Cytokine Signaling Plays Central Roles in the Initiation of Limb Regeneration

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Citation Tsai, Stephanie. 2019. Cytokine Signaling Plays Central Roles in the Initiation of Limb Regeneration. Doctoral dissertation, Harvard University, Graduate School of Arts & Sciences.

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Cytokine signaling plays central roles in the

initiation of limb regeneration

A dissertation presented

by

Stephanie Tsai

to

The Department of Molecular and Cellular Biology

in partial fulfillment of the requirements

for the degree of

Doctor of Philosophy

in the subject of

Biology

Harvard University

Cambridge, Massachusetts

April 2019

© 2019 Stephanie Tsai

All rights reserved.

Dissertation Advisor: Professor Douglas A. Melton Stephanie Tsai

Cytokine signaling plays central roles in the initiation of limb regeneration

Abstract

Newts and salamanders possess the unique ability amongst vertebrates to regenerate their limbs. While mammals respond to limb amputations with a scarring response, salamanders instead form an undifferentiated cellular structure distal to the amputation plane known as the blastema, which eventually differentiates into a new limb. Formation of the blastema is a complex process requiring coordinated migration and proliferation of progenitors derived from many tissues (muscle, bone, connective tissue etc.) and the exact molecular mechanisms underlying the initiation of this process remain unknown.

Here, we have elucidated the distinct transcriptional programs that are active in blastemal progenitors as well as the surrounding tissues during early stages of regeneration. These analyses suggested that dividing progenitors selectively establish an autocrine TGF- signaling network and further revealed the enriched expression of several cytokines, indicating progenitors may play an immunomodulatory role.

Functional analyses of the role of one such cytokine, interleukin-8 (il-8), indeed led us to discover that blastemal progenitors are immunomodulators during early stages of regeneration through interleukin-8/cxcr-1/2 signaling.

We also examined how these gene expression programs change when we prevented the formation of the wound epidermis, an important epithelial support structure for blastema formation. Our data suggest that the early wound epidermis plays

iii a large role in modulating inflammation, ECM degradation, and tissue histolysis. These analyses further identified the decreased expression of one growth factor cytokine, midkine (mk), in the absence of the wound epidermis. Gain- and loss-of-function analyses of mk during limb regeneration revealed that it is a critical regulator of wound epidermis maturation during blastema formation. Altogether, these studies emphasize the importance of cytokine signaling in the initiation of limb regeneration and provide the field with resources that may lead to the discovery of more putative regulators of blastema formation.

iv Table of Contents

Title page………………………………………………...... ………………...... i

Copyright………………………………………………...... ………………...... ii

Abstract………………………………………………...... ………………...... iii Table of contents………………………………………………...... ………………...... v List of Figures………………………………………………...... …...... viii

List of Tables………………………………………………...... ………………...... xi Abbreviations………………………………………………...... …………...... xii Acknowledgements………………………………………………...... …………...... xiii

Dedication ………………………………………………...... xvii

Epigraph………………………………………………...... xviii

Chapter 1 Introduction………………………………………………………………...... 1 1.1. Introduction to the Mexican axolotl…………………………………………...... 2 1.2. Structure of the axolotl limb………………………………………...……...... 5 1.3. Stages of limb regeneration…………………………………………………...... 5 1.4. The heterogeneous cellular origins of the blastema……………………...... 8 1.5. Essential components for successful blastema formation…………...... 10

1.5.1. Nerve dependence……………………………………………………...... 10 1.5.2. Wound epidermis/AEC………………………………………………...... 11 1.5.3. Injury…………………………………………………………………...... 14 1.6. Immunological influences on regeneration...... 16

1.7. Topics addressed in this dissertation……………...………..……………...... 19

1.8. References………………………………………………………...... 20

Chapter 2 Transcriptional profiles of early regenerating limb tissues...... 28

2.1. Preface……………………………………………………………………………...... 29

2.2. Abstract…………………………………………………………………………...... 29

2.3. Introduction……………………………………………………………………...... … 30

2.4. Results...... 32 2.3.1. Transcriptional profiling of dividing cells during the initiation of limb regeneration……………………………………………………...... …32 2.3.2. Growth factor signaling pathways are largely repressed within

regenerating stump tissues………………………………………...... 36 2.3.3. The transcriptional landscape of early dividing cells indicates

roles in shaping the blastemal niche...... 38

2.5. Discussion...... 41

2.6. Materials and methods...... 43

2.7. References...... 48

Chapter 3 Blastemal progenitors modulate immune signaling during early

limb regeneration...... 55

3.1. Preface...... 56

3.2. Abstract...... 56

v 3.3. Introduction...... 56

3.4. Results...... 59 3.4.1. Interleukin-8 (il-8) is expressed in early blastemal progenitors...... 59 3.4.2. IL-8 is sufficient to induce myeloid cell recruitment and proliferation in bone/perichondrium and epidermis in non-

regenerating limbs...... 61 3.4.3. Knockdown of IL-8 results in delayed blastemal outgrowth and regeneration...... 66 3.4.4. IL-8 knockdown leads to defective retention of myeloid cells during the wound healing transition to blastema formation...... 68 3.4.5. CXCR-1/2 signaling is necessary for limb regeneration...... 70

3.5. Discussion...... 74 3.6. Materials and methods...... 76 3.7. References...... 81

Chapter 4 Wound epidermis-dependent transcriptional programs during

early limb regeneration...... 86

4.1. Preface...... 87 4.2. Abstract...... 87 4.3. Introduction...... 88

4.4. Results...... 90 4.4.1. Full skin flap sutured limbs exhibit divergent transcriptional

profiles from normal regenerating limbs...... 90 4.4.2. Early dividing cells fail to activate key signaling pathways

in the absence of the wound epidermis...... 93 4.4.3. Non-dividing cells in regenerating stump tissues display aberrant inflammation and extracellular matrix (ECM)

regulation profiles...... 96 4.4.4. Full thickness epidermis exhibits dysregulation of transcripts involved in cell proliferation, inflammation, and ECM regulation.....100 4.5. Discussion...... 103 4.6. Materials and methods...... 107

4.7. References...... 108

Chapter 5 Midkine (mk) regulates the wound epidermis-to-AEC transition...... 112 5.1. Preface...... 113 5.2. Abstract...... 113

5.3. Introduction...... 114

5.4. Results...... 114 5.4.1. Midkine is strongly expressed in the basal layers of the

wound epidermis and connective tissue blastemal progenitors..... 114 5.4.2. Chemical inhibition of mk inhibits limb regeneration...... 119

5.4.3. iMDK-treatment impairs wound epidermis maturation...... 121 5.4.4. iMDK-treated limbs display persistant inflammatory profiles and dysregulated wound epidermis gene expression...... 124 5.4.5. Midkine F0 mutants have delayed blastemal outgrowth and

vi impaired wound epidermis maturation...... 128 5.4.6. Overexpression of mk during regeneration leads to

uncontrolled wound epidermis growth...... 131

5.4.7. Sdc-1 is expressed in the wound epidermis...... 132

5.5. Discussion...... 135 5.6. Materials and methods...... 138

5.7. References...... 143

Chapter 6 Conclusions and future perspectives...... 147 6.1. Revealing mechanisms of blastema formation is essential to understanding limb regeneration...... 148 6.2. Early transcriptional programs of blastemal progenitors...... 150 6.3. Inflammation, ECM regulation, and tissue histolysis

are likely regulated by the wound epidermis...... 152

6.4. Blastemal progenitors as early immunomodulators...... 153

6.5. Regulators of early wound epidermis to AEC maturation...... 155 6.6. New frontiers and future perspectives for the field...... 157 6.6.1. Exploring the roles of unannotated genes in salamander regeneration...... 157 6.6.2. Cell-type specific roles in blastema formation...... 158 6.6.3. Genetic regulatory mechanisms of key genes...... 159

6.7. Concluding remarks...... 159

6.8. References...... 160

Appendix...... 165

A.1. Supplemental materials for Chapter 2...... 166

A.2. Supplemental materials for Chapter 3...... 178

A.3. Supplemental materials for Chapter 4...... 180

A.4. Supplemental materials for Chapter 5...... 189

vii List of Figures

Chapter 1

1.1. Introduction to the axolotl and limb regeneration……………………...... 4

1.2. Stages of blastema development………………………………………...... 7

Chapter 2

2.1. Technical validation of FACS-method to purify 4N and 2N fractions from

regenerating stump tissue………………………………………...... 34

2.2. Transcriptional profiles of dividing, non-dividing, and wound epidermal

cells reveal fold change enrichment of well-known early regeneration genes...... 35

2.3. Growth factor signaling pathways are inhibited in regenerating stump tissues...... 37

2.4. Identification of highly enriched transcripts in dividing cells during early regeneration...... 40

Chapter 3

3.1. Il-8 is strongly expressed in blastemal progenitors during early stages of

limb regeneration...... 60

3.2. Il-8 induces recruitment of monocytes and granulocytes in intact limbs...... 63

3.3. Il-8 is sufficient to induce proliferation of bone/perichondrial cells and epidermis...... 65

3.4. Il-8 knockdown results in delayed blastemal outgrowth and regeneration...... 67

3.5. Il-8 knockdown results in defective retention of myeloid cells during the

transition from wound healing into blastema formation...... 69

3.6. Cxcr-1/2 is expressed in subsets of monocytes and granulocytes...... 70

3.7. Early CXCR-1/2 signaling is necessary for limb regeneration...... 71

3.8. CXCR-1/2 inhibition impacts myeloid cell survival during early limb

regeneration...... 73

viii Chapter 4

4.1. Suturing full thickness skin over the amputation plane prevents wound epidermis formation...... 91

4.2. Schematic of transcriptional profiling experiment in the presence and absence

of the wound epidermis...... 92

4.3. PCA analysis of normal and full skin flap sutured limb samples...... 93

4.4. Early dividing cells exhibit minor differences in transcriptional profiles in

the absence of the wound epidermis...... 95

4.5. Inflammation and ECM regulation in stump-derived non-dividing cells are

heavily affected in the absence of the wound epidermis...... 98

4.6. Wound epidermis and full skin flap sutured skin exhibit transcriptional

differences in inflammation, ECM regulation, and cell proliferation...... 106

Chapter 5

5.1. Normalized transcripts per million (TPM) levels of midkine (mk) in regenerating and full skin flap conditions...... 115

5.2. Mk is strongly expressed in the basal layers of the wound epidermis/AEC and connective tissue blastemal progenitors...... 116

5.3. MK protein localization during limb regeneration...... 118

5.4. Chemical inhibition of mk prevents limb regeneration...... 120

5.5. iMDK treatment does not affect early induction of proliferation in limb

regeneration...... 122

5.6. iMDK-treated limbs exhibit an abnormally thin wound epidermis and increased levels of cell death...... 123

5.7. Treatment with iMDK leads to dysregulation of wound epidermis gene

expression and persistent inflammation...... 126

5.8. iMDK-treated limbs exhibit higher levels of monocytes...... 127

5.9. Mk knockout animals exhibit delayed blastema formation...... 129

5.10. Mk mutants exhibit an abnormally thin wound epidermis...... 130

ix

5.11. Mk mutants display a mild increase in the levels of monocytes...... 131

5.12. Overexpression of mk in regenerating limbs leads to uncontrolled growth

of the wound epidermis...... 133

5.13. Sdc-1 is expressed in normal skin and the wound epidermis during limb regeneration...... 134

Appendix

A.1.1. Immune signaling pathways are predominantly activated in the wound

epidermis and non-dividing cells...... 177

A.2.1. Validation of the il-8 overexpression construct and il-8 morpholino

knockdown...... 178

A.2.2. IL-8 morpholino knockdown does not affect blastemal cell or wound

epidermis proliferation...... 179

A.4.1. Mk is not expressed in monocytes...... 193

A.4.2. Validation of custom polyclonal rabbit anti-MK antibody...... 193

A.4.3. PCA analysis of DMSO and iMDK samples...... 194

A.4.4. Mk targeting strategy for mk mutant generation...... 194

A.4.5. Validation of pCAG-MK overexpression construct...... 195

x List of Tables

Appendix

A.1.1. Table of Ingenuity Pathway Analysis (IPA) Z-scores for Immune Signaling Pathways...... 166

A.1.2. Table of Ingenuity Pathway Analysis (IPA) Z-scores for Developmental

Growth Factor and Intracellular Signaling Pathways...... 167

A.1.3. Blastemal-progenitor enriched transcripts...... 168

A.3.1. Top 100 differentially expressed transcripts in stump-derived dividing cells

of full skin flap sutured limbs...... 180

A.3.2. Top 100 differentially expressed transcripts in stump-derived non-dividing

cells of full skin flap sutured limbs...... 183

A.3.3. Top 100 differentially expressed transcripts in epithelial cells of full

skin flap sutured limbs...... 186

A.4.1. Top 100 differentially expressed genes in DMSO/iMDK treatments...... 189

A.4.2. Select enriched pathways in iMDK-treated limbs...... 192

A.4.3. Select enriched pathways in DMSO-treated limbs...... 192

xi Abbreviations AAALAC, Assessment and Accreditation of Laboratory Animal Care AEC, apical epithelial cap AER, apical ectodermal ridge AGSC, Ambystoma genetic stock center cxcr-1/2, C-X-C Motif Chemokine Receptor ½ DMSO, Dimethyl sulfoxide Dpa, Days post-amputation EdU, 5-ethynyl-2'-deoxyuridine ECM, extracellular matrix EMT, Epithelial-mesenchymal transition FACS, Fluorescence-activated cell sorting FSF, Full skin flap FGF, Fibroblast growth factor GFP, Green fluorescent protein GO, Gene Ontology Hpt, hours post-transfection HUGO, Human Gene Nomenclature Committee IACUC, Institutional Animal Care and Use Committee il-8, interleukin-8 IPA, Ingenuity Pathway Analysis iMDK, 3-(2-(4-Fluorobenzyl)imidazo[2,1-b]thiazol-6-yl)-2H-chromen-2-one mk, midkine MO, morpholino NCAE, Naphthol AS-D Chloroacetate NGF, Nerve growth factor NO, Nitric oxide NSE, -Naphthyl Acetate ROS, Reactive oxygen species PBS, Phosphate buffered saline PCA, Principal components analysis ptprz, Protein Tyrosine Phosphatase, Receptor Type Z1 PPAR, Peroxisome proliferator-activated receptor  RXR, Retinoic acid receptor RXR-alpha SB-225002, N-(2-Bromophenyl)-N'-(2-hydroxy-4-nitrophenyl)urea sgRNA, Single guide RNA shh, Sonic hedgehog sdc-1, Syndecan-1 tdT, tdTomato TGF-, Transforming growth factor  TLR, Toll-like receptor TPM, transcripts per million TUNEL, Terminal deoxynucleotidyl transferase dUTP nick end labeling VEGF, Vascular endothelial growth factor WE, Wound epidermis YFP, Yellow fluorescent protein

xii Acknowledgements

I would like to take this opportunity to thank everyone who has supported me throughout the course of my PhD and scientific career. First and foremost, I would like to thank Doug for his continued mentorship and guidance over my entire PhD. I have felt incredibly lucky to have had him as my PhD advisor and could not have asked for a more supportive mentor. Throughout the course of my PhD, Doug has given me the freedom to pursue all of my experimental ideas, even if they did not ultimately pan into successful projects. He has also taught me to think critically about designing and interpreting results of experiments. I know that throughout the rest of my scientific career, I will always be asking myself the question: “How can I be wrong? What are alternative explanations for these results?” With his guidance over the years, I know that I have grown into a more independent and critical scientist and I can only hope to aspire to one day become as great of a scientist as him. Science aside, Doug is one of the most compassionate people that I know, and always cares about the personal well-being of everyone in the lab. I will definitely miss having regular conversations with him not just about science and my career, but also about life in general.

I would also like to thank the members of my committee. Cliff has been a mentor to me since my undergraduate years in his lab, and has been a great advisor in the fields of limb development and regeneration. Cassandra has also provided valuable career advice and support throughout the course of my projects. Finally, Jessica has been an important scientific mentor to me since my years as an MIT undergrad in Cliff’s lab.

Working closely with her as an undergrad on axolotls, she taught me a lot about the limb regeneration field. Over the years, she has continued to provide valuable career guidance

xiii and scientific input on my projects over my PhD and her enthusiasm about salamander limb regeneration and science in general is always energizing.

Next, I was incredibly lucky to have the opportunity to work with a very talented undergraduate, Clara Baselga-Garriga, over the last two and a half years. Clara always comes to lab with a smile and ready to work. She has basically been my partner in crime on the midkine project in particular and I cannot begin to describe what a pleasure it has been to mentor her during her time in the Melton lab. She has always reminded me that science is more fun when working with others. Bonding over late nights and early mornings injecting eggs to make axolotl mutants, getting excited over results, having intense experimental discussions, and also just talking about life in general. I’ll really miss working with her. To see her grow scientifically over the time we have spent together has been really rewarding and I am proud of all of she has achieved. I know she will be a famous doctor/writer some day! I look forward to the day when I can ask for her autograph.

The Melton lab has also been an incredible environment to work in over the last five years and I owe a lot of the successes in my PhD to the support from all of the lab members. Even though I am the only person who works with axolotls, I have always had immense support from all of the many past and present members of the lab and I’d like to thank them all. I would like to specially thank several lab mates who helped me a lot through my PhD. José has been my main go-to person for scientific insight, questions, and funny anecdotes. Not only is he super knowledgeable about classical methods in biology, but his daily jesting always brightens my days in the lab. Edwin has been an amazing bay mate over the years and his laugh ironically always makes me laugh as well.

I always enjoy having scientific and life discussions with him. Jeff in particular has also

xiv been a great friend throughout graduate school and one of the most thorough scientists that I know. Coffee breaks with both Jeff and Edwin over the years have been one of my favorite past times. Ronny and I will always be the first to flock to free food as if we have a special sense of smell if it’s free. I am also looking forward to our future matchups and trades in Fantasy Football. Joking aside, Ronny has also been an amazing influence on me as a scientist, and the high standards that he holds himself to as a scientist are inspirational. Nadav is one of the most critical scientists I know in the lab, and he has always provided good input on my projects. In addition, I enjoy hearing about all of the fun facts and anecdotes he has to share. Jenny has also been an amazing help when it comes to literally anything in the lab whether its ordering or animal-related. Her smile and laugh is contagious. Elad has also been a great person to talk to about science, life and our careers. Some past members of the lab who supported me throughout my PhD who

I’d also like to thank are Quinn Peterson, George Kenty, Danny Ben-Zvi, Dani Swain, Chi-

Yang Chen, and David Gonzalez. I am also very grateful to those who have helped me with setting up the axolotl facility and also caring for the axolotls over the years: Isaac,

Lauren, Eny, Mimi, Sergio, Quang, and Joe Vaughn.

Aside from my lab, I’d like to thank my friends who have been a main block of support throughout my PhD. In particular, I’d like to thank my graduate classmates in the

MCO program, who have made this an unforgettable experience. In particular, Shristi,

Denise, Haneui, Vika, Jenelle, and Brenda, for their amazing friendship and support over the years. So many great memories. My long-time roommates, Denise and Jocelyn, have also been my support beams for my PhD. Grad school would not have been nearly as fun without my roomies! I’d like to also thank my best friend in college, Sharon, for her support

xv and friendship over the years. Lastly, I am grateful to previous members of the Arlotta lab including Ryoji, for his friendship and guidance in the axolotl field as well as my scientific career over the years, and Dennis, for a fun first few years working on axolotls together in the lab.

Importantly, I’d like to thank my mom, dad, and brother, Victor, who have provided me with every opportunity since I was young and have supported me on this amazing journey. From driving me to Kentucky to work on axolotls over the summer to now completing my PhD. Words cannot describe how much their support and love means.

They’ve always inspired me to pursue my dreams and I really owe all of my successes to them. I hope that I have made them proud.

Finally, I’d like to thank my amazing boyfriend Joe, who has been my superman throughout my PhD. Not only has he been an inspiration scientifically to me, but I feel as though we have grown together over the course of our PhDs. Whenever times were tough, he was my rock and kept me grounded. I could not have asked for a better person to share this success with.

xvi

For my mom, dad, and Victor

xvii

“If I set out to prove something I am no real scientist-- I have to learn to follow where the facts lead me-- I have to learn to whip my prejudices...” ― Lazzaro Spallanzani

xviii

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Chapter 1

Introduction

Since it was first reported in the 18th century by Lazzaro Spallanzani (Spallanzani,

1768), salamander limb regeneration has fascinated scientists for centuries. Why humans lack the ability to similarly accomplish such a complex biological feat despite the striking anatomical resemblance between human and salamander limbs remains a longstanding enigma. While mammals traditionally respond to limb amputation with a scarring response, salamanders respond by forming a new cellular structure distal to the amputation plane known as the blastema. How this structure forms lies at the heart of the limb regeneration field and elucidating the molecular mechanisms underlying the initiation of blastema formation may direct our efforts to potentially stimulate regenerative potential in humans.

1.1. Introduction to the Mexican axolotl

In the lab, we have utilized the Mexican axolotl (Ambystoma mexicanum) as the model organism to study salamander limb regeneration. This diploid urodele amphibian species is endemic to the canals of Lake Xochimilco in the outskirts of the modern-day

Mexico City. Once a staple in the Aztec diet hundreds of years ago, axolotls are now mostly utilized for research purposes in developmental and regenerative biology. While most other salamander species undergo metamorphosis and become terrestrial, axolotls are naturally neotenic, remaining in a larval aquatic state throughout their lifespan and retaining juvenile bodily features including gills. However, axolotls retain the ability to undergo metamorphosis experimentally if injected or immersed with the hormone thyroxine (Page et al., 2008; Page et al., 2009; Khattak et al., 2014a) and others have applied these methods to study differences in regenerative capabilities pre- versus post- metamorphosis (Monaghan et al., 2014).

The most commonly used axolotl strain is the white leucistic strain (d/d). White axolotls contain an unknown recessive mutation that disrupts pigment cell migration and development during embryogenesis, which leads to a pale pink appearance (Figure 1.1A)

(Woodcock et al., 2017). The lack of pigment cells allows for reduced autofluorescence and easy visualization in live imaging or immunofluorescence studies. In addition to white mutants, wild-type pigmented (dark green appearance) and albino (light yellow appearance) axolotls also exist, however, these strains still harbor various pigment- producing cell types (known as melanophores, xanthophores, and/or iridophores) that contribute to autofluorescence throughout the animal, making white axolotls the preferred strain in research studies.

2 Over the last couple of decades, the axolotl has become the most widely utilized salamander for research studies due to increased genetic and technical tractability

(Khattak et al., 2014a) compared to other salamander species. Axolotls breed year-round with ease in the laboratory setting and generate large clutches of fertilized eggs (typically at least 300 eggs). In fact, the major populations of axolotls in the world are all bred in captivity, as their numbers in the wild are quickly declining. Moreover, the development and optimization of germline transgenesis methods in the axolotl over the last decade via

CRISPR (Flowers et al., 2014; Fei et al., 2018), Tol2, and I-sceI meganuclease-mediated transgenesis (Khattak et al., 2014a), as well as inducible genetic systems (Whited et al.,

2012; Khattak et al., 2014b) has made it an attractive model for research purposes.

Coupled with the recent release of the axolotl genome (Nowoshilow et al., 2018; Smith et al., 2019), these methods allow for genetic lineage tracing and targeted knock-out/knock- in studies to functionally test molecular candidates. Moreover, the relatively slow embryonic development (~6 hours at the 1-cell stage) allows for high penetrance of mutations via injection of targeted Cas9-ribonucleoprotein (Cas9-RNP) complexes into fertilized eggs (Fei et al., 2018), allowing phenotyping to occur in the F0 generation and offsetting the challenges associated with their long generation time (10-12 months).

