of yolky endoderm cells to the mature gut epithelium as an increase in Sonic Hedgehog signaling in carnivores rescues apoptosis of the inner most cells and increases epithelial surface area.

Taken together, our data suggests that variation in gut morphology may have arisen due to developmental constraints on endoderm cell polarization and cell rearrangement and variations in Shh signaling and also implies that changes in scale during tubulogenesis may be an unexpected source of novel form and function during evolution.

227

Figure C- 1) Variation in tadpole feeding ecology correlates with variation in egg size (yolk reserves) and gut length Frogs inhabit every continent on Earth except for Antarctica and have evolved a variety of reproductive strategies in order to reproduce in diverse environments. The size of the egg and the amount of yolk reserves correlates with feeding strategy. (A) Most frogs lay smaller eggs that are about 1 mm in diameter into aqueous environments like a pond or lake. These embryos develop into tadpoles that have a long, coiled, yolk-depleted intestine in order to absorb the maximum amount of nutrition from the vegetable matter that they consume. B) Some frogs lay eggs in ephemeral or temporary pools of water. These embryos develop rapidly in order to metamorphose before the water evaporates. In many cases, these tadpoles are carnivorous (even cannibalistic). The eggs are typically about 2 mm in diameter. Their gastrointestinal tracts are yolk-depleted and compartmentalized, consisting of a more “adult- like” stomach in order to digest protein. Most tadpoles have a yolk supply in order to support the embryo until tadpole feeding stages. However, some tadpoles develop in environments without a food source, like in the rain water collected in pitcher plants or in foam nests and are endotrophic. C) Endotrophic frogs usually lay large eggs of up to 5 mm in diameter. Tadpoles with this developmental strategy will develop a short, yolk-filled, gut as the tadpoles will not eat and must rely on their yolk reserves for nutrients and energy until metamorphosis. D) Other frogs will skip the tadpole stage all together and develop directly from an embryo into a froglet inside of a thick egg capsule on dry land. These eggs are very large, up to 6mm in diameter, have massive yolk reserves that acts as nutritional endoderm and develop gastrointestinal tracts with a short, adult-like morphology. E) Box and Whiskers plot comparing differential developmental/feeding strategies to gastrointestinal tract physiology and egg size. Frogs that employ traditional developmental strategies (i.e. herbivorous tadpoles) and have long coiled intestinal tracts, typically have the smallest eggs. Carnivorous tadpoles and those that employ endotrophic tadpole feeding strategies both have shortened gastrointestinal lengths and moderately sized eggs, although yolk reserves differ. Frogs that employ direct development strategies with shortened “adult-like” gut morphology have the largest eggs. (Figure was adapted with permission from Nanette-Nascone Yoder).

228

229

Figure C- 2) All endodermal cells contribute to the gut epithelium in exotrophic and terrestrial, delayed herbivorous tadpoles, regardless of egg size Embryos prior to lumen formation (GS 20) (A, C, E) and at tadpole feeding stages (GS 25) (B, D, E) were analyzed for endodermal contribution to the final gut epithelium in three different exotrophic herbivorous tadpoles with different egg sizes by performing immunohistochemistry on transverse cross sections (G, H, I, J, K ,L) and accessing cellular morphology (-tubulin (a-tub): green, -catenin (b-cat): red, topro-3 (nuclei): blue). White lines indicate thickness of epithelium. (G, H, I) Prior to lumen formation, endoderm cells are multiple layers thick, spanning the entire primitive gut tube before beginning to polarize at the basement membrane. Polarization progresses inward as the cells radially intercalate and form a single layered epithelium with multiple gastrointestinal loops and a central lumen. (J, K, L) Regardless of size, all of the endodermal cells eventually contribute to the final gut epithelium and, presumably, contribute to gut length (GS 25). In smaller (X. laevis, A-B, G, J), and moderately sized embryos (E. pustulosus, C-D, H, K), by tadpole feeding stages (GS 25) all cell have become incorporated into a single layered epithelium. In larger embryos that are terrestrial (e.g. eggs laid on or in plants), like those of T. corticale, (E-F, I, L), complete cellular intercalation and integration takes longer and is delayed, as some gut loops are single-layered (*) and some are still multiple layers thick (**) at this stage. This is likely due to the large size and increased number of cells layers that need to undergo radial intercalation. Scale bars: 75m.

230

231

Figure C- 3) The innermost endoderm cells do not contribute to the gut epithelium in rapidly-developing, exotrophic carnivorous tadpoles, regardless of egg size Embryos prior to lumen formation (GS 20) (A, C, E) and at tadpole feeding stages (GS 25) (B, D, E) were analyzed to determine if all endodermal cells of the primitive gut tube contributed to the final single-layered intestinal epithelium. Three different carnivorous tadpoles with different egg sizes were analyzed (H. boettgeri: oophagus or egg-eating, A,B,G,J,M; C. cranwelli: carnivorous, C,D,H,K,N; and L. laevis: cannibalistic, E,F,I,L,O) by performing immunohistochemistry on transverse cross sections (G,H,I,J,K,L) and accessing cellular morphology (-tubulin (-tub): green, smooth muscle actin (SMAC): green, - catenin (-cat): red, caspase-3 (cas): red, topro-3 (nuclei): blue). At GS 20, endodermal cell fill the entire primitive gut tube in multiple cell layers (G- I). The number of cells layers is proportionate to the size of the embryo. At NF 25 (J-O), the innermost endodermal cells undergo apoptosis (white arrow) to form an enlarged lumen and do not contribute to the final epithelium, resulting in a shortened gut. The number of cells that undergo apoptosis appears to be correlated with embryo size. Scale bars: 75 m.