Various viral infection and plasmid electroporation methods have also been developed for use in molecular gain-of-function studies (Khattak et al., 2013; Whited et al., 2013;

Oliveira et al., 2018). Altogether, these techniques have opened the doors for detailed molecular genetic studies in the axolotl.

3

Figure 1.1. Introduction to the axolotl and limb regeneration. (A) An image of an adult white leucistic (d/d) axolotl adapted from Learn, 2019. (B) A diagram depicting the anatomy of an axolotl forelimb. The solid lines denote the plane of amputation used for all experiments within this dissertation. (C) Stages of limb regeneration depicting the time scale of wound healing, blastema formation, and re- differentiation stages. This figure was adapted from McCusker et al. (2015).

4 1.2. Structure of the axolotl limb

Axolotls are tetrapod vertebrates with limbs that resemble those of humans anatomically, containing similar skeletal structures and tissues (e.g. muscle, bone, cartilage, connective tissues etc.). Fore- and hindlimbs can be divided into three regions proximodistally based on the residing skeletal structures: the stylopod, zeugopod, and autopod (Figure 1.1B). The stylopod encompasses the upper-limb long bones, either the humerus or femur in the fore- or hindlimb, respectively. The zeugopod contains the two long bones in the forearm or lower leg skeletal structures, known as the radius/ulna or tibia/fibula in the fore- or hindlimb, respectively. Finally, the autopod is the most distal segment and contains all of the bones that construct the hands or feet. These include the carpals, metacarpals, and phalanges in the forelimb and tarsals, metatarsals, and phalanges in the hindlimb. Axolotl forelimbs and hindlimbs have four and five digits, respectively. In the salamander limb regeneration field, researchers generally perform amputations at either the mid-humerus/femur (known as a proximal amputation) or the mid-radius/ulna or -tibia/fibula (known as a distal amputation). All of the amputations performed in the studies presented within this dissertation were distal amputations.

1.3. Stages of limb regeneration

Limb regeneration can be divided into three phases: wound healing, blastema formation, and re-differentiation (Figure 1.1C, Figure 1.2) (McCusker et al., 2015). Briefly, wound healing occurs during the first 3 to 7 days post-amputation (dpa). Immediately post-amputation, epithelial cells quickly migrate over the exposed amputation plane to form a thin wound epidermis in as little as 12 hours post-amputation. Formation of the

5 wound epidermis is followed by immediate inflammatory responses mediated primarily by the innate immune system in response to injury and the beginnings of extracellular matrix degradation. Following wound healing, progenitors derived from many different tissues in the limb (e.g. muscle, connective tissues etc.) (Kragl et al., 2009) re-enter the cell cycle, migrate to the amputation plane, and proliferate locally to form the blastema, the hallmark structure of limb regeneration. Blastema formation is tightly coupled with histolysis of tissues at the amputation plane (e.g. myofibers and bone/cartilage) and deposition of pro- regenerative extracellular matrix molecules including tenascin (Seifert et al., 2012).

Concomitantly, the wound epidermis thickens to form an epithelial cap structure called the apical epithelial cap (AEC). This thickening is primarily attributed to proliferation of epithelial cells in the periphery and subsequent migration towards the amputation plane

(Repesh and Oberpriller, 1978). The AEC is necessary for successful limb regeneration and secretes mitogens that maintain blastemal cell proliferation (Thornton, 1957; Boilly and Albert, 1990). Blastema development occurs over 1-4 weeks depending on the age of the animal and the stage of regeneration is defined by the size of the blastema (e.g. early, medium, and late bud blastema stages). Over the next several weeks, the blastema will gradually re-differentiates and re-patterns into a new limb. Complete regeneration takes approximately 4 weeks in younger juvenile animals and 10 weeks in adults. The work presented in this dissertation is primarily focused on the earliest stages of regeneration during the transition from wound healing into blastema formation. Blastema formation takes approximately 2 weeks to occur in most of the animals utilized in these studies.

6

Figure 1.2. Stages of blastema development. Blastema formation occurs through many phases. The first phase is wound healing. Immediately post amputation, epithelial cells migrate quickly over the exposed amputation plane to form a thin wound epidermis. Immune cells subsequently invade the injured area in order to respond to pathogen-associated and/or tissue damage-associated cues. These characteristic immediate inflammatory responses are also coupled with the beginnings of extracellular matrix (ECM) degradation and tissue histolysis. During the later stages of wound healing, progenitors derived from many different tissues (collectively referred to as blastemal progenitors) begin to re-enter the cell cycle and migrate towards the amputation plane, while wound healing begins to resolve and immune cells efflux the damaged area. This is followed by localized proliferation at the distal amputation plane, which gives rise to the blastema. As the blastema forms, the wound epidermis continually thickens to form the AEC, which is necessary for regeneration. AEC, apical epithelial cap.

7 1.4. The heterogeneous cellular origins of the blastema

Much of the complexity of limb regeneration lies in the cellular heterogeneity within the blastema. Researchers previously thought that the blastema was comprised of pluripotent stem cells based on their homogenous appearance; however, nuclear transfer experiments of blastema cell nuclei from regenerating axolotl limbs into enucleated X. laevis eggs did not give rise to cloned animals and provided support against this theory

(Dasgupta, 1970). Decades later, researchers demonstrated that the blastema is instead comprised of tissue-lineage restricted progenitors by labeling tissues within the axolotl limb via different regional embryonic transplantation experiments between GFP+ and

GFP- axolotl embryos (Kragl et al., 2009). For example, by transplanting presomitic mesoderm from GFP+ to GFP- embryos, they achieved GFP labeling of the muscle in the axolotl limb. Upon amputation, they discovered that only the muscle in the regenerated limb was GFP+, suggesting that muscle only regenerates itself and no other tissues. Using this labeling technique, they showed the same lineage restriction occurred for other tissues including cartilage tissue, epidermis, and Schwann cells. The only tissue that displayed a higher level of plasticity was the dermis, which contributed to regenerated cartilage and tendon tissue as well.

More recent studies have elucidated the specific origins of muscle-derived blastema cells. Sandoval-Guzmán et al. (2014) utilized genetic lineage tracing of mature muscle fibers to demonstrate different cellular origins for regenerated muscle between two salamander species, the axolotl and the Eastern spotted newt (Notophthalmus viridescens). By genetically labeling mature muscle fibers with YFP in both species, they observed that YFP+ cells contributed to regenerated muscle in newts, but not in the

8 axolotl. These results interestingly suggested different cellular sources and mechanisms for muscle regeneration in newts and axolotls: dedifferentiation of mature myofibers in newts versus activation of the resident muscle stem cell population, known as satellite cells, in axolotls. By performing embryonic somatic mesoderm transplantations as in Kragl et al. (2009), they achieved GFP labeling of muscle tissue including both the mature multinucleated fibers and satellite cells. Examination of the blastema of regenerating limbs from GFP labeled animals revealed that all of the GFP+ cells in the blastema expressed pax7, a satellite cell marker (von Maltzahn et al., 2013), suggesting that axolotls regenerate muscle via activation of satellite cells rather than dedifferentiation of mature muscle myofibers. Lineage tracing of pax7+ cells during axolotl limb regeneration also confirmed these conclusions (Fei et al., 2017). These studies collectively determined that satellite cells are the source of regenerated muscle in axolotl limb regeneration.

Aside from muscle, the specific origins of blastema cells derived from other tissues remain elusive, due to a lack of reliable markers for different cell types within the axolotl limb. However, recent single cell studies have begun to shed light on these questions by assessing the cellular heterogeneity within the mature and regenerating limb (Gerber et al., 2018; Leigh et al., 2018). Unbiased single cell sequencing and computational pseudotime lineage reconstruction of cell state trajectories from wound healing, early, and mid-bud stage regenerates suggest that most cell-types exhibit distinct lineage trajectories. However, synovial fibroblasts, joint, and cartilage cells appear to arise from a shared progenitor population (Leigh et al., 2018). Additionally, Gerber et al. (2018) utilized a driver element of the pan-connective tissue marker prrx-1 to label all connective tissues (Prrx1:Cre-ER;Caggs:lp-Cherry) including tendons, cartilage, and dermal

9 fibroblasts. Single cell analysis and lineage reconstruction of connective tissue progenitors revealed that cells derived from different types of connective tissues converge on a similar molecular blastemal cell identity as the blastema forms, and diverge upon re- differentiation at the end of regeneration. Brainbow clonal lineage labeling of connective tissue-derived blastemal cells also suggested that this convergent molecular blastemal cell state is multipotent and capable of giving rise to both skeletal and non-skeletal connective tissues. Most notably, these studies provide the field with reliable markers for cell types present within the axolotl limb to more specifically address questions about the exact cellular sources of the blastema via genetic experimentation in future studies.

1.5. Essential components of successful limb regeneration

Classical studies have illustrated that salamander limb regeneration requires an injury, the presence of nerves, and the wound epidermis (Tassava and Mescher, 1975).

Previous studies have suggested that each plays a different role in the process: an injury is necessary to initiate tissue histolysis and cell cycle re-entry, while nerves and the wound epidermis are important for the early and later proliferative capacity of blastemal cells, respectively (Mescher and Tassava, 1975; Tassava and Loyd, 1977). The studies demonstrating the known roles of each of these components are reviewed below.

1.5.1 Nerve dependence

The nerve dependence of limb blastema formation has been well-known for centuries (Todd, 1823). Denervation prior to blastema formation hinders regeneration, whereas denervation post-blastema formation results in normal regeneration,

10 emphasizing the important early influence of nerves on successful regeneration (Singer,

1952; Farkas and Monaghan, 2017). While denervated limbs post-amputation exhibit many of the same early responses as that of innervated limbs including tissue histolysis and wound epidermis formation (Schotte and Butler, 1941), they display low levels of cellular proliferation, which is integral for blastema formation. Moreover, microarray analyses revealed lower levels of transcripts involved in cell proliferation, axonal regeneration, and epidermal maturation in denervated limbs (Monaghan et al., 2009). The importance of nerves on blastema formation led researchers to believe that nerves secreted a neurotrophic factor crucial for the initial stages of regeneration (Singer, 1952).

To date, some important nerve-derived factors have been identified including Neuregulin-

1 (NRG-1) and Newt Anterior Gradient (NAG) protein (Kumar et al., 2007; Farkas et al.,

2016). Supplementation of NRG-1 or NAG is sufficient to rescue regeneration in denervated limbs of axolotls or newts, respectively, demonstrating the importance of these factors. Moreover, this nerve dependence appears to be widely conserved in the regeneration of many other organ systems in other species as well (Buckley et al., 2012;

Simoes et al., 2014; Mahmoud et al., 2015).

1.5.2. Wound epidermis/AEC

The wound epidermis is a transient epithelial structure that is necessary for regeneration (Godlewski, 1928; Goss, 1956; Thornton, 1957; Mescher, 1976). The wound epidermis is distinct from normal skin. Normal non-regenerating skin is comprised of two layers, the epidermis and dermis (Seifert et al., 2012), and similar to mammalian skin, the epidermal cells follow an outward differentiation trajectory (Leigh et al., 2018). These two

11 layers are separated by a basement membrane layer called the basal lamina, which acts as a collagen heavy physical barrier between the skin and the underlying tissues. In contrast, the wound epidermis is comprised of only an epidermal layer and lacks a basal lamina. The absence of the basal lamina is thought to allow for signaling between the wound epidermis and underlying limb tissues, which is necessary for blastema formation

(Mescher, 1976). During wound healing and the initial stages of blastema formation, the wound epidermis is a thin epithelium, only 1-4 cell layers thick (Salpeter and Singer, 1960;

Singer and Saltpeter, 1961; Campbell and Crews, 2008), formed by proliferating keratinocytes that migrate radially from the peripheral normal skin to cover the amputation site. As the blastema forms, the wound epidermis matures into the AEC and thickens considerably via continued cell proliferation and migration of peripheral keratinocytes.

Considering the major differences in the cellular environment between early and late stages of limb regeneration, it is likely that the early wound epidermis and the AEC have discrete functional roles. For instance, the early wound epidermis is exposed to highly pro-inflammatory conditions associated with early injury-associated immune responses, in contrast to the AEC, which is not.

Studies during later stages of regeneration post-blastema formation have demonstrated that the AEC functionally resembles the apical ectodermal ridge (AER) during limb development by playing similar roles in maintaining cell proliferation and limb patterning (Boilly and Albert, 1990; Christensen and Tassava, 2000; Han et al., 2001).

Continuous removal of the AEC from the blastema inhibits regeneration (Thornton, 1957) and co-culturing blastemal cells with protein extract derived from the AEC stimulates proliferation (Boilly and Albert, 1990). In addition, the AEC expresses important mitogens

12 known to play a role in limb development including FGF-8 (Han et al., 2001; Christensen et al., 2002), collectively suggesting that the AEC mirrors the developmental roles of the

AER.

While the roles of the AEC have been more defined, the specific functions of the early wound epidermis are less well known. Preventing the wound epidermis from forming entirely either by suturing full thickness skin (containing a basal lamina) over the amputation surface immediately (Godlewski, 1928; Mescher, 1976) or via insertion of the amputated limb directly into the body cavity (Goss, 1956) inhibits regeneration, demonstrating the necessity of the early wound epidermis. Interestingly, normal induction of cell cycle re-entry and some tissue histolysis within non-epidermal stump tissues still occurs in the absence of the wound epidermis, suggesting that it may not play a major role in these processes. Nevertheless, signaling molecules secreted from the early wound epidermis including axolotl MARCKS-like protein (axMLP) (Sugiura et al., 2016) have been shown to be sufficient to induce cell cycle re-entry in various tissues within axolotl limbs, indicating it likely still participates in these processes. Furthermore, the early wound epidermis expresses many matrix metalloproteinases (Yang et al., 1999; Kato et al.,

2003), suggesting it may direct the formation of the pro-regenerative extracellular niche.

The maturation of the early wound epidermis to the AEC is essential for successful limb regeneration, yet little is known about factors that are important for the maturation from the early wound epidermis to the AEC. The early wound epidermis becomes heavily innervated during the first week of regeneration and classical studies suggested that early innervation was key for the functional maturation to the AEC (Singer, 1949). However, the demonstration of successful limb regeneration in aneurogenic larvae (Thornton and

13 Steen, 1962) and in limbs in which the innervation of the wound epidermis was prevented

(Sidman and Singer, 1961) suggested that direct innervation may not be important.

Others have suggested that contact between epithelial cells originating from opposing positions within the limb is key for wound epidermis function and maturation, coined the positional discontinuity model (Campbell and Crews, 2008). In other words, contact between epithelial cells from the dorsal and ventral side, or anterior and posterior side of the limb, is essential for forming a wound epidermis capable of regenerating a limb.

However, contact between epithelial cells from the same position in the limb (ex. anterior with anterior) will not form a regenerative wound epidermis. This hypothesis was supported by work that demonstrated that an accessory limb will form in a non- regenerating limb simply by juxtaposing a generic wound epidermis (from just a skin wound) on the anterior side of the limb with a posterior piece of skin from the contralateral limb and deviating a nerve to the wound epidermis. This accessory limb will not form without the skin graft (Endo et al., 2004), suggesting that positional discontinuity is important for the generation of a wound epidermis capable of giving rise to a limb.

Understanding the molecular regulation downstream of this discontinuity could help us gain insights into wound epidermis maturation in future studies.

1.5.3. Injury

Interestingly, the studies delineated above altogether demonstrate that cell cycle re-entry and tissue dedifferentiation still occur to some extent regardless of the absence of nerves or wound epidermis (Tassava and Mescher, 1975), suggesting that other more immediate processes associated with injury are important for initiating regeneration.

14 One such process is hemostasis. Limb amputation is followed by a quick hemostatic response in which the blood clots in under a minute (Repesh and Oberpriller,

1978). This is a much quicker response than in mammals including humans, which takes several minutes. Previous in vitro studies determined that newt myotubes re-entered the cell cycle when cultured in the presence of fetal bovine serum. Co-culturing myotubes in the presence of a variety of known growth factors such as TGF- and bFGF did not induce cell cycle re-entry (Tanaka et al., 1997). However, activity-guided fractionation of serum identified two peaks capable of inducing cell cycle re-entry in newt myotubes: thrombin and an unidentified thrombin-cleaved product (Tanaka and Brockes, 1998; Tanaka et al.,

1999). Thrombin is an important enzyme that catalyzes the formation of fibrin from fibrinogen, a necessary step to forming a blood clot. While the cell cycle re-entry activity of thrombin was serum dependent, the thrombin-cleaved product retained activity without serum and induced cell cycle re-entry of multinucleated newt myotubes. Furthermore, murine myotubes did not re-enter the cell cycle in contrast to their newt counterparts

(Tanaka et al., 1997), highlighting potentially fundamental differences between mammalian and newt responses to injury. To date, researchers have identified two secreted molecules capable of initiating cell cycle re-entry in the axolotl and newt:

MARCKS-like protein (axMLP) and serum protease cleaved BMP-4/7, respectively

(Sugiura et al., 2016; Wagner et al., 2017).

Others have investigated different aspects of immediate injury-associated processes in various appendage regeneration models including the production of reactive oxygen species (ROS) and the dysregulation of natural voltage gradients. Previous studies have identified the importance of tightly controlled amputation-induced ROS

15 production during the first 24-48 hours post-injury in the zebrafish fin and Xenopus tail post-amputation (Gauron et al., 2013; Love et al., 2013). These studies demonstrate that

ROS production is essential for proper modulation of important downstream signaling pathways (ex. Wnt) and cellular processes (ex. apoptosis, leukocyte recruitment). In addition, researchers have investigated the changes in bioelectric gradients following amputation in appendage regeneration (McLaughlin and Levin, 2018). Amputation considerably disrupts the voltage gradient throughout the limb and many studies perturbing these natural gradients via channel blockers or gradient reversal severely impaired newt limb regeneration (Borgens, 1982; Jenkins et al., 1996). Altogether, these studies suggest that more immediate processes associated directly with amputation itself play important roles in initiating limb regeneration.

1.6. Immunological influences on regeneration

Recent decades have elucidated the importance of the immune system on successful regenerative outcomes across many organ and appendage model systems

(Godwin et al., 2017b). Any injury elicits an immediate immunological response to defend the organism against potential infectious pathogens that may have entered the body and to clear cellular debris caused from the damage (Tang et al., 2018). The wound healing response is common to all vertebrates. However, while mammals typically produce a scar after wound healing, highly regenerative vertebrates including the salamander exhibit perfect regeneration. Thus, identifying potential molecular and cellular differences in the wound healing response between highly- and less- regenerative species may help direct pro-regenerative outcomes in humans.

16 Wound healing is mediated primarily by the innate immune system, which is comprised of myeloid cells that circulate throughout the body. Myeloid cells encompass several different phagocytic cell types including macrophages, which are derived from precursors known as monocytes, and neutrophils, basophils, and eosinophils, which are collectively known as granulocytes (Chaplin, 2010). Myeloid cells are most well-known for their exceptional ability to phagocytose dead cells and debris at the wound. In addition to the innate immune system, lymphoid cells of the adaptive immune system (T- and B- cells) also home to the site of injury and mediate specific responses against recognized antigens as well as the long-term humoral response for future encounters with new pathogens. While classical immunological roles have been well delineated for both innate and adaptive immune cells during injury and infection, myeloid cells, T- and B- cells have been more recently been shown to be important paracrine regulators in development, regeneration, and tumorigenesis (Aurora and Olson, 2014; Naik et al., 2018).

Furthermore, many of these cell types have also been shown to normally reside in tissues and mediate homeostatic behaviors of tissues (Naik et al., 2018).

While the axolotl immune system contains all of the myeloid and lymphoid cell- types present in mammalian species (Lopez et al., 2014), the work of others suggests that their adaptive immune response is heavily attenuated compared to other amphibians including Xenopus and fish species (Ching and Wedgwood, 1967). Some have hypothesized that the complexity of the immune system is inversely correlated with regenerative capacity (Mescher and Neff, 2005; Mescher et al., 2013), as amphibians including Xenopus can regenerate their hindlimbs and tails pre-metamorphosis, but lose their full regenerative capacity post-metamorphosis, once their immune system has

17 matured. Similarly, mammals are capable of regenerating dermal injuries during fetal stages of development, but also lose this regenerative capacity and form a scar instead post-natally (Aurora and Olson, 2014). In addition, suppressing the immune system with immunosuppressants or immune cell depletion is sufficient to stimulate Xenopus tail regeneration in a non-regenerating developmental period, the refractory period

(Fukazawa et al., 2009), suggesting the immune system is an important mediator of regeneration. Yet, organisms with highly evolved immune systems including zebrafish are capable of regenerating many organs and appendages (Trede et al., 2004), suggesting an added layer of regulatory complexity that may be species-dependent.

Nevertheless, cells from both the innate and adaptive immune system are present in early regenerating limb tissues and continue to reside within the blastema (Godwin et al., 2013; Leigh et al., 2018), suggesting active roles in regeneration. Furthermore, macrophages are necessary for both axolotl limb and heart regeneration (Godwin et al.,

2013; Godwin et al., 2017a) playing roles including induction of the expression of limb regeneration-associated genes, deposition of pro-regenerative ECM components, and clearance of senescent cells (Yun et al., 2015). Interestingly, macrophage-depleted regenerating axolotl limbs and hearts exhibit characteristics of scarring, suggesting that macrophages are key players in tipping the balance between a non-regenerative and regenerative outcome. Given the importance of different immune cells in the regeneration of other systems, it is likely that other innate and adaptive cell types including neutrophils and T-cells also play regenerative roles that have yet to be elucidated in the axolotl. In all, it is clear that differences exist between the behavior of immune cells in regenerative

18 and non-regenerative systems. Whether these behaviors are driven by cell autonomous or non-cell autonomous mechanisms remains to be discovered.

1.7. Topics addressed in this dissertation

My doctoral thesis broadly focused on characterizing the early transcriptional programs of dividing progenitors and the surrounding tissues during the cell cycle re-entry stages of axolotl limb regeneration in order to identify new regulators of blastema formation. To this end, we developed and validated a method to use DAPI cell cycle analysis to isolate dividing cells in regenerating stump tissues (enriching for blastemal progenitors) in Chapter 2. We utilized this method to isolate and transcriptionally profile three distinct populations during early stages of regeneration: stump-derived dividing cells, stump-derived non-dividing cells, and the wound epidermis. These data allowed us to identify and differentiate the gene expression programs active in progenitors that likely give rise to the blastema from those of the surrounding tissues (non-dividing cells and the wound epidermis) at the beginning stages of cell cycle re-entry. Furthermore, we provided the field with a list of blastemal progenitor-enriched transcripts that may play important roles in early blastemal cell biology. Exploration of the functional relevance of one blastemal progenitor-enriched candidate, a well-known inflammatory cytokine interleukin-

8 (il-8), in Chapter 3 revealed a newly identified role of blastemal progenitors as immunomodulatory players in the early stages of regeneration. Finally, we were interested in exploring the functional role of the wound epidermis during early stages of regeneration. To this end, we expanded upon the regenerating transcriptional dataset from Chapter 2 and added an additional condition in which we prevented wound

19 epidermis formation in Chapter 4. These data allowed us to ask how the transcriptional programs of blastemal progenitors and surrounding tissues change when wound epidermis formation is prevented. Finally, in Chapter 5, we functionally examined the role of one candidate, a pleiotropic growth factor cytokine midkine (mk), that was down- regulated in the absence of the wound epidermis. Functional gain- and loss-of-function experiments collectively suggest that mk is likely a regulator of wound epidermis maturation to the AEC.

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26 Thornton, C. S. and Steen, T. P. (1962). Eccentric blastema formation in aneurogenic limbs of Ambystoma larvae following epidermal cap deviation. Dev Biol 5, 328-343.

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27

Chapter 2

Transcriptional programs during early stages of axolotl limb regeneration

2.1. Preface

The work presented in this chapter was modified and adapted to fit the guidelines of this dissertation from the following publication: Tsai S.L., Baselga-Garriga C., and

Melton D.A. (2019). Blastemal progenitors modulate immune signaling during early limb regeneration. Development:146(1). The conceptualization and experimental design was performed in collaboration with Doug. I performed all of the experiments and the data analysis.

2.2. Abstract

Blastema formation, a hallmark of limb regeneration, requires proliferation and migration of progenitors to the amputation plane. While blastema formation has been well- described, the transcriptional programs that drive blastemal progenitors remain unknown.

Here, we devised a method to utilize the cell cycle as a marker to enrich for early blastemal progenitors. To investigate the gene expression signatures within blastemal progenitors apart from the surrounding tissues, we transcriptionally profiled dividing and non-dividing cells in regenerating stump tissues, as well as the wound epidermis, during early stages of axolotl limb regeneration. Surprisingly, pathway analysis of differentially expressed transcripts from each population indicated the suppression of several core developmental signaling pathways including FGF signaling in early regenerating stump tissues. Furthermore, the transcriptional profiles of early dividing cells suggest selective establishment of an autocrine TGF- signaling network and roles in shaping the regenerative niche. Finally, our analysis reveals a candidate list of blastemal progenitor-

29 enriched transcripts that may provide new insights into blastemal cell biology during the initiation of regeneration.

2.3. Introduction

Salamanders, including newts and axolotls, possess the ability to fully regenerate their limbs throughout their lifespan. This regenerative capacity requires the formation of a transient cellular structure distal to the amputation plane known as the blastema which is comprised of progenitors derived from multiple different tissues (Kragl et al., 2009;

Sandoval-Guzman et al., 2014; Currie et al., 2016). While salamanders can perform this process regularly, mammals are incapable of forming a blastema that can give rise to an entire limb. Understanding the mechanisms underlying the initiation of blastema formation may provide important insights into unlocking human regenerative potential.

Blastema formation requires coordinated proliferation and migration of progenitors derived from muscle, bone, dermal fibroblasts, connective tissue, and other tissues (Kragl et al., 2009). Researchers have shown that while the blastema itself is transcriptionally similar to the limb bud during development, regenerating limbs exhibit unique transcriptional profiles during early stages of regeneration before the blastema has formed (Knapp et al., 2013). Elucidating these signals, as well as their specific activities, is crucial for understanding why and how salamanders respond to amputations with blastema formation. Several important signaling molecules and pathways have already been identified during early stages of appendage regeneration. For instance, axolotl

MARCKS-like protein (axMLP) is a secreted molecule from the wound epidermis required for the induction of tail regeneration and capable of inducing cell proliferation in intact

30 limbs (Sugiura et al., 2016). Moreover, several core developmental signaling pathways including FGF and Wnt are required for appendage regeneration in axolotls and other species (McCusker et al., 2015; Haas and Whited, 2017; Stocum, 2017).

Several studies have investigated bulk transcriptional changes during different stages of axolotl limb regeneration and successfully identified genes that may play important roles in blastema formation and maintenance (Monaghan et al., 2009; Campbell et al., 2011; Knapp et al., 2013; Stewart et al., 2013; Wu et al., 2013; Voss et al., 2015;

Bryant et al., 2017; Gerber et al., 2018). More recently, single cell analysis specifically in the connective tissue lineage has elucidated the molecular transitional states during dedifferentiation of mature connective tissue to a progenitor state during limb regeneration (Gerber et al., 2018). Additionally, unbiased single cell sequencing in regenerating limbs has provided the field with novel markers for different cell types within the limb to begin to examine regeneration at finer resolution (Leigh et al., 2018).

In the present study, we sought to investigate the distinct transcriptional programs active within blastemal progenitors irrespective of lineage, as well as the surrounding tissues, to better understand the genetic programs and signaling interactions that govern the initiation of blastema formation. We reasoned that blastemal progenitors would be among early proliferating cells in stump tissues following amputation. Therefore, to enrich for blastemal progenitors, we transcriptionally profiled dividing cells from regenerating stump tissues during early stages of axolotl limb regeneration. This approach allowed us to examine the transcriptional signature of early dividing cells, inclusive of blastemal progenitors, and identify potential novel modulators of early blastemal cell induction and maintenance. We additionally profiled non-dividing cells in regenerating stump tissues

31 and the wound epidermis at similar stages for comparison. Pathway analysis of the discrete transcriptional programs of all three subpopulations revealed differential patterns of signaling pathway activation/inhibition and unexpectedly uncovered the repression of several core developmental signaling pathways throughout regenerating stump tissues.

2.4. Results

2.4.1. Transcriptional profiling of dividing cells during the initiation of limb regeneration

Cell cycle re-entry is an integral event for the initiation of limb regeneration prior to blastema formation. We therefore reasoned that we would be able to exploit differences in the cell cycle, such as DNA content, as a means of enriching for blastemal progenitors during early stages of regeneration. We developed and optimized a protocol to perform

DAPI staining in conjunction with FACS to separate 4N and 2N cells in the regenerating stump during early regeneration (Figure 2.1). Intact limb tissue is referred to as the 0 dpa timepoint which was collected to serve as a non-regenerating control. Utilizing this method, we transcriptionally profiled three total cellular populations at 0, 4, and 5 days post-amputation (dpa): stump-derived dividing (4N) and non-dividing (2N) cells, as well as the whole wound epidermis (Figure 2.3A).

Principal component analysis (PCA) of the transcriptional profiles of all samples revealed four clusters representing tissue type (stump-derived or epidermis) and regeneration status (Figure 2.2A). Differential expression analysis of transcripts between the 4N (proliferating) and 2N (non-dividing) fractions at all three timepoints showed the expected enrichment of cell cycle gene expression in the proliferating fraction (Figure

2.2B). Moreover, known blastemal cell markers, including prrx-1 (Satoh et al., 2011;

32 Gerber et al., 2018) and kazd1 (Bryant et al., 2017), were enriched in the stump dividing cells. Genes known to be up-regulated early upon amputation but more likely to be modulating the extracellular matrix degradation rather than marking progenitor cells, such as mmp9 and mmp3 (Vinarsky et al., 2005; Monaghan et al., 2012; Stewart et al., 2013), were enriched in non-dividing stump tissues (Figure 2.2C). To investigate the specificity of our strategy to enrich for blastemal progenitors rather than other dividing cell types, such as immune cell infiltrates, we examined the predicted activation and inhibition of immune signaling pathways by applying Ingenuity Pathway Analysis (IPA) software to the differentially expressed transcripts in each cell population relative to the non-regenerating limb. Most innate immune signaling pathways were inhibited within dividing cells (Figure

A.1.1, Table A.1.1), while activated in the other fractions, suggesting that we did not preferentially enrich for dividing immune cells. In all, these observations validated the enrichment of dividing blastemal progenitors using this protocol and allowed us to examine unique gene expression patterns of these cells.

Overall, a total of 21,077 transcripts (19,417 genes at 4 dpa) and 16,510 transcripts

(15,205 genes at 5 dpa) were differentially expressed across all three fractions during early regeneration (Figure 2.3B-C). Of these, only a strikingly small percentage were commonly up- or down-regulated across all tissues relative to non-regenerating tissue at

0 dpa (1.51% at 4 dpa and 0.58% at 5 dpa), strongly suggesting that all three subpopulations initiate distinct transcriptional programs following amputation. Lists of differentially expressed transcripts for all analyses are deposited on GEO (accession number: GSE111213). The most highly expressed and commonly upregulated transcripts corresponded to enzymes involved in modulating extracellular matrix (ECM) degradation

33

Figure 2.1. Technical validation of FACS-method to purify 4N and 2N fractions from regenerating stump tissue. (A) Representative FACS plot and gating for 2N (P3) and 4N (P5) populations in dissociated stump tissue based on DAPI staining. (B) Representative Bioanalyzer traces of RNA isolated from sorted stump-derived 2N and 4N fractions and non-sorted, but dissociated epidermis. (C) RT-PCR of cDNA generated from sorted 2N and 4N fractions for cell cycle genes enriched in the actively dividing cells.

34

Figure 2.2. Transcriptional profiles of dividing, non-dividing, and wound epidermal cells reveal fold change enrichment of well-known early regeneration genes. (A) Principal component analysis (PCA) plot of individual samples based on top 500 most highly expressed transcripts. Colors denote the specific sample and shapes denote the replicate of the sample. (B) Graph demonstrating high fold change enrichment of 20 cell cycle genes in 4N vs. 2N across all three timepoints. (C) Fold change enrichment of well- known genes involved in early limb regeneration in either the 4N or 2N fraction. Transcripts for blastemal cell markers such as kazd1 and prrx-1 were correctly enriched in the dividing cells.

35 (mmp18, mmp2, timp1, tena, adam8) and transcription factors involved in limb development and regeneration including sall4 (Neff et al., 2005; Akiyama et al., 2015;

Erickson et al., 2016) and runx-1 (Umansky et al., 2015; Deltcheva and Nimmo, 2017), indicating that they may play a role in orchestrating regeneration across all tissues.

2.4.2. Growth factor signaling pathways are largely repressed within regenerating stump tissues

We examined signaling interactions between regenerating subpopulations through

IPA analysis of differentially expressed transcripts in each fraction at both timepoints. IPA software utilizes algorithms that account for the expression levels of signaling components, activators, and inhibitors of well-known signaling pathways to predict whether a particular pathway is activated or inhibited. Surprisingly, pathway analysis revealed that several growth factor (e.g. FGF, Notch) (Figure 2.3D) and intracellular (e.g. calcium and mTOR) signaling pathways implicated in regeneration were strongly inhibited in regenerating stump tissues during these early timepoints (Table A.1.2). Notable exceptions include a handful of pathways: Hippo, canonical Wnt, and TGF- signaling.

Among the repressed pathways were neurovascular signaling pathways (CNTF, NGF,

Neuregulin, VEGF signaling) that were activated in the wound epidermis and suppressed in stump tissues. These data further suggest that the wound epidermis orchestrates early neurovascular regeneration and that early repression of these pathways in regenerating stump tissues may be required. In all, these results show that exact spatiotemporal modulation of growth factor signaling pathways in specific tissues occurs during early stages of regeneration.

36

Figure 2.3. Growth factor signaling pathways are inhibited in regenerating stump tissues. (A) Schematic of transcriptomic profiling experiment. Limbs were amputated and 2-3 mm of tissue proximal to the amputation site was collected at 0, 4, or 5 days post- amputation (dpa). Three fractions were isolated from each sample for sequencing: stump- derived dividing cells, non-dividing cells, and the whole wound epidermis. DAPI cell cycle analysis and FACS was performed to isolate dividing and non-dividing cells in the stump tissue. (B, C) Venn diagrams of the distribution of differentially expressed transcripts at 4

37 Figure 2.3. (Continued) or 5 dpa are shown in B and C, respectively. (D) Ingenuity Pathway Analysis of differentially expressed transcripts in stump-derived dividing, non- dividing, or wound epidermal cells. Positive Z-scores depict predicted activation whereas negative Z-scores depict predicted inhibition of the respective pathway. Color coding of labels for each fraction of tissue as follows: stump-derived 2N, blue, stump-derived 4N, green, wound epidermis, purple.

2.4.3. The transcriptional landscape of early dividing cells indicates roles in shaping the blastemal niche

We next utilized our dataset to identify transcripts enriched in dividing cells at both regenerating timepoints (see Materials and Methods for filtering criteria). A total of 1,217 transcripts (1,181 genes) were enriched in dividing cells and of these, only 298 transcripts

(265 genes) were annotated. A heatmap representing the top 75 annotated and most highly expressed enriched transcripts, with little to no expression in the wound epidermis, is shown in Figure 2.4A. A complete list of all annotated enriched transcripts identified can be found in Table A.1.3. The small fraction of annotated enriched transcripts suggests that many key modulators of early regenerative events might be novel genes or at least may not have identifiable orthologous genes in other species present in existing datasets.

Among the most highly expressed and enriched transcripts in dividing cells were regenerative ECM components including tenascin (tena), collagens (co5a2, co5a1, co1a2, coba1, coca1), emilin1 (emil1), and fibrillin-2 (fbn2), suggesting that dividing cells play an early role in building the blastemal niche. Most notably, the transcriptional signatures of early dividing cells suggest that they may be directly regulated by TGF- signaling. Pathway analysis revealed that TGF- signaling was specifically activated in early dividing cells (Figure 2.3D). Further examination of TGF- signaling components

38 revealed upregulation of both up- and down-stream regulators (tgf-1, tgf-1r, smad2, ltbp1, inhba) as well as direct targets of TGF- signaling associated with epithelial-to- mesenchymal transition (EMT) such as snail1 and twist1 (Figure 2.4B). These data indicate that an auto-regulatory TGF- signaling network is established both intra- and extracellularly in dividing cells, including blastemal progenitors, during early stages of regeneration.

To validate our differential-gene expression findings and to learn more about where these transcripts are expressed, we performed time course RNA in situ hybridization on two candidates with high enrichment in dividing cells, transmembrane protein 119 (tm119) and E3 ubiquitin-protein ligase (lin41) (Figure 2.4C-H). In situ hybridization confirmed that these transcripts were specific to early dividing cells and likely expressed in blastemal progenitors during early stages of regeneration. Tm119 has been shown to play a role in bone development and osteoblast proliferation (Kanamoto et al., 2009; Mizuhashi et al., 2012; Mizuhashi et al., 2015), whereas lin41 has a highly conserved role in stem cell maintenance as well as cellular reprogramming (Slack et al.,

2000; Worringer et al., 2014). Both candidates showed little to no expression in uninjured limbs (Figure 2.4C, F). Co-expression of tm119 and lin41 with a dividing cell marker, top2a, via double in situ hybridization at 7 dpa validated that these transcripts were expressed within dividing cells (Figure 2.4D, G). Moreover, in situ hybridization of tm119 and lin41 at 21 dpa revealed strong expression of tm119 across the blastema, and lower, but pan-blastemal expression of lin41 as well (Figure 2.4E, H), suggesting that they were indeed expressed within blastemal progenitors at earlier stages.

39 A B log2(TPM) Z-score

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F lin41 G top2a lin41 H lin41 0 dpa 7 dpa 21 dpa

4N 2N WE

Figure 2.4. Identification of highly enriched transcripts in dividing cells during early regeneration. (A) Heatmap of the expression levels (log2TPM) of the top 75 transcripts enriched in regenerating dividing cells in wound epidermis and non-dividing cells in the stump tissue. Green arrows denote transcripts for the blastemal markers kazd1 and prrx- 1; red arrows point to tm119 and lin41. (B) Heatmap of expression levels of select TGF-  signaling pathway regulators, targets, and downstream effectors. Color coding of labels in A and B for each fraction of tissue as follows: stump-derived 2N, blue, stump-derived 4N, green, epidermis or wound epidermis, purple. (C-E) In situ hybridization of tm119 at 0, 7, and 21 dpa. Double in situ hybridization of tm119 with top2a is depicted in panel D.

40 Figure 2.4. (Continued) Arrowheads in D demarcate co-positive tm119+top2a+ cells. (F- H) In situ hybridization of lin41 at 0, 7, and 21 dpa. Double in situ hybridization of tm119 and top2a is depicted in panel G. Arrowheads denote co-positive lin41+top2a+ cells. Scale bars represent 20 µm. Images were taken at 40x magnification. Insets in the bottom left corner denote where in the overall section the higher magnification image was taken. WE, wound epidermis.

2.5. Discussion

Elucidating the molecular mechanisms underlying the initiation of regeneration is key to understanding the difference in responses to tissue loss between regenerative and non-regenerative organisms. Here we transcriptionally profiled distinct regenerating subpopulations during early pre-blastemal stages of limb regeneration to differentiate gene expression changes that occur in early dividing cells, inclusive of blastemal progenitors, from those of surrounding tissues. Using this dataset, we have gained insights into the patterns of suppression and activation of signaling pathways present within subsets of early regenerating limb tissues. Notably, our examination of the expression profiles of early dividing cells revealed an immunomodulatory role for blastemal progenitors during early stages of regeneration.

The transcriptional signatures of early dividing cells and blastemal progenitors suggest that the formation of the early blastemal niche may be regulated by canonical

Wnt, Hippo, and TGF- signaling. Dividing cells showed enriched expression of regenerative ECM components, many of which are important for regeneration (Calve et al., 2010; Godwin et al., 2014), suggesting that they are drivers of the transition to a regenerative ECM. In addition, Hippo, Wnt, and TGF- signaling pathways were activated within dividing cells. As down-stream effectors of all three pathways interact in

41 development and tumorigenesis (McNeill and Woodgett, 2010; Attisano and Wrana,

2013), it is likely that synergy between these pathways is essential for early blastemal cell establishment and maintenance. Interestingly, TGF- signaling, which is necessary for axolotl limb regeneration (Levesque et al., 2007; Denis et al., 2016), was specifically active in dividing cells. Our data further suggest that TGF- signaling is sustained through autocrine feedback in early dividing cells, which exhibit exclusive up-regulation of tgf-1, tgf-1r, and smad-2 as well as regulators and direct targets including ltbp1, twist1, and snail1. Snail-1 directs epithelial-to-mesenchymal transition (EMT) behaviors (Fuxe et al.,

2010) and activates expression of both regenerative ECM components and twist family members, which are expressed in limb blastemal cells (Kragl et al., 2013; Bryant et al.,

2017). Moreover, dividing cells highly expressed ECM components such as emilin-1 and fibrillin-2, which modulate TGF- signaling through interactions with ltbp1 (Neptune et al.,

2003; Randell and Daneshtalab, 2017), indicating that both an intra- and extra-cellular

TGF- signaling network is established.

Surprisingly, we observed the strong repression of many growth factor signaling pathways in early regenerating stump tissues. FGF, Notch, IGF-1, PDGF, and non- canonical Wnt signaling pathways (PCP and Wnt/Ca+) all appear to be inhibited, yet many of these signaling pathways are necessary and/or sufficient for appendage regeneration in amphibians as well as zebrafish (Poss et al., 2000; Yokoyama et al., 2000; Yokoyama et al., 2001; Stoick-Cooper et al., 2007; Chablais and Jazwinska, 2010; Satoh et al., 2011;

Grotek et al., 2013; Makanae et al., 2014; Rodrigo Albors et al., 2015; Currie et al., 2016;

Nacu et al., 2016; Shibata et al., 2016). Furthermore, signaling pathways involved in neuronal (e.g. Neuregulin) and vascular (e.g. VEGF) regeneration (Yu et al., 2014; Farkas

42 et al., 2016; Farkas and Monaghan, 2017; Ritenour and Dickie, 2017) were active in the wound epidermis, yet suppressed in regenerating stump tissues, signifying that early neurovascular regeneration is coordinated by the wound epidermis and that repression of these pathways within regenerating stump tissues may be important. A potential explanation for this apparent dichotomy lies in the timing of activation. Premature activation of signaling pathways such as Notch (Grotek et al., 2013) or non-canonical Wnt signaling (Stoick-Cooper et al., 2007) inhibits blastemal growth during zebrafish fin regeneration. Therefore, our findings suggest that repression of these pathways in early stages of regeneration may be necessary and that precise, timely release of inhibition ensures successful regeneration. It is interesting to note that the analysis did not reveal strong predictions for activation or repression of other pathways essential for limb regeneration such as sonic hedgehog (shh) signaling (Singh et al., 2015). These other pathways may be downstream and act at later stages of regeneration (or are regulated post-transcriptionally). Nevertheless, these findings could provide targetable insights for improving regenerative outcomes.

2.6. Materials and Methods

Animal procedures

Axolotl (Ambystoma mexicanum) husbandry and surgeries were performed in accordance with the Association for Assessment and Accreditation of Laboratory Animal

Care (AAALAC) and Institutional Animal Care and Use Committee (IACUC) guidelines at

Harvard University. Sub-adult white and albino axolotls (15-18 cm) provided by the

43 Ambystoma Genetic Stock Center (AGSC, University of Kentucky, KY) were used for the

RNA-sequencing experiment.

FACS and RNA isolation

Briefly, animals were anesthetized in 0.1% Tricaine (Sigma-Aldrich, St. Louis, MO) and all four limbs were amputated at the mid-radius/ulna level. Bone was trimmed back to facilitate wound closure and regeneration. Approximately 2-3 mm of tissue directly proximal to the amputation plane was collected at either 0, 4, or 5 dpa. We chose to transcriptionally profile regenerating limbs at 4 and 5 dpa because preliminary experiments showed a distinct increase in cellular proliferation in the regenerating stump beginning at 3 dpa (data not shown here) and we wanted to capture the transcriptional signatures of dividing cells during the duration of cell-cycle re-entry. In order to obtain enough material for the sorting and sequencing protocol, we chose to perform the experiments at 4 and 5 dpa. Twelve limbs were pooled for each biological replicate per timepoint and the experiment was performed in biological triplicate.

To prep the tissue for FACS, the intact epidermis or wound epithelium was micro- dissected off and placed into 0.25% Trypsin-EDTA for 15 minutes with agitation at room temperature to dissociate epithelial cells from the dermal layer. The remaining stump tissues were micro-dissected further into small pieces with dissecting scissors and chemically dissociated in a solution composed of 5 mg/mL collagenase (Sigma-Aldrich),

7.3 mg/mL dispase II (Roche Diagnostics, Indianapolis, IN), and 1.36 mg/mL D-Glucose in 80% PBS (Kumar and Brockes, 2007) for 15 minutes with agitation at room temperature. In order to obtain representation of dermal cells in the stump tissue fraction,

44 we also chemically dissociated micro-dissected intact epidermis or wound epithelia along with the stump tissues. For regenerating samples, the wound epidermis portion, which is visibly transparent, was carefully removed to isolate adjacent full thickness skin with both epidermal and dermal layers and then dissociated with the stump tissue fraction.

Dissociated cells were then transferred to a new tube (non-dissociated chunks of tissue were left behind) and the dissociation was serum-inactivated. The following protocol for staining, FACS, and RNA isolation was optimized and adapted from Hrvatin et al. 2014.