232

233

Figure C- 4) Endoderm rearrangement increases epithelial surface area, but not length, in endotrophic (direct developing) tadpoles In order to determine if the entire population of endodermal cells contributes to the final intestinal epithelium of large, endotrophic, direct developing embryos (E. coqui), immunohistochemistry was performed on transverse cross sections at the “yolk sac” stage (A) and the “yolk resorption” stage (B) (-tubulin: green; caspase-3: red, and topro-3 (nuclei): blue). During the “yolk sac” stage, the gut tube is completely occluded with endodermal cells (C, E). During the “yolk resorption” stage, tall, elongated cells align in a single layered epithelium (D, F). All cells contribute to the final epithelium but form a convoluted tube, increasing surface area at the expense of length. Scale bars (C-D): 675 m. Scale bars (C-F): 75 m

234

235

Figure C- 5) Sonic Hedgehog signaling increases intestinal surface area and reduces apoptosis in cannibalistic tadpoles In a chemical genetics screen, Sonic Hedgehog (Shh) was found to have an effect on gut length. In fact, X. laevis embryos treated with a Shh signaling antagonist (Cyclopamine) caused the gastrointestinal tract to become shortened in length (data not shown). Additionally the innermost endoderm cells underwent apoptosis (data not shown), similar to normal gastrointestinal organogenesis in carnivorous species. In order to determine the effect of increased Shh signaling on carnivore gut morphogenesis, L. laevis embryos were treated with the hedgehog signaling agonist, Purmorphomine, (B) or an equal volume of a DMSO solvent control (A). In control embryos (A), the gut developed normally, with a shortened, “carnivore-typical” morphology. In embryos treated with Purmorphomine (B), the gastrointestinal did not increase in length, however, surface area was increased. In order to determine the cellular level effects of increased Shh signaling, immunohistochemistry was performed on transverse cross sections (E-cadherin: green, topro (nuclei): blue) (C-F). In control embryos (C, E), the innermost endodermal cells undergo apoptosis instead of contributed to the mature, single-layered gut epithelium. In embryos treated with Purmorphomine (D, F), these innermost cells are rescued from apoptosis, as all of the cells are incorporated into the final epithelium. Interestingly, instead of contributing to length, the intestine is convoluted and “folded” on itself with increased epithelial surface area, similar to the normal gastrointestinal morphology in direct developing, endotrophic species (Fig. C4). Scale bars: 75 m.

236

237

Figure C- 6) Changes in endoderm morphogenesis may underlie anuran gut evolution Gastrointestinal morphology varies depending on tadpole diet and developmental strategy (i.e. herbivore, carnivore/cannibal, terrestrial/delayed, or direct developer). In all tadpoles, prior to lumen formation, endodermal cells completely fill and occlude the primitive gut tube. Cells become polarized and radially intercalate with their neighbors, starting at the basement membrane and progressing towards the forming central lumen. In all tadpoles studied, except for carnivores, all cells of the endoderm contribute to the final gastrointestinal tract epithelium. In carnivores, especially the cannibalistic tadpole, L. laevis, innermost endoderm cells undergo apoptosis, perhaps due to reduced Hedgehog signaling. In guts with more yolk (from larger eggs), cells tend to rearrange more slowly, as more cell layers need to undergo radial intercalation. In direct, developing endotrophs, the endoderm contributes more to epithelial surface area than to length. Therefore, changes in egg size and the method or degree to which endodermal/yolky cells are incorporated into the final intestinal epithelial may guide anuran gut evolution.

238

239

Appendix D: Foregut gene expression in X. laevis. Mandy Womble

The cellular level morphogenetic mechanisms of foregut organ development are just beginning to be understood. The foregut begins as an anterior region of the primitive gut tube

before being regionalized through the expression of tissue and organ specific genes. The foregut is divided into the esophagus and stomach. Many accessory organs also bud off of the developing foregut including the lungs, liver, and pancreas. In order to interpret cellular level morphogenetic events, it is important to first understand the anatomy of both transverse and frontal cross sections. In the following figures, I used in situ hybridization at multiple stages of organogenesis in the model frog Xenopus laevis to probe for a variety of genes including:

the liver specific transcription factor, hhex; the glycoprotein, fibrinogen, known to be expressed in the liver and intestine; the LR determining transcription factor, pitx2; a marker

for vein angiogenesis, msr (aplnr); mesodermally expressed transcription factors, barx and

foxf1; the pancreas specific transcription factor, ptf1, and the signaling ligand, shh. The

expression patterns of these genes allow visualization of the anatomic landmarks and localization of organs during development that will be vital to interpreting other experimental

data including histological images and embryonic manipulations.