The cell suspension was pelleted, re-suspended, passed through a 70 uM cell strainer, washed twice with 80% PBS, and fixed in 4% paraformaldehyde/0.1% saponin with a

1:50 dilution of RNasin plus RNase Inhibitor (Promega, Madison, WI) for 30 minutes at

4C. Fixed cells were then washed twice with a 1% BSA/ 0.1% saponin (1:40 dilution

RNasin plus RNase Inhibitor, Promega) in PBS wash buffer and DAPI staining (10 ug/mL) of cells was performed for 30 minutes at 4C in a 0.1% saponin solution (1:20 RNasin plus RNase Inhibitor, Promega). The stump fraction of DAPI-stained cells were immediately sorted into 2N and 4N fractions using FACS, whereas the corresponding epithelial fraction was subjected to the same staining treatment, but not sorted. Cells were sorted into RNAlater solution (Invitrogen, Carlsbad, CA) and RNA was isolated with the

RecoverAll Total Nucleic Acid Isolation Kit for FFPE (Invitrogen). The quality of the RNA samples was assessed using the Agilent RNA 6000 Pico kit on the Bioanalyzer 2100

(Agilent Technologies, Santa Clara, CA).

cDNA library preparation and sequencing

45 cDNA was synthesized from RNA samples using the NuGEN Ovation RNA-seq

System V2 protocol (Integrated Sciences, Sydney, Australia) according to the manufacturer’s instructions using 10 ng of RNA as starting material. The quality and concentration of cDNA preps was then assessed using the Agilent DNA 1000 kit on the

Bioanalyzer 2100 (Agilent Technologies). RT-PCR for cell cycle markers ccnb3, ccna2, and cdk1b was then performed on the cDNA generated from 4N and 2N cells to check for proper enrichment of dividing cells. Primer sequences are as follows:

ccnb3-For, 5’-CACAAGAATCCAGTGCCACA-3’

ccnb3-Rev, 5’-CCTCCTTTGCAACAGTGTCC-3’

ccna2-For, 5’-GAACGTACAGCCTGGCAAG-3’

ccna2-Rev, 5’-CTGACGGCTGCTCCTTTG-3’

cdk1b-For, 5’-GCCAAACAACGAAATCTGGC-3’

cdk1b-Rev, 5’-AGGGTGGTTCAATGCCTCTT-3’

To prepare cDNA sequencing libraries, 200 ng of cDNA was first sheared to a peak size of 200 bp using the Covaris S220 (Covaris, Woburn, MA) according to the manufacturer’s protocol. Good quality and correct size distribution of sheared DNA fragments was assessed with the Agilent DNA High Sensitivity kit on the Bioanalyzer

2100 (Agilent Technologies). Sequencing libraries were then generated using the

Wafergen PrepX Complete ILMN DNA Library kit (Takara Bio, Mountainview, CA) protocol on the Apollo 324 NGS Library Prep System (Takara Bio). Quality of the DNA sequencing libraries was performed with the Agilent DNA High Sensitivity kit on the

Bioanalyzer 2100 (Agilent Technologies). Concentration of the DNA libraries was doubly confirmed using the Qubit dsDNA HS Assay kit (ThermoFisher Scientific, Waltham, MA)

46 and Kapa Illumina Library Quantification kit (Kapa Biosystems, Wilmington, MA). Libraries were multiplexed and sequenced on either the Illumina Hiseq 2500 system (125 bp reads) or Nextseq 500 (150 bp reads) (Illumina, San Diego, CA) at the Harvard Bauer Core

Sequencing Facility.

Sequencing analysis

Reads were trimmed using Trimmomatic (Bolger et al., 2014) to a minimum length of 100 bp and poor quality reads were removed from the sequencing analysis. Alignment was performed on the trimmed reads using Kallisto (Bray et al., 2016) and the previously published well-annotated axolotl transcriptome (Bryant et al., 2017). Raw read data and the processed data matrix containing TMM-normalized TPM values for each sample are accessible on the NCBI GEO database: accession GSE111213. Differential expression analysis of genes and transcripts relative to the non-regenerating timepoint (0 dpa) for each fraction was performed using DESeq2 (Love et al., 2014) with an adjusted p-value cutoff of 0.05. Core and comparison analysis of differentially expressed transcript lists for each cellular population was performed using Ingenuity Pathway Analysis software

(Qiagen, Hilden, Germany). Uniprot IDs of transcript blast hits (extracted using the associated Trinotate file in Bryant et al. (2017) were converted to human IDs for IPA analysis. We focused on strongly activated or inhibited growth factor signaling pathways i.e. absolute value of the Z-score > 1 and signal detected across at least 3 differentially expressed analyses out of the 6 total.

To identify transcripts enriched within dividing cells at both timepoints, we first compared dividing and non-dividing cells averaged at both regenerating timepoints (4 and

47 5 dpa) and identified 6,834 differentially expressed transcripts (3,510 of which were up- regulated within dividing cells). Transcripts that were normally differentially expressed in non-regenerating dividing and non-dividing cells (at 0 dpa), such as cell cycle-associated transcripts, were filtered out. Of these, 628 total transcripts (583 genes) were regeneration-specific, annotated, and had at least a 2-fold change between dividing and non-dividing cells and only 298 transcripts in this list were up-regulated in dividing cells.

In situ hybridization and quantification

Tissue was collected at 0, 7 and 21 dpa and fixed in 4% paraformaldehyde overnight at 4C, washed in PBS, brought up a sucrose gradient to 30% sucrose, and embedded in OCT. The blocks were serially sectioned and 16 um sections were collected.

Custom RNAscope probes for the axolotl orthologs of top2a, tm119, and lin41 were generated (Advanced Cell Diagnostics, Newark, CA) in either the C1 or C2 channels.

Double chromogenic section in situ hybridization was performed using the RNAscope 2.5

HD Duplex Detection Kit (Advanced Cell Diagnostics) according to the manufacturer’s instructions on frozen cryosections.

2.7. References

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54

Chapter 3

Blastemal progenitors modulate immune signaling during early limb regeneration

3.1. Preface

The work presented in this chapter was largely modified and adapted to fit the guidelines of this dissertation from the following publication: Tsai S.L., Baselga-Garriga

C., and Melton D.A. (2019). Blastemal progenitors modulate immune signaling during early limb regeneration. Development:146(1). The conceptualization and experimental design was performed in collaboration with Doug. I performed all of the experiments and the data analysis, except for the blinded counting analysis of the myeloid cells in the il-8 and control morphants, which was performed by Clara.

3.2. Abstract

The list of blastemal progenitor-enriched transcripts during early limb regeneration generated in Chapter 2 was a rich source of gene candidates that could help us and others in the field gain further insight into blastema cell biology. In this chapter, we examined the functional role of one enriched candidate, il-8, in limb regeneration. Through the experiments detailed within this section, we demonstrate that blastemal progenitors play an immunomodulatory role during early stages of regeneration via il-8 signaling. In addition, we also identify and establish the importance of early CXCR-1/2 signaling in limb regeneration. In all, these results suggest that bi-directional cross-talk between the blastemal cells and immune system is essential for successful regeneration.

3.3. Introduction

Many advances have been made in the last decades revealing crucial roles that immune cells play in modulating tissue repair and injury responses. Beyond their

56 canonical functions in debris clearance and preventing infection as a consequence of injury, immune cells have been shown to actively stimulate progenitor stem cell activation and differentiation in a variety of organ and tissue regenerative contexts (Aurora and

Olson, 2014).

Out of the entire immune cell repertoire, macrophages are perhaps the most well studied in regeneration, although others have begun to demonstrate the importance of other cell types including Tregs and eosinophils (Burzyn et al., 2013; Heredia et al., 2013).

Macrophages can be polarized into two interchangeable functional phenotypes, M1 (pro- inflammatory) and M2 (pro-regenerative) depending on extracellular cues from the surrounding environment (Sica and Mantovani, 2012). Various studies have demonstrated the significance of the shift from M1 to M2 states between early and late stages of repair or regeneration in different contexts including skeletal muscle (Mounier et al., 2013) and central nervous system (CNS) regeneration in mice (Miron et al., 2013).

Interestingly, while M1 macrophages have traditionally been thought to be detrimental for regenerative outcomes, they also play essential early roles in these contexts as well including progenitor cell activation and debris clearance. In fact, premature activation of

M2 polarization impairs skeletal muscle regeneration (Perdiguero et al., 2011), indicating that fine temporal modulation of M1/M2 polarization is necessary for successful repair.

The essential roles that macrophages play in organ and tissue injury contexts in mice have translated to highly regenerative species as well. Macrophages have been shown to be necessary for initiating appendage, heart, and ear hole regeneration in the axolotl, zebrafish, and African spiny mouse (Godwin et al., 2013; Petrie et al., 2014;

Godwin et al., 2017; Simkin et al., 2017a). In addition, most of the work in the regenerative

57 models of salamanders and zebrafish has focused on the importance of myeloid cells in the innate immune system including macrophages and neutrophils. These cells are among some of the most abundant first responders post-amputation.

We noticed that several inflammatory cytokines were enriched within blastemal progenitors in the dataset, one of which was interleukin-8 (il-8), a well-known chemoattractant of neutrophils and macrophages (de Oliveira et al., 2013; Xu et al.,

2018). While many have shown the important paracrine effects that the immune system has on successful regeneration, less is known about non-immune cell cues that direct myeloid cell behaviors in these contexts. Our transcriptional data therefore suggested that blastemal progenitors may directly modulate the immune system during regeneration.

To determine whether this was indeed the case, we investigated the expression of il-8 throughout the course of limb regeneration and examined its potential role with gain- and loss-of-function approaches. Il-8 expression was highly induced in blastemal progenitors of the connective tissue lineages specifically during wound healing stages of limb regeneration. Ectopic il-8 expression in non-regenerating limbs induced myeloid cell recruitment, while IL-8 knockdown resulted in defective myeloid cell retention during late wound healing, delaying regeneration. Furthermore, the il-8 receptor, cxcr-1/2, was expressed in myeloid cells and inhibition of CXCR-1/2 signaling during early stages of limb regeneration prevented regeneration. Altogether, our findings suggest that blastemal progenitors are active, early mediators of immune support and identify CXCR-1/2 signaling as an important early immunomodulatory pathway in limb regeneration.

58 3.4. Results

3.4.1. Interleukin-8 (il-8) is expressed in early blastemal progenitors

Time course in situ hybridization for il-8 during limb regeneration revealed a strong, transient up-regulation during early stages of regeneration (Figure 3.1). Il-8 was undetectable in non-regenerating limbs (Figure 3.1A-A’) and highly induced upon amputation (Figure 3.1B-E’). At 1 dpa, il-8 was expressed in cells lining the bone (Figure

3.1B-B’) and in the dermis. Il-8 expression peaked at 3 dpa in mesenchymal cells within regenerating stump tissues and began to decrease by 7 dpa (Figure 3.1C-D’). By early blastemal stages at 14 dpa, il-8 was weakly and sparsely expressed throughout the blastema and basal layers of the apical epithelial cap (AEC) (Figure 3.1E-E’).

Previous studies have shown that il-8 is highly expressed in invading monocytes in wound healing and tumorigenic contexts (Fu et al., 2015; Williams et al., 2016). In contrast, double in situ hybridization of il-8 with two blastemal cell markers prrx-1 and kazd1, as well as csf1-r, a monocyte marker in mammals and amphibians (Grayfer et al.,

2014; Stanley and Chitu, 2014) confirmed that il-8 was indeed expressed within blastemal progenitors and not monocytes. At both 3 and 7 dpa, il-8 was co-expressed in a subpopulation of prrx-1+ cells (Figure 3.1F), and exhibited little to no expression in csf1- r+ monocytes (Figure 3.1G). Quantification of prrx-1+ and il-8+ populations revealed high concordance between prrx-1 and il-8 expression. At 3 dpa, 99.0% of prrx-1+ cells expressed il-8 and 96.6% of il-8+ cells expressed prrx-1. At 7 dpa, only 67.9% of prrx-1+ cells expressed il-8, but 97.5% of il-8+ cells expressed prrx-1, consistent with a decrease in il-8 expression over time (Figure 3.1H). In contrast, at 3 dpa, only 3.8% of il-8+ cells were

59 A 0 dpa B 1 dpa C 3 dpa D 7 dpa E 14 dpa

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Figure 3.1. Il-8 is strongly expressed in blastemal progenitors during early stages of limb regeneration. (A-E’) Time course double in situ hybridization of il-8 at 0, 1, 3, 7, and 14 dpa. Il-8 is not expressed in non-regenerating limbs (0 dpa), begins expression at 1 dpa, peaks at 3 dpa, and remains strongly expressed, but in fewer cells at 7 dpa. By early blastemal stages,14 dpa, il-8 expression has largely diminished, with weak expression in sparse blastema cells and in the basal layers of the wound epidermis. The amputation plane is demarcated with a solid line in panels A-E. Higher 40x magnification images of the insets are shown in A’-E’. The dotted line in E’ marks the wound epidermis boundary. (F) Double in situ hybridization of il-8 with the blastemal cell marker prrx-1 at

60 Figure 3.1. (Continued) both 3 and 7 dpa shows colocalization between il-8 and prrx-1. Examples of co-positive cells are denoted with arrowheads. (G) Double in situ hybridization of il-8 with the monocyte marker csf-1r at both 3 and 7 dpa shows little to no colocalization between il-8 and csf-1r. Representative single positive csf-1r+ cells are denoted with arrows, while single positive il-8+ cells are denoted by arrowheads. (H-I) Quantification of double in situs shown in panels F and G. Graph in H shows percentage breakdown of total counted il-8 and/or prrx-1 expressing cells (3 dpa, n=298 cells total, 7dpa, n=412 cells total). Graph in I shows percentage breakdown of total counted il-8 and/or csf-1r expressing cells (3 dpa, n= 500 cells total, 7 dpa, n=630 cells total). (J) Double in situ hybridization of il-8 (blue) and kazd1 (red), a highly expressed blastemal cell marker reveals co-expression of il-8 in a subset of kazd1+ blastemal cells (denoted by arrowheads). Image was taken at 40x magnification. Scale bars in A-E, 200 µm, A’-E’, 50 µm, F, G, and J, 50 µm.

csf1r+ and 1.7% of csf1r+ cells were il-8+, while at 7 dpa, 4.4% of il-8+ cells were csf1r+ and 1.5% of csf1r+ cells were il-8+ (Figure 3.1I). Il-8 was also expressed in a subset of kazd1+ blastemal progenitors at 7 dpa (Figure 3.1J). Altogether, these data provide evidence that il-8 is strongly expressed within a subset of blastemal progenitors primarily during early stages of limb regeneration.

3.4.2. IL-8 is sufficient to induce myeloid cell recruitment and proliferation in bone/perichondrium and epidermis in non-regenerating limbs

Because il-8 was strongly expressed in blastemal progenitors, we examined whether il-8 expression was sufficient to induce cell behaviors characteristic of the initiation of blastema formation, such as immune cell recruitment or cellular proliferation.

We designed a myc-tagged il-8 overexpression vector (pCMV-IL8myc-T2A-tdTomato) and control vector (pCMV-T2A-tdTomato) and validated secretion of myc-tagged IL-8

61 protein in 293T cells (Figure A.2.1). We injected and electroporated either the control tdTomato vector or the il-8 overexpression vector into intact limbs of non-regenerating animals, briefly pulsed the animals with EdU at 3 days post-injection, and collected samples 24 hours later (Figure 3.2A).

As IL-8 is a well-known inflammatory cytokine (Zeilhofer and Schorr, 2000), we first examined whether ectopic expression of il-8 induced monocyte and granulocyte recruitment by performing -Naphthol Acetate (NSE) staining and Naphthol AS-D

Chloroacetate (NCAE) staining, respectively. A 2.3-fold increase in NSE+ monocytes

(117.53 vs. 51.94 cells/mm2, p = 0.0041) and a 2.0-fold increase in NCAE+ granulocytes

(121.54 vs. 61.97 cells/mm2, p = 0.0353) was observed in limbs expressing il-8 relative to controls (Figure 3.2B-D’), indicating that IL-8 is sufficient to recruit myeloid cells.

We also assessed whether IL-8 could induce cellular proliferation by comparing the percentage of total EdU+ cells between tdTomato and il-8 expressing limbs. We observed a modest, yet significant increase in the total number of EdU+ cells in limbs overexpressing il-8 compared to control tdTomato limbs (13.92% vs 11.27%, p = 0.0015)

(Fig 3.3A, C). Quantification of dividing bone/perichondrial, epidermal, endothelial, and satellite cells revealed that this total increase in dividing cells was primarily due to an increase in the percentage of dividing bone/perichondrial cells (17.78% vs. 8.71%, p =

0.004), though there is a modest increase in the percentage of dividing epidermal cells

(24.60% vs. 19.70%, p = 0.0146) (Figure 3.3B-E). No increase in proliferation was observed in CD34+ endothelial cells or pax7+ muscle satellite cells. These data suggest that ectopic il-8 expression is sufficient to promote proliferation of bone/perichondrial and epidermal cells in a non-regenerative context, i.e. without amputation.

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Figure 3.2. Il-8 induces recruitment of monocytes and granulocytes in intact limbs. (A) Experimental schematic of il-8 overexpression experiment. The left limbs of axolotls were injected and electroporated with a T2A-tdTomato control construct, whereas the right limbs were injected and electroporated with an il8myc-T2A-tdTomato construct. Animals were pulsed with EdU 24 hours prior to tissue collection at 4 days post-injection (dpi). (B) Quantification of monocytes and granulocytes per mm2 revealed a statistically significant increase in both monocytes (**pval = 0.0041, n=9) and granulocytes (*pval = 0.0353, n=9) in il-8 overexpression limbs. Statistical analyses were performed using a two-tailed paired t-test. Error bars represent mean ± SD. (C-D’) Representative 10x montage images of NSE/NCAE stained limbs in control or il-8 overexpression limbs are shown in C and D, respectively. Scale bars denote 200 µm. Higher 20x magnification images of the insets in panels C’ and D’, respectively. NSE+ monocytes are stained in

63 Figure 3.2. (Continued) black and NCAE+ granulocytes are stained in purple. Arrowheads point to representative monocytes and arrows point to representative granulocytes. Scale bar denotes 20 µm and the dotted lines denote bone.

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C tdTomato EdU DAPI EdU

o

t

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EdU DAPI EdU DAPI Figure 3.3. Il-8 is sufficient to induce proliferation of bone/perichondrial cells and epidermis. (A-B) Quantification of total EdU+ cells and cell-type specific EdU+ cells in control vs. il-8 overexpression limbs revealed a mild statistically significant increase in the total percentage of EdU+ cells (**pval = 0.0015, n=8), a strong significant increase in the percentage of dividing bone/perichondrial cells (***pval = 0.0004, n=8), and a mild increase in the percentage of dividing epidermal cells (*pval = 0.0146, n=8). No significant increase was detected in CD34+ endothelial cells or pax7+ satellite cells. All statistical analyses were done using a two-tailed paired t-test. Error bars represent mean ± SD. (C)

65 Figure 3.3. (Continued) Representative 10x montage images from control or il-8 overexpression limbs. Scale bars denote 200 µm. (D-E) High magnification images of insets from C. Representative 20x images of the bone and surrounding perichondrium of control (D) or il-8 (E) overexpression limbs. Arrowheads denote EdU+ perichondrial cells. Scale bars denote 20 µm. The yellow dotted lines in panels C-E denote bone within the tissue. Ns, non-significant.

3.4.3. Knockdown of IL-8 results in delayed blastemal outgrowth and regeneration

We next examined whether loss of function of il-8 may impair limb regeneration.

To this end, we performed a dual pulse morpholino experiment (Figure 3.4A) using translation-blocking il-8 targeted or control morpholinos, injected into either the right or left forelimb, at 2 days pre-amputation and 3 dpa. This treatment produced a strong and transient knockdown of endogenous IL-8 protein levels (Figure A.2.1F). Il-8 morpholino treated limbs displayed delayed blastemal outgrowth (Figure 3.4B). Quantification at 21 dpa revealed a 37% decrease in blastemal length (572.89 vs. 915.40 µm, p < 0.0001) and a 32% reduction in blastemal area (491.29 vs. 723.90 µm, p = 0.0005) (Figure 3.4C-

F). In addition, control limbs reached digit stages of differentiation before il-8 morphant limbs (Figure 3.4B), indicating that transient knockdown of IL-8 delays limb regeneration.

At this time point, there was no defect in cell proliferation in either blastemal cells or the wound epidermis (Figure A.2.2) and no visible phenotypes with the blastema aside from the difference in size. These observations in conjunction with the strong expression of il-

8 during early stages of limb regeneration suggested that il-8 morphant limbs were likely exhibiting defects at earlier timepoints in regeneration.

66 A Amputation

MO Injection MO Injection Collect Tissue

dpa -2 0 3 21 (days post-amputation)

B control MO il-8 MO C control MO D il-8 MO

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1500 1500

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Figure 3.4. Il-8 knockdown results in delayed blastemal outgrowth and regeneration. (A) Schematic of the dual pulse morpholino experiment. The forelimbs of an axolotl were injected with either control morpholino (left limb) or translation-blocking il- 8 morpholino (right limb) at 2 days pre-amputation and 3 days post-amputation. Limbs were then collected at 21 dpa. (B) Representative brightfield images of the delay in blastemal outgrowth and regeneration. Scalebars denote 1 mm. (C-D) Picromallory stained sections of control and il-8 MO injected limbs. Scalebars represent 500 µm. The dotted lines demarcate the amputation plane. (E) Quantification of blastema length in control vs. il-8 MO injected limbs. Il-8 MO injected limbs exhibited a statistically significant decrease in blastema length (****pval < 0.0001, n=19). (F) Quantification of blastema area in control vs. il-8 MO injected limbs. Il-8 MO injected limbs exhibit a significant decrease in blastemal area (***pval < 0.0005, n=19). Paired two-tailed t-tests were employed for statistical analyses. Error bars represent mean ± SD.

67 3.4.4. IL-8 knockdown leads to defective retention of myeloid cells during the wound healing transition to blastema formation

Since ectopic il-8 expression was sufficient to recruit immune cells, we hypothesized that il-8 morphant limbs may exhibit deficiencies in myeloid cell recruitment.

Surprisingly, we observed no significant deficiency in recruitment of either NCAE+ granulocytes or NSE+ monocytes to the amputation site at either 1 dpa or 5 dpa (Figure

3.5A, D). However, the mean density of granulocytes was lower overall than that of controls at both of these timepoints, suggesting that il-8 may play a minor role in recruiting granulocytes during wound healing. More interestingly, there was a significant decrease in both granulocytes (374.16 vs. 483.24 cells/mm2, p < 0.01) and monocytes (198.20 vs.

290.92 cells/mm2, p<0.05) during later stages of wound healing at 7 dpa (Fig 3.5). We did not observe a difference in overall cell proliferation or cell death at this time point.

Therefore, the data suggest that il-8 likely plays a role in retaining both granulocytes and monocytes during the transition from wound healing to blastema formation.

To determine whether IL-8 was directly regulating myeloid cell behaviors, we examined the expression pattern of its cognate receptors, cxcr-1 and cxcr-2, during early stages of regeneration. While other organisms including mammals have both receptors, analysis of publicly available transcriptomes for the axolotl (Bryant et al., 2017;

Nowoshilow et al., 2018) revealed only one receptor with homology to both cxcr-1 and cxcr-2 in different species. We therefore refer to the detectable cognate receptor as cxcr-

1/2. Double in situ hybridization of cxcr-1/2 with csf1r or mpo, a neutrophil marker (Walters et al., 2010), at 3 dpa revealed co-expression of cxcr-1/2 with subpopulations of csf1r+ and mpo+ cells (Figure 3.6), suggesting that IL-8 likely directly acts on subpopulations of

68 A B control MO B’ control MO il-8 MO

800 ns

** * ns

2 600

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/ granulocytes

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control MO D control MO il-8 MO E E’

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0 * monocytes 1 dpa 5 dpa 7 dpa

Figure 3.5. Il-8 knockdown results in defective retention of myeloid cells during the transition from wound healing into blastema formation. (A) Quantification of NCAE+ granulocytes in control or il-8 MO limbs at 1, 5, or 7 dpa revealed a statistically significant decrease at 7 dpa (**p < 0.01, n=9). Error bars represent mean ± SD. (B-C’) Representative 10x montage images of control and il-8 MO limbs at 7 dpa are shown in B and C, respectively. Insets represent 40x magnification images shown in B’ and C’. (D) Quantification of NSE+ monocytes in control or il-8 MO limbs at 1, 5, or 7 dpa revealed a statistically significant decrease at 7 dpa (*p < 0.05, n=9). Error bars represent mean ± SD. (E-F’) Representative 10x montage images of control and il-8 MO limbs at 7 dpa are shown in E and F, respectively. Insets represent 40x magnification images shown in E’ and F’. Paired two-tailed t-tests were employed for statistical analyses. Scalebars in B-C and E-F denote 200 um; in B’-C’ and E’-F’ represent 50 µm. Asterisks (*) in B-C and E-F denote the bone. Dotted lines in B’-C’ and E’-F’ demarcate the boundary of the bone.

69 A csf1r cxcr1/2 B mpo cxcr1/2

Figure 3.6. Cxcr-1/2 is expressed in subsets of monocytes and granulocytes. Double in situ hybridization of cxcr-1/2 and either csf1r, a monocyte marker (A), or mpo, a neutrophil marker (B). Both co-positive cxcr-1/2+ csf1r+ cells and cxcr-1/2+ mpo+ cells were apparent at 3 dpa (denoted by orange arrows). However, not all csf1r+ and mpo+ cells were positive, suggesting cxcr-1/2 is expressed in a subset of monocytes and granulocytes. Insets show where in the section the 63x magnification image was taken. Scale bar denotes 20 m.

monocytes and granulocytes including neutrophils during wound healing. Altogether, these data provide one of the first examples of blastemal progenitors as immunomodulators, specifically as the source of canonical chemokines, during wound healing and further suggests that retention of myeloid cells during the transition to blastema formation is critical.

3.4.5. CXCR-1/2 signaling is necessary for limb regeneration

Finally, we asked whether inhibition of CXCR-1/2 signaling would also impair limb regeneration. We perturbed the pathway by treating regenerating axolotls with a small molecule inhibitor of CXCR-1/2, SB-225002 (White et al., 1998) (Figure 3.7A). Inhibitor treatment beginning from the time of amputation (0 dpa) completely inhibited limb regeneration, while DMSO-treated limbs regenerated normally (Figure 3.7B-E). Due to

70

Figure 3.7. Early CXCR-1/2 signaling is necessary for limb regeneration. (A) Experimental schematic of different SB-225002 drug treatments beginning at 0, 3, or 15 dpa. (B-E) SB-225002 treatment beginning from 0 dpa inhibits limb regeneration. Brightfield images of regenerating limbs treated with DMSO or SB-225002 at 17 dpa are shown in B and C, respectively. Alcian stained DMSO- or SB-225002-treated limbs at 40 dpa are shown in D and E. (F-I) SB-225002 treatment beginning at 3 dpa prevents limb regeneration. Brightfield images of DMSO- or SB-225002-treated limbs at 17 or 40 dpa are shown in panels F-I. (J-K) Limbs regenerate normally if SB-225002 treatment begins after blastema formation (15 dpa). Brightfield images of DMSO- or SB-225002-treated limbs at 30 dpa are shown in panels J and K, respectively. Arrows in B-K denote the amputation plane. Scale bars in B-K represent 1 mm.

71 the well documented role of CXCR-1/2 signaling in immediate wound-associated inflammatory responses (Ha et al., 2017), we hypothesized that failure to regenerate in this context was mainly due to prevention of immediate inflammatory responses stimulated by limb amputation. Therefore, we treated regenerating animals with SB-

225002 beginning at later stages of wound healing (3 dpa) and found that this also prevented regeneration (Figure 3.7F-I), suggesting that prolonged CXCR-1/2 signaling is essential. Yet, treatment beginning after blastema formation (15 dpa) led to normal limb regeneration (Figure 3.7J-K). In all, these data suggest that CXCR-1/2 signaling is necessary during early stages of regeneration, but dispensable for later stages.

Since il-8 morphants displayed defective retention of myeloid cells during late wound healing, we examined whether inhibition of CXCR-1/2 signaling affected myeloid cell behavior similarly during early stages of regeneration. Unexpectedly, monocytes and granulocytes in SB-225002-treated limbs (from 0 dpa) displayed unhealthy morphologies at 7 dpa. The majority of NSE+ and NCAE+ cells at the distal amputation site had formed apoptotic bodies, suggestive of myeloid cell death (Figure 3.8A). Quantification of healthy

NCAE+ and NSE+ myeloid cells revealed statistically significant decreases in both populations in SB-225002-treated limbs at 7 dpa compared to DMSO controls (Figure

3.8B-C, NCAE+ granulocytes: 128.9 vs. 362.3 cells/mm2, p<0.0001, NSE+ monocytes:

94.71 vs. 240.6 cells/mm2, p=0.0022). In concurrence with these observations, TUNEL staining of inhibitor treated limbs revealed higher levels of cell death near the amputation plane at 7 dpa (6.03% vs. 1.47%, p=0.0154) (Figure 3.8D), suggesting that myeloid cells were likely undergoing apoptosis. Interestingly, increased cell death was not observed in il-8 morphant limbs, suggesting that other cytokines in addition to il-8 may synergistically

72

A Granulocytes Monocytes TUNEL

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Figure 3.8. CXCR-1/2 inhibition impacts myeloid cell survival during early limb regeneration. (A) Images of NCAE+, NSE+, or TUNEL+ cells in DMSO- or SB-225002- treated limbs at 7 dpa with treatment beginning from 0 dpa (DMSO, n=4, SB-225002, n=5). Insets of fully stained sections in bottom left corners show where the higher magnification image was taken. Scale bars represent 50 µm. Asterisks denote autofluorescent bone in TUNEL-stained sections. White arrows denote healthy NCAE+ or NSE+ cells, whereas white arrowheads denote unhealthy NCAE+ or NSE+ cells exhibiting apoptotic bodies. (B) Quantification of NCAE+ granulocytes with healthy morphology revealed a statistically significant decrease in SB-225002-treated limbs (362.3 vs. 128.9 cells, ****p<0.0001). (C) Quantification of NSE+ granulocytes with healthy morphology revealed a statistically significant decrease in SB-225002-treated limbs (240.6 vs. 94.71 cells, **p=0.0022). (D) Quantification of TUNEL+ nuclei revealed higher levels of cell death in SB-225002-treated limbs (6.033% vs. 1.46%, *p=0.0172). Unpaired two-tailed t-tests were employed for statistical analyses. All error bars represent mean ± SD.

73 modulate myeloid cell behaviors. Altogether, these data newly identify CXCR-1/2 signaling as an immunomodulatory pathway that is in part, regulated by IL-8, and crucial for successful blastema formation.

3.5. Discussion

Here we provide one of the first examples that blastemal progenitors play an early paracrine immunomodulatory role in appendage regeneration. Others have shown that

IL-8 recruits neutrophils and macrophages in zebrafish organ and appendage regeneration (de Oliveira et al., 2013; Xu et al., 2018); however, the source of IL-8 was not examined. Contrary to other injury and tumorigenic contexts, where IL-8 is secreted from macrophages (Arango Duque and Descoteaux, 2014), we found that the primary source of IL-8 in early stages of limb regeneration is a subpopulation of blastemal progenitors. Furthermore, the high concordance of expression between il-8 and prrx-1, recently demonstrated as a connective tissue blastemal cell marker (Gerber et al., 2018), suggests that il-8 is primarily expressed in early regenerating connective tissue.

Meanwhile, its receptor cxcr1/2 is expressed in myeloid cells. Our functional data further shows the importance of signaling between blastemal progenitors and the immune system. Il-8 knockdown delayed regeneration and CXCR-1/2 inhibition prevented limb regeneration. We also demonstrate that Il-8 can recruit both myeloid cell-types in non- regenerating limbs. Thus, while leukocytes have been shown to regulate blastemal cells in other models (Nguyen-Chi et al., 2017), our findings newly suggest that blastemal progenitors direct immune support and that this bi-directional signaling is important for early stages of regeneration.

74 Failure to retain myeloid cells during the transition from wound healing to blastema formation may account for the regenerative delay in il-8 morphants. Monocyte-derived macrophages are necessary for successful appendage regeneration in the axolotl and other model systems, playing roles including directing blastemal outgrowth, clearing senescent cells, and modulating blastemal cell proliferation (Li et al., 2012; Godwin et al.,

2013; Petrie et al., 2014; Yun et al., 2015; Nguyen-Chi et al., 2017; Simkin et al., 2017b).

Others have demonstrated the importance of pro-regenerative (M2) rather than pro- inflammatory (M1) macrophage subtypes in regeneration (Pei et al., 2016; Simkin et al.,

2017b). Therefore, it is possible that IL-8 may retain M2 macrophages during the initiation of blastema formation. The extent of the role of granulocytes in regeneration, however, seems to be more context dependent (Nakayama et al., 2011; Li et al., 2012; Kurimoto et al., 2013; Paris et al., 2016; Lindborg et al., 2017). Granulocytes, including neutrophils, clear debris during wound healing (Wang, 2018). Therefore, low levels of granulocytes in il-8 morphants may have slowed wound healing, delaying regeneration. Furthermore, in the zebrafish, IL-8 can act as a potent neutrophil chemoattractant (de Oliveira et al., 2013; de Oliveira et al., 2016) or a chemokinetic molecule, aiding in neutrophil reverse migration away from sites of injury through Cxcr2 signaling (Powell et al., 2017). In contrast, our findings suggest that IL-8 may retain granulocytes including neutrophils during late wound healing rather than facilitate their exit strategy as in the zebrafish, highlighting potential species-specific differences in immune responses during regeneration. As IL-8 morphant limbs eventually regenerate, it is clear that other pathways compensate to ensure wound healing resolution.

75 Lastly, we show that CXCR-1/2 signaling is necessary during early, but not late, stages of limb regeneration. CXCR-1/2 inhibitor-treated limbs beginning during early and late stages of wound healing failed to form a blastema. The high level of myeloid cell death, which was not observed in il-8 morphants, during early regeneration was likely the main cause of failure to regenerate, suggesting that CXCR-1/2 signaling may serve as an important survival pathway for monocytes and granulocytes in early limb regeneration.

Additionally, the difference in severity of phenotypes between IL-8 morphant and CXCR-

1/2 inhibitor-treated limbs is likely attributed to the fact that CXCR-1/2 binds to other interleukins including IL-1 and IL-6 (Baggiolini et al., 1997). Therefore, it is possible that other cytokines are acting in concert with IL-8 to control CXCR-1/2 specific myeloid cell behaviors during early limb regeneration. Altogether, these data highly suggest that

CXCR-1/2 signaling may be key in bridging communication between early blastemal cells and the immune system during the initiation of regeneration.

3.6. Materials and Methods

Animal procedures

Axolotl (Ambystoma mexicanum) husbandry and surgeries were performed in accordance with the Association for Assessment and Accreditation of Laboratory Animal

Care (AAALAC) and Institutional Animal Care and Use Committee (IACUC) guidelines at

Harvard University. Juvenile white axolotls (5-8 cm) were used for all morpholino and overexpression experiments. For CXCR-1/2 inhibitor experiments, animals (3-5 cm) were immersed in either 500 nM SB-225002 (Tocris Bioscience) or DMSO (Sigma) beginning at either 0, 3, or 15 dpa and solutions were changed daily.

76 In situ hybridization and quantification

Tissue was collected at 0, 7 and 21 dpa and fixed in 4% paraformaldehyde overnight at 4C, washed in PBS, brought up a sucrose gradient to 30% sucrose, and embedded in OCT. The blocks were serially sectioned and 16 um sections were collected.

Custom RNAscope probes for the axolotl orthologs of cxcr-1/2, csf1r, il-8, kazd1, mpo, and prrx-1 were generated (Advanced Cell Diagnostics, Newark, CA) in either the C1 or

C2 channels. Double chromogenic section in situ hybridization was performed using the

RNAscope 2.5 HD Duplex Detection Kit (Advanced Cell Diagnostics) according to the manufacturer’s instructions on frozen cryosections.

The total number of prrx-1+only, il-8+ only, csf1r+ only or double positive cells were quantified by counting cells that exhibited 10 or greater puncta for either probe. These are categorized as strongly expressing cells according to RNAscope standards (score of

4+). These numbers were used to quantify the percentages of co-positive and single positive cells.

Generation and validation of the myc-tagged il-8 overexpression construct

To generate an il-8 overexpression construct, the open reading frame for il-8 was amplified out of cDNA from regenerating limbs at 7 dpa with primers that added on a NheI site and a kozak sequence directly upstream of the ATG start codon and a HindIII site directly downstream of the coding sequence. This PCR product was cloned into a pCMV-

TVA-T2A-tdTomato backbone (gift from J. Whited) in place of TVA. An empty pCMV-T2A- tdTomato vector was used as a control.

77 To validate the expression of the vector, 293T cells were transfected with 50 ug of either the il-8 overexpression or tdTomato control construct using Lipofectamine 2000.

The media and cells were separately collected at 48 hours post-transfection (hpt). Media was concentrated with a 15 mL Amicon filter (3K) (Millipore Sigma, Burlington, MA).

Protein was extracted from cells using Trizol (ThermoFisher Scientific). Western blotting was performed on both il-8 overexpression and control transfected cell lysates and media using a mouse monoclonal anti-GAPDH (MAB374, Millipore Sigma, 1:2000) and polyclonal rabbit anti-myc tag antibody (ab9106, Abcam, Cambridge, MA, 1:2000). Cells were additionally immunostained at 72 hpt for the myc tag (same antibody as above) and tdTomato using a polyclonal goat anti-tdTomato antibody (LS-C348313, LSBio, Seattle,

WA, 1:1000).

Ectopic expression of il-8 in intact limbs

Intact limbs of axolotls were injected and electroporated with myc tagged il-8 overexpression construct (1 ug/uL in PBS). At three days post-injection, animals were injected intraperitoneally with EdU (Life Technologies, Carlsbad, CA) at a concentration of 8 mg/kg four hours prior to tissue collection. Tissue was prepped and embedded in

OCT in a similar manner to previously described above. The blocks were serially sectioned at a thickness of 16 um. EdU staining was performed using the Click-it EdU

Alexa Fluor 488 Imaging Kit (ThermoFisher Scientific). For immunostaining, mouse anti- chick Pax7 (DSHB, University of Iowa, IA, 1:200) and rabbit anti-human CD34 (ab81289,

Abcam, 1:200) antibodies were used.

78 The imaging analyses were all conducted blinded. For the quantification of il-8 overexpression limbs, 3-4 sections were imaged per limb and quantified. Total numbers of EdU+, CD34+, pax7+, CD34+ EdU+, and pax7+ EdU+ cells were counted and percentages were calculated out of total either DAPI+ nuclei for total EdU percentages, pax7+ nuclei for dividing satellite cells, and CD34+ cells for dividing endothelium. Dividing bone/perichondrial or epidermal cells were counted based on EdU+ cells of the cell type out of DAPI total of the cell type (counted by morphology). A two-tailed paired t-test was used for statistical analysis.

NSE/NCAE Staining and Analysis of Myeloid Cells

Staining of monocytes and granulocytes was performed using the -Naphthyl

Acetate (Non-specific Esterase) (NSE) kit or Naphthol AS-D Chloroacetate (Specific

Esterase) (NCAE) Kit (Sigma-Aldrich), respectively, according to the manufacturer’s instructions. The only modification to the protocol was a 10-minute fixation step in citrate- acetone-methanol fixative. For the overexpression experiment, the area of each cross- section was measured using ImageJ analysis software. For characterization of the il-8 morphant limbs, tissue was collected and prepared at 1, 5, and 7 dpa. The area of the section 500 um from the amputation plane was measured and the quantification was performed blinded. Every effort was made to ensure quantification around the same area in all limbs using the humerus and ulna as landmarks. A two-tailed paired t-test was used for all statistical analyses and all of the quantification was performed blinded (averaging

2-3 sections/limb). For characterization of DMSO- or SB-225002-treated limbs, limbs from

79 animals treated from time of amputation were collected at 7 dpa, stained, and analyzed as described above.

IL-8 morpholino knockdown

Axolotls were anesthetized in 0.1% tricaine and intact forelimbs were injected and electroporated with either control (left limb) or il-8-targeted morpholino (right limb).

Approximately 3-4 uL of morpholinos were injected at a final concentration of 5 uM in

PBS. At two days post-injection, both forelimbs of axolotls were amputated at the mid- radius/ulna level and control or il-8 morpholinos were injected again at 3 days post- amputation into regenerating stump tissue. Both translation-blocking and five-point mutation control morpholino anti-sense oligonucleotides were designed and generated by GeneTools LLC (Philomath, OR) against the il-8 ORF with the following sequences:

Control: 5'-CCGATCTTGATGCTCACCTCCTG-3'

il-8: 5'-CCGATGTTCATGGTGACCTGCTG-3'

To validate morpholino knockdown, il-8-targeted and control morpholino injected limbs from the same animal were collected and protein was extracted using Trizol

(ThermoFisher Scientific). Western blotting was performed on protein extracts using the anti-GAPDH antibody described above as well as a cross-reacting polyclonal mouse anti- chick IL-8 antibody (MBS2018201, MyBioSource, San Diego, CA, 1:400).

For analysis at the blastemal stages, animals were injected intraperitoneally with

EdU (Life Technologies) at a concentration of 8 mg/kg four hours prior to tissue collection.

Tissue was collected at 21 dpa and prepped as described above. Picro-mallory staining was performed on sections for histological analysis to analyze blastema length and area.

80 Cell proliferation was assessed using the Click-it EdU Alexa Fluor 488 Imaging Kit

(ThermoFisher Scientific). All imaging analysis was done in ImageJ and conducted blinded. A two-tailed paired t-test was used for statistical analysis.

TUNEL staining

TUNEL staining was performed using the In Situ Cell Death Detection Kit (Roche) as described previously (Zhu et al., 2012). The percentage of TUNEL+ nuclei (out of DAPI total) were quantified from 2 sections per limb and averaged.

Alcian blue staining

Alcian blue staining of DMSO or SB-225002-treated 40 dpa limbs was performed as describe previously (Whited et al., 2013).

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82 Kurimoto, T., Yin, Y., Habboub, G., Gilbert, H. Y., Li, Y., Nakao, S., Hafezi- Moghadam, A. and Benowitz, L. I. (2013). Neutrophils express oncomodulin and promote optic nerve regeneration. J Neurosci 33, 14816-14824.

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Miron, V. E., Boyd, A., Zhao, J. W., Yuen, T. J., Ruckh, J. M., Shadrach, J. L., van Wijngaarden, P., Wagers, A. J., Williams, A., Franklin, R. J. M., et al. (2013). M2 microglia and macrophages drive oligodendrocyte differentiation during CNS remyelination. Nat Neurosci 16, 1211-1218.

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85

Chapter 4

Wound epidermis-dependent transcriptional programs during early limb regeneration

4.1. Preface

The conceptualization and experimental design was performed in collaboration with Doug for the work presented in this chapter. I performed all of the experiments and the data analysis presented in this chapter and these data are part of a manuscript in progress.

4.2. Abstract

The wound epidermis is a transient epithelial structure that is required throughout limb regeneration. During blastema formation, the wound epidermis matures from a thin epidermal structure to a thick structure known as the apical epithelial cap (AEC) that is tightly coupled with the blastema. While the roles of the AEC have been more well- defined, the specific roles that the early wound epidermis plays in blastema formation remain largely unknown. In this chapter, we prevented wound epidermis formation by suturing full skin over the amputation plane and performed transcriptional profiling at 5 dpa of the stump-derived dividing and non-dividing cells, as well as the full thickness skin epithelial cells in order to investigate which transcriptional programs in the stump tissues are dependent on the wound epidermis, and which wound epithelial programs are potentially dependent on signaling from the stump tissues. Surprisingly, the overall transcriptional profile of early dividing progenitors was quite similar in the absence of the wound epidermis, while the profiles of the surrounding tissues diverged considerably.

Most of the differentially expressed transcripts in both the non-dividing stump tissues and full skin flap epithelial cells pertained to genes involved in inflammation, extracellular matrix (ECM) regulation and tissue histolysis, suggesting that the wound epidermis

87 modulates these processes. In addition, pathway analysis of differentially expressed transcripts in regenerating stump tissues revealed that dividing progenitors likely fail to activate key growth factor signaling pathways. In all, these data identify wound epidermis- dependent processes in early stages of regeneration and elucidate new potential regulators of the initiation of limb regeneration.

4.3. Introduction

Limb regeneration is an incredibly complicated process. One obvious layer of complexity lies in the cellular heterogeneity of the limb as it is comprised of many different tissues (muscle, bone, connective tissue etc.), which can be parsed into sub-tissue cell- types (Gerber et al., 2018; Leigh et al., 2018). Each of these cell types must execute tissue-specific regenerative programs in a coordinated manner spatiotemporally. To do so, cell-cell communication is key and layered on top of the cellular heterogeneity is the significance of ongoing cross-talk between all of the different regenerating tissues in the limb, which can be illustrated by notable examples including the nerve dependence of regeneration (Farkas and Monaghan, 2017) and the requirement of supportive cells/tissues including macrophages (Godwin et al., 2013) and the wound epidermis/AEC

(Goss, 1956; Thornton, 1957; Mescher, 1976). In addition, the necessity of nerves and macrophages differs depending on the stage of regeneration, illustrating temporal dynamics of communication between tissues. These added layers of complexity make it paramount to deconvolute the important roles that specific tissues may play during limb regeneration.

88 As mentioned in the introductory chapter, one of the requirements for successful limb regeneration is the formation of a thin wound epidermis during early wound healing stages and the subsequent maturation of this epithelial structure into the apical epithelial cap (AEC) (Campbell and Crews, 2008). While it is known that the AEC plays roles that mirror that of the AER in limb development, including maintaining blastemal cell proliferation and likely pattern formation (Boilly and Albert, 1990; Han et al., 2001), less is known about the roles of the early wound epidermis. Preventing early wound epidermis formation by suturing full thickness skin over the amputation plane prevents blastema formation (Mescher, 1976). Yet, early induction of proliferation in progenitors from many tissues still occurs despite the lack of a wound epidermis, suggesting that it does not play a major role in inducing cell cycle re-entry in regenerating limbs (Mescher, 1976; Johnson et al., 2018). Nevertheless, the early wound epidermis expresses signaling molecules including axMLP that are known to be sufficient to induce proliferation in axolotl tails and limbs (Sugiura et al., 2016), suggesting that it may still participate in inducing cell cycle re-entry in the limb.

Here, we investigated the roles of the early wound epidermis by performing transcriptional profiling of full skin flap sutured limbs. In conjunction with the transcriptional dataset generated in chapter 2, we compared the gene expression profiles of dividing cells and the surrounding tissues of regenerating limbs in normal regenerating and full skin flap sutured limbs at 5 dpa in order to identify 1) wound epidermis-dependent transcriptional programs in regenerating stump tissues and 2) transcriptional programs in the wound epidermis that are potentially dependent on signals from regenerating stump tissues.

89 4.4. Results

4.4.1. Full skin flap sutured limbs exhibit divergent transcriptional profiles from normal regenerating limbs

In order to investigate the role(s) of the early wound epidermis in limb regeneration, we wanted to examine the transcriptional differences between regenerating limbs in the presence and absence of the wound epidermis. We performed full skin flap surgeries immediately post-amputation to prevent wound epidermis formation (Figure 4.1)

(Mescher et al., 1976). As mentioned in the introductory chapter, suturing full thickness skin prevents migration of epithelial cells from the periphery over the exposed amputation plane, in essence preventing formation of the wound epidermis due to the presence of the dermal layer. Utilizing the protocol that we developed in chapter 2, we expanded our previous dataset on early regenerating limbs to include full skin flap sutured limbs.