240

Figure D- 1) Gene Expression patterns of transverse cross sections at NF. 35.36 Transverse sections through embryos at NF 35.36 were processed through in situ hybridization probing for foxf1 (A-I), shh (J-Q), hhex (R-Y), pitx2 (Z-FF), ptf1 (GG-LL), or

msr (MM-TT). Sections are displayed anterior to posterior. Hrt- Heart, Eso- Esophagus,

NtCh- Notochord, MG- Midgut, Liv- Liver, Sto- Stomach, Panc- Pancreas, D. Panc- Dorsal

Pancreas, GB- Gall Bladder, A-Anterior, P- Posterior, R-Right, L-Left.

241

Figure Continued on pg. 242

242

243

Figure D- 2) Gene expression patterns of frontal sections at NF 37.38. Frontal sections through embryos at NF 37.38 were processed through in situ hybridization

probing for barx (A-K), Fibrinogen (L-W), ptf1 (X-HH), or pitx2 (II-TT). Sections are displayed dorsal to ventral. Eso- Esophagus, MG- Midgut, Sto- Stomach, Panc- Pancreas, A-

Anterior, P- Posterior, R-Right, L-Left.

244

Figure Continued on pg. 245

245

246

Figure D- 3) Gene expression patterns of transverse cross sections at NF 37.38 Transverse sections through embryos at NF 37.38 were processed through in situ hybridization probing for hhex (A-I), pitx2 (J-N), ptf1 (O-V), or msr (W-CC). Sections are

displayed anterior to posterior. Hrt- Heart, Eso- Esophagus, MG- Midgut, Liv- Liver, Sto-

Stomach, Panc- Pancreas, D. Panc- Dorsal Pancreas, GB- Gall Bladder, A-Anterior, P-

Posterior, R-Right, L-Left.

247

Figure continued on pg. 248

248

249

Figure D- 4) Gene expression patterns of frontal sections at NF 39 Frontal wax sections through embryos at NF 39 were processed through in situ hybridization probing for fibrinogen (A-J), ptf1 (K-T), or pitx2 (U-DD). Sections are displayed dorsal to ventral. Eso- Esophagus, MG- Midgut, Sto- Stomach, Dor Panc- Dorsal Pancreas, Ven Panc-

Ventral Pancreas, A-Anterior, P- Posterior, R-Right, L-Left.

250

Figure continued on pg. 251

251

252

Figure D- 5) Gene expression patterns of transverse cross sections at NF 39.40 Transverse wax sections through embryos at NF 39.40 were processed through in situ

hybridization probing for foxf1 (A-E), shh (F-H), hhex (I-M), ptf1 (N-R), or msr (S-W).

Sections are displayed anterior to posterior. Hrt- Heart, Eso- Esophagus, MG- Midgut, Liv-

Liver, Sto- Stomach, Panc- Pancreas, D. Panc- Dorsal Pancreas, GB- Gall Bladder, A-

Anterior, P- Posterior, R-Right, L-Left.

253

254

Figure D- 6) Gene expression patterns of frontal sections at NF 40.41 Frontal wax sections through embryos at NF 40.41 were processed through in situ hybridization probing for barx (A-M), fibrinogen (N-Z), ptf1 (AA-MM), or pitx2 (NN-ZZ).

Sections are displayed dorsal to ventral. Eso- Esophagus, MG- Midgut, Sto- Stomach, Dor

Panc- Dorsal Pancreas, Ven Panc- Ventral Pancreas, A-Anterior, P- Posterior, R-Right, L-

Left.

255

Figure Continued on pg. 256

256

257

Figure D- 7) Gene expression patterns of transverse cross sections at NF 40.41 Transverse sections through embryos at NF 40.41 were processed through in situ

hybridization probing for fibrinogen (fib) (A-J), ptf1 (K-T), pitx2 (U-DD), or barx (EE-

NN). Sections are displayed anterior to posterior. Hrt- Heart, Eso- Esophagus, MG- Midgut,

Liv- Liver, Sto- Stomach, Panc- Pancreas, D. Panc- Dorsal Pancreas, GB- Gall Bladder, A-

Anterior, P- Posterior, R-Right, L-Left.

258

Figure Continued on pg. 259

259

260

Figure D- 8) Gene expression patterns of transverse cross sections at NF 41 Transverse sections through embryos at NF 41 were processed through in situ hybridization

probing for hhex (A-E), pitx2 (F-J), ptf1 (K-O), msr (P-T), or foxf1 (U-Y). Sections are

displayed anterior to posterior. Hrt- Heart, Eso- Esophagus, MG- Midgut, Liv- Liver, Sto-

Stomach, Panc- Pancreas, D. Panc- Dorsal Pancreas, GB- Gall Bladder, A-Anterior, P-

Posterior, R-Right, L-Left.

261