To this end, we performed full skin flap surgeries immediately post-amputation and collected tissue at 5 dpa in order to isolate and transcriptionally profile three subpopulations: stump-derived dividing, stump-derived non-dividing, and epidermal cells

(Figure 4.2). Utilizing the previous dataset from regenerating limbs at 0 and 5 dpa, we compared the transcriptional profiles of all three populations in the presence and absence of the wound epidermis. Principal component analysis (PCA) of full skin flap and normal regenerating samples at 5 dpa revealed four clusters separated by tissue type (epidermal or non-epidermal tissues) and condition (full skin flap sutured or normal regenerating limbs) (Figure 4.3).

90 Figure 4.1. Suturing full thickness skin over the amputation plane prevents wound epidermis formation. Picro-mallory stained sections of normal regenerating and full skin flap sutured non-regenerating limbs at 8 dpa are shown in A and B. Higher magnification images of insets in A and B are shown in A’ and B’, respectively. A normal wound epidermis is formed by migration of peripheral epithelial cells (stained red) over the amputation plane and lacks a collagen-thick dermal layer (stained blue). The direct contact between the epithelial cells and the underlying stump tissues is thought to facilitate signaling that is essential for blastema formation. Suturing full thickness skin (epidermis and dermis) immediately over the amputation plane prevents the formation of a wound epidermis. The presence of the dermis is thought to be a physical barrier that prevents signaling between the epithelial cells and the regenerating stump tissues, inhibiting limb regeneration. dpa, days post-amputation.

91 Figure 4.2. Schematic of transcriptional profiling experiment in the presence and absence of the wound epidermis. We collected and performed RNA-sequencing on stump derived-dividing, stump derived-non-dividing, and full thickness skin cells from full skin flap sutured limbs at 5 dpa utilizing the method developed in Tsai et al. (2019). We then compared the transcriptional profiles of each subpopulation between normal regenerating and full skin flap sutured limbs to identify differentially expressed transcripts.

92

Figure 4.3. PCA analysis of normal and full skin flap sutured limb samples.

4.4.2. Early dividing cells fail to activate key signaling pathways in the absence of the wound epidermis

As determined previously in Chapter 2, HIPPO, canonical Wnt, and TGF- signaling were the only growth factor signaling pathways predicted to be active within normal regenerating stump tissues, while most other pathways appeared to be repressed.

In order to investigate whether absence of the wound epidermis impacted the activation of these pathways, we similarly utilized Ingenuity Pathway Analysis (IPA) to examine the potential activation/suppression of growth factor signaling pathways in stump-derived dividing and non-dividing cells of full skin flap sutured limbs.

This analysis revealed that similar growth factor signaling pathways (HIPPO and canonical Wnt) were predicted to be active in non-dividing cells in both conditions,

93 suggesting that the activation of these pathways is independent of the wound epidermis.

In contrast, the absence of the wound epidermis led to failed activation of HIPPO, canonical Wnt, and TGF- signaling in early dividing cells (Figure 4.4A). Analysis of the transcript levels of components of all three pathways in dividing cells revealed that many components of TGF- signaling (e.g. inhba, ltbp1, smad2, smad5, pai1, snai1, tgfr1), canonical Wnt signaling (e.g. lef1, dkk1, pard3, dvl2, ppard), and HIPPO signaling (e.g. mst1, lats1, tead3) are indeed down-regulated, further suggesting that these pathways are likely inactive (Figure 4.4B). Notably, our previous data suggested that early blastemal progenitors selectively establish an autocrine TGF- signaling network during early stages of limb regeneration based on the enrichment and induction of many TGF- signaling components in early dividing cells. While many of these components are down- regulated in full skin flap sutured limbs, some remain induced to similar levels including the upstream regulator tgfb1 and a downstream target twist1, suggesting that the induction of these genes in early progenitors is wound epidermis-independent. In all, these data suggest that the wound epidermis may play a role in the induction of these pathways in early blastemal progenitors.

The differential activation of growth factor signaling pathways in early dividing cells was quite surprising given that the overall transcriptional profiles in the absence of the wound epidermis were relatively similar. Only 596 transcripts were differentially expressed in dividing cells and of these, 313 of these transcripts were annotated. Some of the most highly differentially expressed transcripts are shown in Figure 4.4C and the top 100 transcripts can be found in Table A.3.1. Intriguingly, the top most differentially expressed hit was an RNA binding protein ews which plays a role in transcriptional

94 Figure 4.4. Early dividing cells exhibit minor differences in transcriptional profiles in the absence of the wound epidermis. (A) Ingenuity pathway analysis (IPA) of differentially expressed transcripts in stump-derived dividing and non-dividing cells suggests that early dividing cells lose predicted activation of Hippo, canonical Wnt, and TGF- signaling pathways. (B) Heatmap of normalized and averaged TPM values of differentially expressed transcripts associated with Hippo, canonical Wnt, and TGF- signaling at 0 dpa, 5 dpa, and 5 dpa full skin flap sutured limbs. (C) Heatmap of normalized TPM levels of select differentially expressed transcripts in early dividing cells between both conditions. WE, wound epidermis; dpa, days post-amputation; FSF, full skin flap.

95 regulation and has been most well-studied in tumorigenic contexts when partially fused with other transcriptional activators as a result of aberrant chromosomal translocations

(Lee et al., 2018). In addition, blastemal progenitor-enriched transcripts discovered in the previous regenerating dataset including lin41, mmp13, and k2c8 (Tsai et al., 2019) were down-regulated in full skin flap sutured limbs, suggesting that the expression of these genes is wound epidermis-dependent. Altogether, these data suggest that while the wound epidermis does likely play a role in inducing the activation of key pathways in early dividing progenitors, it does not seem to strongly impact the overall transcriptional profiles of these cells.

4.4.3. Non-dividing cells in regenerating stump tissues display aberrant inflammation and extracellular matrix (ECM) regulation profiles

While the dividing cells displayed similar transcriptional profiles in the absence of the wound epidermis, the non-dividing cells in regenerating stump tissues exhibited divergent profiles in full skin flap sutured limbs from their regenerating counterparts. In total, 3,911 transcripts were differentially expressed and of these, 982 were annotated. A of the top 100 differentially expressed transcripts can be found in Table A.3.2 of the

Appendix. GO analysis of enriched biological processes of differentially expressed transcripts revealed an overwhelming overrepresentation of transcripts involved in immunomodulation, inflammation, and ECM organization (Figure 4.5A). Furthermore, in conjunction with the expression data from non-regenerating intact limbs in our previous dataset, we could parse the differentially expressed transcripts into four clusters: transcripts that either fail to be maintained (cluster 1), fail to be repressed (cluster 2), fail

96 to be induced (cluster 3), or are aberrantly induced in non-dividing cells (cluster 4, Figure

4.5B).

Further analysis of the enriched biological processes in specific clusters revealed that transcripts involved in ECM regulation were overrepresented in cluster 1, suggesting that many ECM components and regulators including fbn2, finc (or fn1), mmp2, mmp14, has2, and comp failed to maintain expression at some level in full skin flap sutured limbs

(Figure 4.5E). In addition, differentially expressed ECM-related transcripts were also present in the other clusters. Notably, transcripts associated with the important pro- regenerative ECM component tenascin (tena) (Godwin et al., 2014), along with ECM- degrading enzymes including catk, mmp14, mmp2, and mmp13 failed to be induced in the absence of the wound epidermis. Interestingly, co1a1, which is a major collagen matrix component of differentiated bone and cartilage (Viguet-Carrin et al., 2006) seemed to aberrantly maintain expression in full skin flap sutured limbs. Coupling this observation with the lack of induction of the bone matrix-degrading enzyme catk (Troen, 2004), these data suggest that normal bone resorption that occurs during early stages of limb regeneration may be inhibited in the absence of the wound epidermis. In all, these data support the notion that the wound epidermis plays a major role in ECM regulation and tissue histolysis during early stages of regeneration.

Furthermore, immune-related transcripts were highly overrepresented in cluster 3

(Figure 4.5C) and a subset of the most highly differentially expressed transcripts are represented in Figure 4.5D. Many of these transcripts were associated with components of pathogen-associated molecular pathways including toll-like receptor (TLR) and interferon // signaling (e.g. myd88, tlr3, tlr7, stat1, irf3, ifih5, mx1, mx2, il18), while

97 Figure 4.5. Inflammation and ECM regulation in stump-derived non-dividing cells are heavily affected in the absence of the wound epidermis. (A) Plot of enriched GO biological processes from all differentially expressed transcripts in non-dividing cells in full skin flap sutured limbs. (B) Clustered heatmap of the normalized TPM values of all differentially transcripts in non-dividing cells between regenerating and full skin flap sutured limbs at 5 dpa. Four main clusters numbered 1 to 4 (color coded pink, orange, green, and purple were apparent. (C) Cluster-specific enriched GO biological processes from cluster 1 (top, pink) and 3 (bottom, green). (D-F) Heatmap of normalized TPM values

98 Figure 4.5. (continued) of differentially expressed transcripts involved in inflammation (D), ECM regulation (E), or transcriptional regulation. dpa, days post-amputation; FSF, full skin flap; FDR, false discovery rate; ECM, extracellular matrix.

mypop, an anti-viral transcriptional regulator (Wustenhagen et al., 2018), was over-

expressed in full skin flap sutured limbs. In addition, the macrophage marker mpeg1

(Zakrzewska et al., 2010) was down-regulated in the absence of the wound epidermis,

potentially indicating less macrophage infiltration into the damaged tissue. Finally,

components of the anti-inflammatory PPAR/RXR signaling pathway (e.g. gpda, rras,

adcy7, m4k4, mp2k2) and anxa1, a well-known anti-inflammatory molecule that promotes

wound healing resolution (Sugimoto et al., 2016), were down-regulated. These data

collectively suggest that inflammatory processes are mis-regulated in the absence of the

wound epidermis, and the failure of induction of many immunomodulatory transcripts

suggests that the early wound epidermis plays a large role in directing the inflammatory

response.

Finally, we were able to identify transcription factors and regulators that were

differentially expressed in the absence of the wound epidermis (Figure 4.5F).

Transcription factors including myc, fosl2, stat1, sp4, and prdm1 as well as epigenetic

regulators including setb1 and smyd3 failed to be induced in non-dividing cells of full skin

flap sutured limbs. In addition, transcription factors including nkx2.5, cbx5, klf10, and

gat1b were aberrantly expressed, indicating that the early wound epidermis may play a

role in inducing and/or suppressing these factors.

99 4.4.4. Epithelial cells in Full thickness skin exhibit dysregulation of transcripts involved in cell proliferation, inflammation, and ECM regulation

Communication between damaged stump tissues and the migratory epithelial cells that form the wound epidermis is essential for proper blastema formation. Full skin flap surgeries prevent this communication due to the presence of the collagen-thick dermal layer. Therefore, we compared the transcriptional profiles between epithelial cells from the full thickness skin flap and the wound epidermis in order to determine which transcripts were dysregulated. These transcripts could provide valuable insight into processes within the wound epidermis that may be dependent on signaling from regenerating stump tissues during early stages of regeneration.

Overall, 1,060 transcripts were differentially expressed between full skin flap epithelial cells and the wound epidermis (Figure 4.6). Of these, 480 were annotated and are shown in Figure 4.6A. A list of the top 100 transcripts can be found in Table A.3.3 of the Appendix. Clustering of the differentially expressed transcripts revealed three main clusters: transcripts that failed to be induced (Cluster 1), were aberrantly expressed

(Cluster 2), or failed to be maintained (Cluster 3, Figure 4.6A). Cluster-specific analysis of overrepresented biological processes and pathways demonstrated that many of the differentially expressed transcripts in Cluster 1 were involved in cell proliferation (e.g. cell cycle checkpoint, spindle organization, DNA conformation change), RNA processing, and pro-inflammatory processes (production of or response to type I interferon and cytokine stimulus) (Figure 4.6B). In addition, transcripts involved in the anti-fungal immunity C-type lectin receptor signaling (e.g. irf1, nalp3) were overrepresented in Cluster 3.

100

Figure 4.6. Wound epidermis and full skin flap sutured skin exhibit transcriptional differences in inflammation, ECM regulation, and cell proliferation. (A) Clustered heatmap of the normalized TPM levels all differentially expressed transcripts between normal wound epidermal and full skin flap epidermal cells at 5 dpa. Three main clusters of transcripts (color coded pink, green, and purple) are apparent. (B) Plot of cluster- specific enriched GO biological processes and pathways for cluster 1 (top) and cluster 3 (bottom). (C-F) Heatmap of differentially expressed transcripts that are immunomodulators (C), ECM regulators (D), transcriptional regulators (E), and cell surface receptors (F). WE, wound epidermis; FSF, full skin flap; FDR, false discovery rate; ECM, extracellular matrix.

101 Notably, similar to the stump-derived non-dividing cells in full skin flap sutured limbs, inflammatory processes were also dysregulated in the full skin flap epithelium. A subset of the highest differentially expressed immune-related transcripts are shown in

Figure 4.6C. Transcripts involved in interferon signaling (e.g. in35, ifi6, ifih1, irf3, stat1, stat6, tri27, tri56) and cytokine signaling (e.g. ccl4, ccl28) were not induced in full skin flap epithelial cells. Anti-pathogenic mucins including muc5a and muc5b were also aberrantly overexpressed. Intriguingly, mpeg1 and anxa1 were also down-regulated in full skin flap epithelial cells similar to stump-derived non-dividing cells. Altogether, these data indicate that pro- and anti-inflammatory processes are mis-regulated in the absence of signaling between the epithelial cells and regenerating stump tissues.

In addition to inflammation, many ECM components were differentially expressed, indicating differences in ECM regulation in the absence of signaling from underlying stump tissues (Figure 4.6D). Transcripts corresponding to structural components of intact skin that maintain expression in the wound epidermis (e.g. otoan, k1c15, k1c19, k2c5, gpc1, and gpc6) lost expression in the full skin flap epithelia. Moreover, strongly expressed secreted molecules involved in ECM regulation (e.g. co7a1, finc, and palld, mmp3) were dysregulated.

Finally, in order to determine potential upstream drivers of these transcriptional differences, we identified differentially expressed transcription factors and receptors

(Figure 4.6E-F). Several transcription factors (e.g. elf3, zn236, smbt2, hmgb2, htf4, foxp4) and transcriptional regulators (e.g. smyd3) failed to be induced in full skin flap sutured epithelial cells, while others (e.g. ctnd1, grsf1, klf11) failed to be maintained or were overexpressed (e.g. gat1b, brd7, tcf7). Furthermore, many of the differentially expressed

102 receptors were normally expressed in intact skin and wound epidermis, but failed to maintain expression in the full skin flap (e.g. ndrg2, jag1, avr2a, tsn15, notc3, tsn3, lifr).

Interestingly, fgfr3 expression was ectopically maintained in full skin flap epithelial cells, and tm100 was overexpressed. These data provide a list of potential candidates that may help us gain insight into wound epidermis biology.

4.5. Discussion

The wound epidermis is essential for proper limb regeneration (Thornton, 1957;

Mescher, 1976) and exists as a transient epithelial structure that eventually differentiates into the epidermis of the regenerated limb (Kragl et al., 2009). Throughout regeneration, the wound epidermis matures from an early thin epithelium during wound healing to the

AEC once the blastema has formed. While more is known about the roles of the AEC during later stages of regeneration (Campbell and Crews, 2008), little is known about the functional significance of the early wound epidermis. Here, we investigated how preventing wound epidermis formation would impact the transcriptional programs of three distinct subpopulations: stump-derived dividing cells (enriching for blastemal progenitors), stump-derived non-dividing cells, and epithelial cells. By suturing full thickness skin over the amputation plane, we simultaneously inhibited formation of the wound epidermis and prevented signaling between the epithelial cells and the underlying stump tissues, allowing us to both examine programs in the regenerating stump that are wound epidermis-dependent and programs in the epidermis that likely require signaling from underlying stump tissues.

103 Surprisingly, our data suggests that transcriptional programs of early blastemal progenitors are largely wound epidermis-independent, as the overall transcriptional profiles of the early dividing cells were not strongly impacted by inhibiting wound epidermis formation. Given that cell-cycle re-entry of progenitors in regenerating limb tissues occurs in the absence of the wound epidermis (Mescher, 1976; Johnson et al.,

2018), our data adds transcriptional evidence to further suggest that cell cycle re-entry is likely directly stimulated by processes inherent to inflicting injury such as blood clotting

(Tanaka and Brockes, 1998; Tanaka et al., 1999). However, dividing progenitors appear to fail to activate the only growth factor signaling pathways that are active in normal regenerative contexts (HIPPO, canonical Wnt, and TGF- signaling), suggesting that the wound epidermis may play a targeted role in directing the cellular behaviors of progenitors after cell-cycle re-entry (ex. EMT transition, cell migration). In addition, it is interesting to note that all three, rather than individual, pathways failed to be activated. As previously mentioned in Chapter 2, these three pathways share many downstream effectors (ex. smad2, smad5 etc.) and synergize in various processes including tumorigenesis (Attisano and Wrana, 2013). Therefore, our results lend further evidence to suggest that these three pathways are likely functionally interconnected and important for early blastemal cell biology. Determining the specific importance of early activation of these pathways in blastemal progenitors in future studies will help us gain insight into the molecular mechanisms that control behaviors of blastemal cells during early stages of limb regeneration.

While preventing wound epidermis formation did not seem to have a strong impact on the overall transcriptional profiles of early dividing cells, the surrounding tissues were

104 more heavily affected. In particular, major inflammatory and ECM regulatory processes were affected in non-dividing regenerating stump tissues and the full skin flap epidermis.

Components of both pro- and anti-inflammatory processes including interferon //, TLR, and PPAR/RXR signaling either failed to be induced or were aberrantly overexpressed in full skin flap sutured limbs, suggesting that both the onset and resolution of inflammation are dysregulated. Interestingly, transcripts corresponding to a key mediator of wound healing resolution anxa1 and the macrophage marker mpeg1 were commonly down-regulated in both populations. Anxa1 is known to play a major role in promoting apoptosis and engulfment of neutrophils by macrophages, a key step in wound healing resolution which decreases production of pro-inflammatory cytokines and stimulates production of anti-inflammatory cytokines in mammalian wound healing (Sugimoto et al.,

2016). In addition, anxa1 can promote monocyte recruitment and therefore impact macrophage infiltration. Therefore, common down-regulation of anxa1 and mpeg1 further suggests that anti-inflammatory processes may not be occurring properly.

Additionally, transcripts corresponding to ECM regulators and components were also dysregulated in both non-dividing stump tissues and full skin flap epithelial cells.

Among these, one of the most highly differentially expressed transcripts corresponded to a collagen-degrading enzyme catk that failed to be induced in non-dividing cells. Catk is highly expressed in osteoclasts, bone-resorbing cells that play a major part in tissue histolysis during early stages of regeneration (Fischman and Hay, 1962; Troen, 2004).

Furthermore, the bone matrix marker co1a1 aberrantly maintains expression and a transcription factor that modulates osteoclast biology, fosl2 (Bozec et al., 2008), is down- regulated in full skin flap sutured limbs. These results suggest that bone resorption during

105 early stages of regeneration is not occurring properly in regenerating stump tissues in the absence of the wound epidermis and in turn, that the wound epidermis plays roles in directing bone resorption. Furthermore, many ECM components failed to be induced

(k1c24, co7a1, and palld) or maintained (otoan, k2c5, k1c15, k1c19) in full skin flap epidermis, suggesting that signaling from regenerating stump tissues may directly influence the expression of these genes. In all, these results suggest both that the wound epidermis plays a major role in immunomodulation and ECM regulation and that ongoing communication between the wound epidermis and underlying stump tissues is crucial for these processes.

Most importantly, we identified differentially expressed potential upstream

(secreted molecules and receptors) and downstream (transcription factors and regulators) modulators that may play a role in controlling or repressing either wound epidermis-dependent transcriptional programs and/or wound epidermis function and maturation. Intriguingly, the histone methyltransferase smyd3 was commonly induced in stump-derived non-dividing cells and the wound epidermis, but down-regulated in full skin flap sutured limbs. As smyd3 has known roles in both development and tumorigenesis

(Du et al., 2014; Giakountis et al., 2017), our data suggest that it may also be an important common epigenetic regulator during limb regeneration. Furthermore, while some of these candidate transcription factors have known roles in development and immunomodulation

(e.g. prdm1, myc, gat1b, nkx2.5, klf10), others are uncharacterized in function (e.g. znfx1, zn236). These data therefore provide a rich resource of new potential regulators of regeneration for the field and characterizing the functional roles of these candidates will help elucidate the molecular mechanisms of limb regeneration.

106 4.6. Materials and Methods

4.6.1. Animal care and husbandry

Axolotl (Ambystoma mexicanum) husbandry and surgeries were performed in accordance with the Association for Assessment and Accreditation of Laboratory Animal

Care (AAALAC) and Institutional Animal Care and Use Committee (IACUC) guidelines at

Harvard University. Sub-adult white and albino axolotls (15-18 cm) provided by the

Ambystoma Genetic Stock Center (AGSC, University of Kentucky, KY) were used for the

RNA-sequencing experiment. Amputations were performed as previously described in

Tsai et al. (2019) and full skin flap surgeries were performed on all four limbs of nine animals as detailed in (Mescher, 1976) immediately post-amputation.

4.6.2. RNA isolation and sequencing

Approximately 2-3 mm of tissue directly proximal to the amputation plane was collected at 5 dpa. Twelve limbs were pooled for each biological replicate per timepoint and the experiment was performed in triplicate. Dissociation, cell cycle analysis, and RNA isolation from the samples was performed as described in Tsai et al. (2019).

4.6.3. Differential Expression Analysis

Raw reads were filtered and trimmed using Trimmomatic (Bolger et al., 2014) and aligned to the axolotl transcriptome (Bryant et al., 2017) using Kallisto (Bray et al., 2016).

Raw TPM values from each sample were analyzed using DESeq2 (Love et al., 2014).

Differentially expressed transcripts were filtered for fold change > 2 and adjusted p-value

< 0.05.

107 For pathway analysis, Uniprot IDs of blastx hits from different species for all differentially expressed transcripts were converted to human Uniprot IDs using HUGO

Gene Nomenclature Committee (HGNC) and/or Uniprot ID conversion tools online.

Ingenuity pathway analysis (IPA) analysis was performed on differentially expressed transcripts to determine growth factor signaling pathway activation or inhibition for the following comparisons: 0dpa 2N vs 5 dpa 2N, 0 dpa 4N vs 5 dpa 4N, 0 dpa epi vs 5 dpa wound epidermis (WE), 0dpa 2N vs 5 dpa FSF 2N, 0 dpa 4N vs 5 dpa FSF 4N, and 0 dpa epi vs 5dpa FSF epi. Heatmaps of normalized TPM levels for differentially expressed transcripts in each fraction (2N, 4N, or epidermis) were generated in RStudio utilizing the

ComplexHeatmap package (Gu et al., 2016) or the gplots package (Gregory R. Warnes,

2015). Clusters of differentially expressed transcripts (5 dpa regenerating vs. 5 dpa FSF) for each cellular fraction were analyzed using WebGestalt (Wang et al., 2017).

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111

Chapter 5

Midkine (mk) regulates the wound epidermis-to-AEC transition

5.1. Preface

The work presented in this chapter is part of ongoing work for a manuscript in preparation. All of the experimental design was performed in collaboration with Doug and

Clara Baselga-Garriga. I performed and analyzed all of the in situ hybridizations, the initial iMDK experiment, genotyping and initial characterization of the mk mutants, and the mk overexpression experiment. The rest of the experiments including further characterization of the iMDK phenotype and generating/characterizing the mk mutant phenotype were performed in collaboration with Clara.

5.2. Abstract

Midkine (mk) is a pleiotropic cytokine involved in many processes including development, inflammation, and tumorigenesis. In this chapter, we investigated the role of mk in limb regeneration which exhibited decreased expression in the absence of the wound epidermis. We demonstrate that mk is strongly expressed throughout regeneration in the basal layers of the wound epidermis/AEC and in early connective tissue blastemal progenitors. Chemical and genetic perturbation of mk impairs wound epidermis maturation, which either completely inhibits or delays regeneration. Sequencing of mk inhibitor (iMDK) treated limbs further reveals dysregulation of wound epidermis gene expression and persistent inflammatory profiles. Finally, overexpression of mk in regenerating limbs leads to uncontrolled growth of the wound epidermis. Altogether, these results suggest that mk is one of the first identified regulators of early wound epidermis maturation to the AEC.

113 5.3. Introduction

Midkine (mk) is a pleiotropic growth factor cytokine that modulates a variety of biological processes including development, inflammation, tumorigenesis, and regeneration (Muramatsu, 2010; Sorrelle et al., 2017). Most notably, in recent decades mk has been shown to be a critical mediator of tumor metastasis and growth (Karaman and Alitalo, 2017; Olmeda et al., 2017), chronic inflammation (Sorrelle et al., 2017), and neural regeneration (Calinescu et al., 2009; Gramage et al., 2015). Many of these processes parallel with those that occur during early stages of limb regeneration.

Nevertheless, its potential role in limb regeneration has yet to be described. Interestingly, we noticed that mk expression was up-regulated in early stages of normal regenerating limbs and down-regulated in non-regenerating full skin flap sutured limbs, suggesting that it may play a role in limb regeneration. We therefore decided to examine the function of mk in limb regeneration through a series of gain- and loss-of-function approaches.

Altogether, our data suggest that mk is a mediator of wound epidermis maturation.

5.4. Results

5.4.1. Midkine is strongly expressed in the basal layers of the wound epidermis and connective tissue blastemal progenitors

We initially observed that mk appeared to be down-regulated in all three subpopulations in full skin flap sutured limbs (Figure 5.1), suggesting that it may be important for regeneration. To determine whether mk was indeed expressed during limb regeneration, we performed timecourse RNAscope in situ hybridization of mk transcripts during regeneration. In situ hybridization revealed that mk was lowly expressed in cells

114

150 2N 4N * Epidermis

100

M

P T 50 ** * ** 0

0 dpa 5 dpa 5 dpa Full Skin Flap Figure 5.1. Normalized transcripts per million (TPM) levels of midkine (mk) in regenerating and full skin flap conditions. At 0 dpa, mk is lowly expressed in both non- dividing and dividing cells within stump tissues, but not expressed in homeostatic skin. Upon amputation, mk expression stays the same in stump-derived non-dividing cells and increases in dividing cells. Furthermore, mk expression is strongly induced within the wound epidermis at 5 dpa. In full skin flap conditions, we observed lower expression of mk in all three populations.

115

Figure 5.2. Mk is strongly expressed in the basal layers of the wound epidermis/AEC and connective tissue blastemal progenitors. (A-F’) Timecourse RNAscope in situ hybridization of mk transcripts at 0, 1, 3, 7, 14, and 21 dpa. Higher 20x magnification insets in A-F are shown in A’-F’, respectively. Arrowhead in C’ denotes the beginning of the wound epidermis. Dotted lines demarcate the wound epidermis/stump tissue boundary in B’-D’ and the amputation plane in E-F. (G) In situ hybridization of mk in full skin flap sutured limbs at 7 dpa. (H) Percentages out of total number of cells counted of mk+ only, prrx-1+ only, or mk+/prrx-1+ co-positive cells at 5 (n=281 cells total) and 7 dpa (n=424 cells total). (I-J) Double in situ hybridization of prrx-1 and mk at 5 and 7 dpa. Images taken at 40x magnification. Arrowheads in I-J denote co-positive cells. Scalebars are 500 µm in A-F and F, 100 µm in A’-F’, and 50 µm in I-J. FSF, full skin flap; dpa, days post-amputation.

116 that are interspersed throughout the muscle in non-regenerating intact limbs (0 dpa) and up-regulated upon amputation beginning at 3 dpa (Figure 5.2A-F’). Specifically, mk is expressed within the basal layers of the wound epidermis and mesenchymal cells within regenerating stump tissues at 3 and 7 dpa (Figure 5.2C-D’). By early and late bud blastema stages (Figure 5.2E-F’), mk remains strongly expressed in the AEC and at lower levels within the blastemal cells. Furthermore, we confirmed that mk expression remains at low levels comparable to that of uninjured limbs in full skin flap sutured limbs (Figure

5.2G).

To determine whether mk was expressed within blastemal progenitors of regenerating stump tissues, we performed double in situ hybridization of mk with the connective tissue blastemal progenitor marker prrx-1 (Gerber et al., 2018). We observed co-expression of mk and prrx-1 transcripts at both 5 and 7 dpa (Figure 5.2H-J).

Quantification of the total number of mk and prrx-1 expressing cells revealed that 87.88% and 67.11% of prrx-1+ cells were co-positive, while 67.70% and 85.28% of mk+ cells were prrx-1+ at 5 and 7 dpa, respectively. These data suggest that mk is expressed in a subset of connective tissue blastemal progenitors. However, single-positive mk+ cells were present at sizeable percentages, indicating that mk may be expressed in blastemal progenitors from other lineages, or other cell types within the regenerating limb. To examine whether other cell types such as immune cells may express mk, we performed double in situ hybridization of mk and csf1r, a monocyte marker (Grayfer et al., 2014;

Stanley and Chitu, 2014), and found little overlap between mk and csf1r transcripts. Only

3.73% of csf1r+ cells were mk+ at 5 dpa (Figure A.4.1), suggesting that mk is not highly expressed in monocytes. Altogether, these data suggest that mk is highly expressed

117

Figure 5.3. MK protein localization during limb regeneration. (A-C’) Immunostaining of axolotl MK protein at 0 (A-A’), 5 (B-B’) and 14 dpa (C-C’). (D-D’) Immunostaining of axolotl MK protein in full skin flap sutured limbs at 7 dpa. Arrowheads denote the amputation plane in A-D. Higher 20x magnification images of the epidermis A-D are shown in A’-D’. Dotted lines demarcate full skin flap in D of the wound epidermis/stump boundary in A’-D’. Scalebars are 500 µm in A-D and 100 µm in A’-D’. FSF, full skin flap; dpa, days post-amputation.

118 throughout regeneration in the basal layers of the wound epidermis and connective tissue blastemal progenitors.

We additionally analyzed the MK protein expression pattern by performing immunostaining with a custom polyclonal antibody against the C-terminus of axolotl MK.

Prior to immunostaining, we validated the specificity of the antibody on regenerating limb extracts at 10 dpa with and without blocking peptide (detailed in Methods, Figure A.4.2).

Immunostaining of MK at 0, 5, and 14 dpa revealed that the protein expression pattern mirrors that of the transcripts (Figure 5.3). In particular, MK protein is absent in non- regenerating skin, and induced in the basal layers of the epidermis at 5 dpa. By 14 dpa,

MK protein is found throughout the AEC and in the blastema, suggesting that MK is diffusing outwards from basal layers of the AEC (Figure 5.3A-C’). Moreover, MK protein levels in full skin flap sutured limbs at 7 dpa are comparable to those of non-regenerating limbs. In all, these data suggest that mk is highly up-regulated during limb regeneration.

5.4.2. Chemical inhibition of mk inhibits limb regeneration

In order to determine whether mk may play a role in limb regeneration, we performed drug treatments with a small molecule inhibitor of MK, iMDK (Masui et al.,

2016) during limb regeneration (Figure 5.4). Treatment with iMDK beginning from 0 dpa or 7 dpa completely prevented blastema formation and limb regeneration (Figure 5.4A,

B). In addition, alcian staining of DMSO or iMDK-treated limbs at 60 dpa confirmed that all skeletal structures regenerated normally in DMSO controls, in contrast to iMDK-treated limbs, which regenerated none (Figure 5.4C). These data collectively suggest that mk may play an important role in limb regeneration.

119

Figure 5.4. Chemical inhibition of mk prevents limb regeneration. (A) Schematic of inhibitor experiments with iMDK. Limbs were amputated and animals were immersed in 10 uM iMDK beginning at either 0 or 7 dpa. (B) Brightfield images of DMSO- and iMDK- treated limbs at 26 and 40 dpa reveals that iMDK treatment inhibits limb regeneration (n=4 DMSO and iMDK for each treatment, 4/4 did not regenerate in each iMDK treatment). The green dotted line denotes the amputation plane. (C) Alcian staining of DMSO- and iMDK-treated limbs at 60 dpa. Scale bars are 500 µm in B and 1 mm in C-D.

120 5.4.3. iMDK-treatment impairs wound epidermis maturation

We next wanted to determine which cellular processes were affected by iMDK treatment. We reasoned that it was possible that treatment with iMDK inhibited limb regeneration by lowering levels of cellular proliferation. In order to address this question, we examined levels of cellular proliferation in DMSO and iMDK-treated limbs at 5 dpa and 14 dpa. We observed no differences in the percentage of EdU+ cells at 5 dpa (24.04% vs 23.27% EdU+ cells) (Figure 5.5A-B), suggesting that iMDK-treatment does not affect early induction of proliferation. Interestingly, we noticed that EdU+ cells still accumulate at the amputation plane in iMDK-treated limbs at 14 dpa, yet a blastema does not form.

Furthermore, histological staining of iMDK-treated limbs demonstrated that fibroblastic blastema-like cells accumulate at the amputation plane even up to 40 dpa, however regeneration remains inhibited (Figure 5.5C). These data collectively suggest that iMDK treatment does not affect cellular proliferation, but may affect the development of other structures necessary for blastema formation, such as the AEC.

To this end, we further examined picro-mallory stained sections from DMSO- or iMDK-treated limbs at 14 dpa. While DMSO-treated limbs displayed a large blastema with a thick AEC, iMDK-treated limbs had an abnormally thin wound epidermis (Figure 5.6A-

B’), suggestive of a defect in the transition from the early wound epidermis to the AEC. In addition, TUNEL staining revealed that iMDK-treated limbs exhibited higher levels of total cell death (5.451% vs. 1.801% TUNEL+ nuclei, pval=0.0145) (Figure 5.6C-E). Intriguingly, we noticed that many TUNEL+ cells were present in the wound epidermis in iMDK-treated limbs and quantification revealed a significant increase in the percentage of TUNEL+ nuclei specifically within the wound epidermis (7.110% vs. 4.384% TUNEL+ nuclei,

121

Figure 5.5. iMDK treatment does not affect early induction of proliferation in limb regeneration. (A) Representative images of EdU-stained regenerating limbs at 5 and 14 dpa from either DMSO- or iMDK-treated limbs. EdU staining is shown in green. Arrowheads denote the amputation plane. (B) Quantification of the percentage of EdU+ cells in DMSO- and iMDK-treated limbs revealed no difference (24.04% vs 23.27% EdU+ cells, pval= ns; DMSO, n=6, iMDK, n=7). (C) Picro-mallory staining of iMDK-treated limbs at 40 dpa. Scale bars in A are 200 µm and C are 500 µm. dpa, days post-amputation.

122

Figure 5.6. iMDK-treated limbs exhibit an abnormally thin wound epidermis and increased levels of cell death. (A-B’) Picro-mallory stained images of DMSO- and iMDK- treated limbs at 14 dpa. Higher magnification images of insets in A and B are shown in A’-B’ respectively. Dotted line denotes the amputation plane. (C-D) Representative images of TUNEL-stained sections from DMSO and iMDK-treated limbs. (E) Quantification of total and wound-epidermis specific TUNEL+ nuclei revealed increased levels of apoptosis both overall (5.451% vs. 1.801% TUNEL+ nuclei, *pval=0.0145) and specifically within the wound epidermis (7.110% vs. 4.384% TUNEL+ nuclei, *pval=0.0232) of iMDK-treated limbs (n=6 for both DMSO and iMDK treatments). Scalebars are 200 µm in A-B and C-D, 100 µm in A’-B’.

123 pval=0.0232) (Figure 5.6E). In all, these data collectively suggest that the defect in wound epidermis development may be due at least in part, to an increase in cell death.

5.4.4. iMDK-treated limbs display persistant inflammatory profiles and dysregulated wound epidermis gene expression

In order to determine how iMDK treatment affected the gene expression programs of regenerating limbs, we transcriptionally profiled DMSO- and iMDK-treated limbs at early stages of blastema formation (11 dpa). Principal component analysis (PCA) of all samples revealed that most of the variation in the dataset was due to the treatment

(Figure A.4.3). Overall, 1,403 transcripts (1,011 annotated) were differentially expressed

(Figure 5.7A). Of the annotated transcripts, 575 were enriched in iMDK-treated limbs and

436 were enriched in DMSO control limbs (Table A.4.1).

Ingenuity pathway analysis (IPA) of enriched transcripts in iMDK-treated limbs revealed an enrichment of transcripts in many pro-inflammatory pathways including toll- like receptor (TLR) signaling, Th1/2 activation, and Oncostatin M signaling (Figure 5.7B,

Table A.4.2, Table A.4.3). Furthermore, transcripts involved in leukocyte extravasation and infiltration of granulocytes, monocytes, and lymphocytes into tissues were also enriched. These data indicate that iMDK-treated limbs exhibit a persistent inflammatory response, and suggest that wound healing resolution is likely defective in these limbs.

Examination of the enriched transcripts in DMSO-treated limbs revealed that many of these transcripts were involved in actin cytoskeletal regulation including RhoA and

RhoGDI signaling (Figure 5.7C). These pathways are heavily involved in contexts of cell migration and invasion (in the case of tumorigenesis) (Newell-Litwa and Horwitz, 2011;

124 Lawson and Ridley, 2018), suggesting that iMDK-treated limbs display potential defects in cell migration. In addition, many DMSO-enriched transcripts pertained to the cell cycle and amino acid metabolism, which was less surprising given that the DMSO-treated limbs were developing blastemas, and likely had higher numbers of highly metabolic dividing cells.

Interestingly, some of the top most differentially expressed transcripts from the analysis were genes that are known to be expressed in the wound epidermis including urom (Campbell et al., 2011; Knapp et al., 2013). Utilizing our dataset as well as published single cell data from regenerating limbs (Gerber et al., 2018; Leigh et al., 2018), we mined the data for transcripts that are known to be expressed specifically in skin during homeostasis or in the wound epidermis during regeneration. Our findings revealed that many wound epidermis genes were mis-expressed in iMDK-treated limbs (Figure 5.7D).

In particular, wound epidermis genes including s10aa, s10ad, agr2, chs1, natt4, and k1c17 were overexpressed in the inhibitor treatment, while three genes m1ip1, elf3, and ptprz were down-regulated. These transcriptional data coupled with our histological data strongly suggest that iMDK treatment impairs wound epidermis development.

Finally, to corroborate the persistent inflammatory transcriptional profile with additional experimental evidence, we examined the levels of monocytes in DMSO- and iMDK-treated limbs at 5 and 14 dpa (Figure 5.8). NSE staining revealed higher levels of monocytes at both 5 and 14 dpa (Figure 5.8A-D). Furthermore, quantification of NSE+ cells at 5 dpa confirmed an early significant increase in monocyte density in iMDK-treated limbs (440.0 vs. 234.6 NSE+ cells/mm2, pval = 0.0168) (Figure 5.8E). In all, these data provide supportive evidence that treatment with iMDK leads to a persistent inflammation.

125

Figure 5.7. Treatment with iMDK leads to dysregulation of wound epidermis gene expression and persistent inflammation. (A) Heatmap of annotated differentially expressed transcripts in DMSO- and iMDK-treated limbs reveals two main clusters (colored pink and orange) of transcripts either enriched in DMSO or iMDK treatments. Transcript expression was normalized per row and plotted as a Z-score. (B) Plot of enriched pathways in iMDK-treated limbs. (C) Plot of enriched pathways in DMSO-treated limbs. (D) Heatmap of mis-regulated wound epidermis genes in DMSO and iMDK-treated limbs. Transcript expression is plotted as a normalized Z-score.

126

Figure 5.8. iMDK-treated limbs exhibit higher levels of monocytes. (A-D) NSE staining of DMSO- and iMDK-treated limbs at 5 dpa (A-B’) and 14 dpa (C-D). Higher magnification images of insets in A and B are shown in A’ and B’, respectively. Arrowheads denote NSE+ monocytes in A’ and B’. Dotted lines in C-D demarcate the amputation plane. (E) Quantification of NSE+ monocytes at 5 dpa revealed a significant increase in the density of monocytes in iMDK-treated limbs (440.0 vs. 234.6 NSE+ cells/mm2, *pval = 0.0168; n=7, DMSO, n=6, iMDK). Scale bars are 200 µm in A-B and 50 µm in A’-B’. Dpa, days post-amputation.

127 5.4.5. Midkine F0 mutants have delayed blastemal outgrowth and impaired wound epidermis maturation

While iMDK treatment yielded a strong phenotype, the mechanism of the drug remains unknown (Masui et al., 2016) and we therefore wanted to perturb mk in a more precise manner genetically. To achieve this, we utilized CRISPR technology and generated mk F0 knockout animals by targeting the start codon (Figure A.4.4A). The slow development of the axolotl allows for high penetrance of mutations throughout the animal, allowing for phenotyping to occur at the F0 stage (Fei et al., 2018). Genotyping of limbs from these animals via deep-sequencing of PCR amplicons from the mk locus revealed that many of these mutants were mosaic null, with no wild-type alleles present (Figure

A.4.4B). Mk F0 mutant animals developed normally and had no obvious phenotypic defects. Furthermore, immunostaining of limbs from tracrRNA non-targeted controls and mk F0 mutants confirmed a complete loss of staining in these null mutants (Figure 5.9A).

We next wanted to determine whether genetic loss of mk similarly impaired regeneration. Brightfield imaging of limbs from tracrRNA controls and mk F0 mutants revealed that mk mutants exhibited delayed blastema formation, but eventually regenerate normally (Figure 5.9C-D). We measured the blastema lengths from sections of regenerating limbs of tracrRNA controls and mk F0 mutants at 14 dpa and observed that the delay in blastema formation segregated based on genotype, with mk null animals

(no detectable wildtype allele) exhibiting the most severe delay (152.06 µm in mk null mutants vs. 404.8 µm in tracrRNA controls, pval = 0.0043) (Figure 5.9E-H). These data collectively suggest that blastema formation is impaired during early stages of regeneration.

128

Figure 5.9. Mk-deficient mutants exhibit delayed blastema formation. (A-B) Immunostaining of regenerating limbs at 7 dpa of both tracrRNA controls (A) and mk mutants (B) mk antibody reveals an absence of staining in the mk mutant. (C-D) Brightfield images of regenerating limbs at 11 dpa of tracrRNA controls or mk F0 mutants reveals a delay in blastema formation. White dotted line denotes the amputation plane. (E) Graph of measurements of blastema length at 14 dpa from regenerating tracrRNA controls, mk null, mk mutants with low % of wildtype alleles, and mk mutants with high % of wildtype alleles reveals a dose dependence effect of mk on the phenotypic delay. (F- G) Representative histological stains from sections of either mk null, mk high% wildtype, and tracrRNA controls at 14 dpa. Scale bars are 500 µm in A-B, 1 mm in C-D, and 200 µm in F-G. wt, wildtype, dpa, days post-amputation.

129

Figure 5.10. Mk-deficient mutants exhibit an abnormally thin wound epidermis. (A) Representative histological stains of sections from regenerating limbs of tracrRNA controls and mk mutants at 10 dpa reveals mk mutants. (C) Quantification of wound epidermis thickness reveals a significant decrease in width of the wound epidermis of mk mutants (152.06 µm in mk null vs. 404.8 µm in tracrRNA control, **pval = 0.0043; n=8, DMSO and iMDK). Scale bars are 500 µm in in A-B.

Since chemical inhibition of mk with iMDK led to abnormal wound epidermis maturation, we decided to examine the wound epidermis of regenerating limbs of control and mk F0 mutant animals. As the phenotypic delay in blastema formation was most severe in mk null animals, we focused the rest of our analyses detailed below on animals with this genotype and refer to them heretofore as mk mutants. Histological staining of limbs from both controls and mk mutants at 10 dpa revealed a striking difference in the thickness of the wound epidermis (Figure 5.10). The wound epidermis of control regenerating limbs had thickened considerably as is natural when maturing into the AEC, whereas mk mutant limbs exhibited a thin wound epidermis. Quantification further revealed that this decrease in wound epidermis thickness was statistically significant

(53.64 µm vs. 115.7 µm, pval=0.0084).

130

Figure 5.11. Mk-deficient mutants display a mild increase in the levels of monocytes. (A-B) Representative NSE stained sections from regenerating limbs of tracrRNA controls and mk mutants. (C) Quantification of the density of NSE+ monocytes revealed a mild significant increase in the levels of monocytes in mk mutants (230.5 vs. 160.4 NSE+ cells/mm2, *pval=0.0441; n=8, DMSO and iMDK). Scale bars are 500 µm in in A-B. dpa, days post-amputation.

In addition, to examine whether there was an increase in the levels of monocytes similar to that observed in iMDK-treated limbs, we performed NSE staining on limbs at 10 dpa and quantified the number of NSE+ cells. Quantification indicated a mild significant increase in the density of monocytes in mk mutant regenerating limbs (230.5 vs. 160.4

NSE+ cells/mm2, pval=0.0441), suggesting there may also be a persistent inflammatory response (Figure 5.11). These results collectively corroborated the chemical inhibition experiments and therefore strongly suggest that loss of mk leads to impaired wound epidermis maturation and persistent inflammation.

5.4.6. Overexpression of mk during regeneration leads to uncontrolled wound epidermis growth

Because genetic loss and inhibition of mk affected the wound epidermis, we next wanted to determine whether overexpression of mk during regeneration would affect the

131 wound epidermis as well. To this end, we injected and electroporated either a tdTomato control (pCAG-tdTomato) and/or a mk overexpression construct (pCAG-MK) at 3 dpa and examined the effects on limb regeneration (Figure 5.12A, Figure A.4.5). Interestingly, while tdTomato control limbs exhibited nice blastemas by 16 dpa (Figure 5.12B-C), mk- overexpressing limbs exhibited protruding bone at 7 dpa coupled with abnormal wound epidermis masses (Figure 5.12D-E), suggesting both aberrant growth and tissue integrity of the wound epidermis. By 16 dpa, the wound epidermis continues to grow uncontrollably and histological staining of sections through tdTomato and mk-overexpressing limbs at

16 dpa confirmed that mk overexpression led to uncontrolled expansion of the wound epidermis (Figure 5.12F-H). Intriguingly, mk-overexpressing limbs also seemed to exhibit potential defects in bone resorption, as the bone structure still remains intact at the amputation plane (Figure 5.12G-H). We further noticed the presence of large cells invading the bone that are reminiscent of osteoclasts, the main bone cells responsible for resorption. These data collectively suggest that overexpression of mk leads to uncontrolled growth of the wound epidermis and potential defects in bone resorption.

5.4.7. Sdc-1 is expressed in the wound epidermis

Gain- and loss-of function perturbation experiments determined that one main role that mk plays during limb regeneration is modulating the development and maturation of the early wound epidermis to the AEC. In order to determine whether mk may directly regulate the wound epidermis, we examined whether any mk receptors (Xu et al., 2014) were up-regulated during limb regeneration in our transcriptional dataset. Of the many

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Figure 5.12. Overexpression of mk in regenerating limbs leads to uncontrolled growth of the wound epidermis. (A) Experimental schematic of the mk overexpression experiment. (B-E) Brightfield images of either tdtomato- or mk-overexpressing limbs at 7 and 16 dpa. Arrowhead in D denotes an abnormal mass of skin forming in mk- overexpressing limbs. Dotted line denotes the amputation plane. (F-H) Representative picro-mallory stained sections of tdtomato- or mk-overexpressing limbs at 16 dpa reveals an abnormally thick wound epidermis. Yellow arrowheads additionally denote the aberrant presence of osteoclasts in mk-overexpressing limbs. n=5 for tdT and MK overexpression limbs. Scale bars are 1 mm in B-E and 200 µm in F-H.

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Figure 5.13. Sdc-1 is expressed in normal skin and the wound epidermis during limb regeneration. Double RNAscope in situ hybridization of mk and sdc-1 at 0, 7, and 14 dpa. Insets show where the 40x magnification image was taken from in the original section. Dotted lines denote the boundary between the normal epidermis/wound epidermis and stump tissues. Scale bars are 50 µm.

receptors, one mk receptor, sdc-1, was the most highly expressed in homeostatic skin at

0 dpa and up-regulated in early dividing cells as well as the wound epidermis at 4 and 5 dpa within our dataset. To validate these transcriptional data, we performed time course

RNAscope double in situ hybridization of mk and sdc-1 at 0, 7, and 14 dpa (Figure 5.13).

As expected, sdc-1 was indeed expressed at low levels in homeostatic skin and up- regulated within the wound epidermis and mesenchymal cells within regenerating stump tissues at 7 and 14 dpa. Interestingly, mk was expressed strongly within the basal layers of the wound epidermis while sdc-1 was expressed more highly in the outer layers at 7 and 14 dpa, suggesting that secreted mk may directly modulate the wound epidermis.

Furthermore, sdc-1 expression overlapped with a subpopulation of mk+ cells within regenerating stump tissues at 7 dpa, however many single-positive sdc-1+ cells were also

134 present. By 14 dpa, sdc-1 expression persists within blastemal cells at strong levels.

Altogether, these data indicate that mk likely directly modulates the wound epidermis and the behavior of cells within regenerating stump tissues to regulate blastema formation.

5.5. Discussion

The wound epidermis/AEC is essential for formation of the blastema, yet little is known about molecules that regulate the maturation of the early wound epidermis to the

AEC during blastema formation. Here, we have identified a novel regulator of wound epidermis maturation. Genetic loss and chemical inhibition of mk both lead to an abnormally thin wound epidermis, indicative of impaired wound epidermis maturation. In addition, iMDK-treated limbs display aberrant wound epidermis gene expression profiles and overexpression of mk during limb regeneration is sufficient to induce uncontrolled wound epidermis growth. Finally, the midkine receptor sdc-1 is expressed within outer layers of the wound epidermis during limb regeneration, suggesting that mk directly modulates the wound epidermis. Altogether, these data suggest that mk plays a central role in modulating the wound epidermis.

It is possible that mk is regulating wound epidermis maturation by acting as a survival factor. One key step to maturation of the wound epidermis is growth in thickness, which is generally due to proliferation and migration of cells from the peripheral skin (Hay and Fischman, 1961; Campbell and Crews, 2008). Inhibitor-treated limbs have higher levels of cell death specifically in the wound epidermis, providing one likely explanation as to why both inhibitor-treated limbs and mk mutants are unable to form an AEC correctly. In fact, mk has been shown to act as a survival factor in other contexts including

135 neural crest cell migration in development (Vieceli and Bronner, 2018) and current experiments are underway to address whether a similar increase in cell death is observed in mk mutants as well. Furthermore, mk overexpression during regeneration resulted in aberrant wound epidermis growth, suggesting that mk may be sufficient to induce epithelial cell proliferation. Therefore, it is possible that mk modulates wound epidermis maturation by balancing cell death and proliferation rates in the wound epidermis.

Notably, wound epidermis genes were dysregulated in inhibitor-treated limbs.

Surprisingly, many of these genes were found to be normally expressed in the wound epidermis (ex. urom, s10ad, s10aa, k1c17 and epcam) as detected in publicly available datasets (Gerber et al., 2018; Leigh et al., 2018; Tsai et al., 2019), and highly overexpressed in iMDK-treated limbs. Notably, chs1 was overexpressed in iMDK-treated limbs. Chs1 is expressed in the wound epidermis specifically during wound healing stages of regeneration (Leigh et al., 2018). Therefore, overexpression of chs1 in inhibitor-treated limbs further suggests that the wound epidermis fails to transition to the AEC. Finally, the midkine receptor ptprz (Xu et al., 2014) failed to be induced in the presence of the inhibitor, suggesting that ptprz may be a wound epidermis-specific receptor mediating the effects of mk in addition to sdc-1. Experiments are currently underway to determine the expression pattern of ptprz and whether small molecule inhibition of ptprz can phenocopy mk inhibition.

Finally, while the strongest phenotype we observed was impaired wound epidermis maturation, it is likely that mk plays more than one functional role during limb regeneration given its widespread expression in connective tissue blastemal progenitors and other as of yet unidentified cell types within regenerating stump tissues. Furthermore, the receptor

136 sdc-1 is also expressed in many cells within regenerating stump tissues as well, lending further evidence to a pleiotropic role for mk in limb regeneration. In addition to a defect in wound epidermis development, inhibitor-treated limbs and mk mutant limbs both have higher levels of monocytes. Moreover, inhibitor-treated limbs display persistent inflammatory profiles and overexpress many transcripts associated with pro-inflammatory pathways that are triggered in response to pathogen and tissue damage-related cues.

These results suggest the possibility that mk may directly modulate the immune response.

As the mk receptor sdc-1 is expressed in many cells within regenerating stump tissues that are not mk+, mk may likely also act as a paracrine immunomodulator as it is known to be involved in regulating inflammation (Weckbach et al., 2011). Another explanation for the observed persistent inflammation is that this is a secondary effect of impaired wound epidermis maturation. As our full skin flap transcriptional data in chapter 4 suggests that the early wound epidermis plays heavy roles in coordinating inflammation, it is conceivable that keeping the wound epidermis in a perpetually early state would impede the resolution of wound healing, leading to persistent inflammation. Finally, we observed higher levels of osteoclasts and aberrant tissue histolysis of the bone in mk- overexpressing limbs. Furthermore, we also observed that catk, a collagen-degrading enzyme expressed by osteoclasts (Troen, 2004) was down-regulated in iMDK-treated limbs as well. Others have shown that mk plays a large role in skeletal remodeling and bone resorption (Liedert et al., 2014), and therefore it is likely that mk may also play a role in bone resorption as well. In all, these data suggest that mk may play pleiotropic roles within limb regeneration and regulate many other aspects of blastema formation.

137 5.6. Materials and Methods

Animal care and husbandry

Axolotl (Ambystoma mexicanum) husbandry and surgeries were performed in accordance with the Association for Assessment and Accreditation of Laboratory Animal

Care (AAALAC) and Institutional Animal Care and Use Committee (IACUC) guidelines at

Harvard University. Juvenile white axolotls (3-5 cm animals) were utilized for the midkine inhibitor (iMDK) experiments and mk overexpression experiments. For drug experiments, limbs of animals were amputated and the animals were immersed in DMSO or 10 uM iMDK solution beginning at either 0 or 7 dpa. Drug solution was changed daily for the duration of the experiments and animals were fed normally.

Imaging analysis

All imaging analysis was conducted randomized and blinded. For most analyses, we utilized a two-tailed unpaired t-test to test for statistical significance unless otherwise stated.

In situ hybridization

Limb tissue was collected and processed as described above. Custom RNAscope probes were generated either the C1 or C2 channel for csf1r, mk, prrx-1, and sdc-1. In situ hybridization was performed using the RNAscope 2.5HD Duplex Detection Kit according to the manufacterer’s instructions. For quantification of single mk+, csf1r+, prrx-

1+, double positive mk+/prrx-1+ and double mk+/csf1r+ cells, we only counted strongly

138 positive cells (greater than 10 puncta per cell). These numbers were utilized to calculate the percentage of single and co-positive cells.

RNA-sequencing and analysis of iMDK-treated limbs

We isolated RNA from DMSO or iMDK-treated limbs at 11 dpa. For the RNA isolation, we homogenized the tissue in Trizol and purified RNA using the Qiagen RNeasy mini kit. We collected the samples in triplicate for each condition. Quality and quantity of

RNA was assessed using the Agilent RNA 6000 pico kit and run on a BioAnalyzer 2100

(Agilent). Sequencing libraries were then synthesized and sequenced on a Nextseq 500 at the Biopolymers Facility at Harvard Medical School.

Fastq data from the RNA-sequencing was filtered and trimmed using Trimmomatic

(Bolger et al., 2014). The data was then aligned to the previously published axolotl transcriptome (Bryant et al., 2017) and analyzed with DESeq2 (Love et al., 2014). For the differential expression analysis, we set the cutoff at an adjusted p-value < 0.05 and fold change > 2. K-means clustering of the transcripts was performed using the Complex

Heatmap package (Gu et al., 2016) and pathway analysis of DMSO- or iMDK-enriched transcripts was performed on differentially expressed transcripts using Ingenuity Pathway

Analysis (IPA) (Qiagen) software.

-Naphthyl Acetate Esterase (NSE) staining

NSE staining was performed using the -Naphthyl Acetate Esterase staining kit

(Sigma) according to the manufacturer’s instructions.

139 TUNEL staining

TUNEL staining was performed according to the manufacturer’s instructions and as described in (Zhu et al., 2012).

Generation of mk mutants and genotyping

In order to generate the mk mutants, we utilized CRISPR-cas9 technology and targeted the start codon of the mk locus with a single guide RNA (sgRNA) (ATG start is bolded): 5’AAGCCCCCACAACTGCATCC -3’. This sequence was also a unique within the axolotl genome. The mk gRNA and generic tracrRNA were ordered from IDT and reconstituted according to their instructions. We injected embryos at the one-cell stage with cas9 ribonucleoprotein (cas9 RNP) complexes. To generate the cas9 RNP complexes, we annealed the sgRNA and tracrRNA as described in Bryant et al. (2017).

We then mixed the mk gRNA (final concentration 200-400 ng/uL) with concentrated cas9

(final concentration 500 ng/uL) (PNA Bio Inc.) and incubated for 10 minutes at 37ºC to generate the cas9 ribonucleoprotein (RNP) complex. For each fertilized egg, we injected

2-3 nl of injection mix and performed the injections as described in (Khattak et al., 2014).

To genotype the animals, we collected genomic DNA from limbs of regenerating tracrRNA controls and mk mutants. Tissue was incubated in 50 mM NaOH for 20 min. at

94ºC and neutralized with TE buffer pH 7.5. The supernatant was then PCR purified using the Qiagen PCR purification kit according to the manufacturer’s instructions. We then performed genomic PCR to amplify the area surrounding the cut site with the following primers:

For 5’-TTGCTTATTCCTTGTGATCATGC-3

140 Rev 5’- GGCACATTATTACACAGAAAGCTC-3’

Next, we performed two rounds of additional PCRs to generate DNA sequencing libraries as described in (Gagnon et al., 2014). One round of nested PCR was performed using the following gene specific primers with universal overhangs:

For 5’- tctttccctacacgacgctcttccgatctGAGGTTTGATTGGACCCTGA-3’

Rev 5’- tggagttcagacgtgtgctcttccgatctGGCACATTATTACACAGAAAGCTC-3’

The next round of PCR was performed to add on i7 indices and P5/P7 sequences to generate DNA sequencing libraries. We pooled up to 24 libraries together and sequenced the pool on a department owned Miseq using the Miseq reagent nano kit v2 (300 cycle)

(Illumina). Fastq data was then analyzed utilizing CRISPResso (Pinello et al., 2016). For the analysis, mutants in which we detected no wildtype alleles were deemed null, with up to 10-15% wildtype alleles deemed low % wildtype, and greater than 15% deemed high

% wildtype (most of these animals were actually closer to 80-90% wildtype).

Characterization of mk mutants

Brightfield images of regenerating limbs were taken of tracrRNA controls and mk mutants during regeneration using an Olympus SZX16 dissecting microscope. Tissue was collected at 10 and 14 dpa for analyses. For the blastema length measurement analysis, we performed picro-mallory staining on sections and measured the length at 14 dpa. A similar analysis was performed when measuring the wound epidermis thickness.

Immunostaining and EdU staining

141 Limb tissue was collected and fixed overnight in 4% paraformaldehyde, brought up a sucrose gradient, and embedded in OCT. Tissue was cryosectioned at a thickness of 12 m. For EdU analysis, animals were injected as described in Tsai et al. (2019) and the Click-it EdU A488 Imaging Kit was utilized (Thermofisher Scientific) according to the manufacturer’s instructions. For MK immunostaining, we first generated a custom polyclonal rabbit antibody against the C-terminus (amino acids 125-142) of axolotl MK and validated it with a peptide blocking assay. We performed western blots on 20 ug of

10 dpa protein extracts and blocked with increasing concentrations of the peptide used to generate the antibody and saw a depletion of signal from the anti-MK staining (Figure

A.4.2).

Overexpression of mk in regenerating limbs

To generate the mk overexpression vector, we cloned in the open reading frame of mk by excising out tdTomato from the pCAG-tdTomato backbone using EcoRI and NotI

(Addgene plasmid #83029) to generate the pCAG-MK overexpression vector. To validate the expression and secretion of mk, we transfected 293T cells with either pCAG-tdTomato or pCAG-MK vectors and collected the transfected cells and media at 72 hours post- transfection (hpt). Protein lysates from the transfected 293T cells were collected using

RIPA buffer and western blots were performed on each of the protein lysates and media samples using anti-tdTomato (1:1000) (LS Bio LS-C340696), our custom generated anti-

MK (1:2000), and anti-GAPDH (1:2000) (Millipore AB2302) antibodies. The presence of axolotl mk was detected in both pCAG-MK transfected 293T cell lysates and media

(Figure A.4.5).

142 Limbs of regenerating animals were injected and electroporated at 3 dpa with either the pCAG-MK overexpression vector and/or the pCAG-tdTomato vector. The electroporation settings were the same as previously described in Tsai et al. (2019).

Approximately 1 ug total was injected into each limb. Brightfield images of regenerating limbs were taken at 7 and 16 dpa using an Olympus SZX16 dissecting microscope.

Alcian staining

Alcian blue staining of DMSO or iMDK-treated limbs at 60 dpa was performed as described previously (Whited et al., 2013).

5.7. References

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Campbell, L. J., Suarez-Castillo, E. C., Ortiz-Zuazaga, H., Knapp, D., Tanaka, E. M. and Crews, C. M. (2011). Gene expression profile of the regeneration epithelium during axolotl limb regeneration. Dev Dyn 240, 1826-1840.

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Gramage, E., D'Cruz, T., Taylor, S., Thummel, R. and Hitchcock, P. F. (2015). Midkine-a protein localization in the developing and adult retina of the zebrafish and its function during photoreceptor regeneration. PLoS One 10, e0121789.

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144 Lawson, C. D. and Ridley, A. J. (2018). Rho GTPase signaling complexes in cell migration and invasion. J Cell Biol 217, 447-457.

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Chapter 6

Conclusions and future perspectives

6.1. Revealing mechanisms of blastema formation is essential to understanding limb regeneration

Salamander limb regeneration is an incredibly complex phenomenon that has fascinated biologists for centuries (Spallanzani, 1769). Upon limb amputation, salamanders are remarkably capable of re-initiating limb developmental programs, unlike most mammals, which initiate fibrotic scarring responses. Why salamanders can accomplish such a feat in contrast to humans is therefore a question that has driven regenerative biologists for many years.

Successful limb regeneration hinges on the formation of a transient dedifferentiated cellular structure distal to the amputation plane known as the blastema, which later re-differentiates and patterns into mature tissues of the regenerated limb. The blastema is comprised of lineage-restricted progenitors derived from many different tissues within the limb including muscle, bone, and connective tissue (Kragl et al., 2009;

Gerber et al., 2018; Leigh et al., 2018) and therefore blastema formation requires precise coordination across a variety of cell-types both through space and time. Naturally, the blastema itself has been likened to the limb bud during development through its morphological resemblance. In fact, studies have shown that the blastema transcriptionally resembles the limb bud during later stages of regeneration (Knapp et al.,

2013; Gerber et al., 2018) and re-activates similar pathways necessary for limb development (Torok et al., 1999; Han et al., 2001; Christensen et al., 2002; Nacu et al.,

2016). Furthermore, the apical epithelial cap (AEC) that is tightly coupled with the blastema has been likened to the apical ectodermal ridge (AER) in limb development

148

based on similar gene expression patterns and roles in maintaining cell proliferation

(Boilly and Albert, 1990; Han et al., 2001; Christensen et al., 2002).

While later stages of regeneration post-blastema formation seem to resemble limb development, earlier stages during wound healing and the initiation of blastema formation display highly divergent transcriptional profiles (Knapp et al., 2013). The beginning stages of limb regeneration are exceptionally intricate involving many concurrent processes including the onset and resolution of inflammation, ECM degradation and tissue histolysis, wound epidermis formation, as well as the induction of proliferation and migration of different lineage-derived progenitors to the amputation plane (Kawasumi et al., 2013).

Therefore, understanding how each of these processes is controlled and coordinated molecularly will help us gain further insight into the mechanisms that drive blastema formation. To this end, many advances have been made in the field to delineate tissues and cell-types that are necessary for blastema formation during early stages. Specifically, it has been shown that the presence of nerves, the wound epidermis, and macrophages are each required for successful blastema formation (Thornton, 1957; Mescher, 1976;

Godwin et al., 2013; Farkas and Monaghan, 2017). Of these, the influence of early innervation has been the most well-studied and to date, several important neurotrophic molecules have been described (Kumar et al., 2007; Farkas et al., 2016). Yet, much remains to be discovered about the molecular regulation of immune cells and the wound epidermis during regeneration. Altogether, these studies make it imperative to further dissect the transcriptional programs of the initial stages of regeneration.

Building off of the important work of others in the field, we have made several important contributions to the limb regeneration field in the work presented in this

149

dissertation. Briefly, using the method that we developed to enrich for dividing cells during early stages of regeneration, we separately identified the gene expression programs that are active within blastemal progenitors and the surrounding tissues (non-dividing cells in regenerating stump tissues and the wound epidermis) (Tsai et al., 2019). We further leveraged our dataset to ask how these transcriptional programs change in the absence of the wound epidermis. These datasets collectively provided insight into early blastemal cell biology as well as the influence of the early wound epidermis. Additionally, these datasets are rich resources of potentially novel regulators of limb regeneration for the field to investigate. Utilizing these datasets, we have elucidated the roles of two cytokines, interleukin-8 (il-8) and midkine (mk), during blastema formation.

6.2. Early transcriptional programs of blastemal progenitors

One of the significant contributions we have made is elucidating the early transcriptional programs of blastemal progenitors in bulk irrespective of lineage during limb regeneration in the presence and absence of the wound epidermis. During normal regeneration, we revealed that early dividing progenitors likely selectively establish an autocrine TGF- signaling network, which is further supported by the work of others in the field that have shown that TGF- signaling is necessary for blastema formation and regeneration (Levesque et al., 2007; Denis et al., 2016). Blastemal progenitors appear to also activate both HIPPO and canonical Wnt signaling in conjunction with TGF- signaling, suggesting these three pathways may synergize to orchestrate global blastemal cell behaviors such as epithelial-to-mesenchymal transition (EMT)-related processes. When we prevent wound epidermis formation, we demonstrate that the

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activation of not one, but all three of these pathways is lost, suggesting both that these pathways are functionally interlinked and that the wound epidermis plays a role in activating these pathways in dividing progenitors. As these pathways synergize in tumorigenesis (Attisano and Labbe, 2004; Attisano and Wrana, 2013), it is likely that similar mechanisms are at play during early stages of regeneration.

As these data were generated from bulk sequencing datasets, it is also possible that different progenitors activate various combinations of these pathways, which would not be discerned from the resolution of our transcriptional dataset. Therefore, it will be key to validate which cell-types specifically activate these pathways in future work. With the release of the genome, it is also now possible to perform targeted knock-ins to potentially generate transgenic reporters for the activation of each of these pathways.

These animals could help further delineate the upstream regulatory mechanisms that regulate these pathways within limb regeneration and could be an exciting avenue to identify new signaling molecules that initiate progenitor activation/cell-cycle re-entry.

Despite the difference in pathway activation profiles in the presence and absence of the wound epidermis, we observed that dividing cells in both contexts had overall relatively similar transcriptional profiles. These data suggest that the wound epidermis does not heavily influence the overall gene expression programs of progenitors. Indeed, several studies have described the induction of proliferation in the absence of the wound epidermis (Mescher, 1976; Johnson et al., 2018), and others have suggested that proliferation is primarily induced in response to injury-related cues such as blood clotting

(Tanaka and Brockes, 1998; Tanaka et al., 1999). Our research therefore lends further transcriptional evidence to support that the wound epidermis is likely not necessary for

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directing the initial cues for progenitor cell activation, i.e. cell cycle re-entry, in limb regeneration. However, the wound epidermis may be essential for directing targeted responses including activating certain signaling pathways that may direct later behaviors including migration and sustained cycling.

6.3. Inflammation, ECM regulation, and tissue histolysis are likely regulated by the wound epidermis

Despite the importance of the wound epidermis, little is known about its functional role(s) during early stages of limb regeneration (Thornton, 1957; Mescher, 1976).

Transcriptional profiling of dividing progenitors and the surrounding tissues in limbs where we prevented wound epidermis formation by suturing full thickness skin over the amputation plane suggested that the early wound epidermis likely plays large roles in regulating inflammation, ECM regulation, and tissue histolysis. Transcripts involved in both pro- and anti-inflammatory pathways were heavily dysregulated in limbs where we prevented wound epidermis formation, suggesting that the wound epidermis may regulate both the onset and resolution of wound healing during the first stages of limb regeneration.

Furthermore, many transcripts involved in ECM regulation and tissue histolysis were also differentially expressed. Notably, we noticed that several genes involved in osteoclast differentiation and function including catk and fosl2 were amongst the most down-regulated transcripts. This was inherently interesting since osteoclasts are important bone-matrix degrading cells that are heavily involved in skeletal remodeling and bone resorption (Troen, 2004), processes that occur during the earliest stages of limb regeneration. Therefore, down-regulation of these genes coupled with the higher

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expression of the bone matrix marker co1a1 in full skin flap sutured limbs highly suggests that the wound epidermis may play a role in directing bone resorption.

6.4. Blastemal progenitors as early immunomodulators

The identification of blastemal progenitor-enriched transcripts provided us with the opportunity to learn about new aspects of early blastemal cell biology. An interesting observation that we made from our dataset was the expression of several inflammatory cytokines in blastemal progenitors, suggesting that early progenitors may play a role in immunomodulation. Indeed, further examination of the functional role of one enriched inflammatory cytokine il-8 through a series of gain- and loss- of function approaches revealed that il-8 is a connective tissue blastemal progenitor-derived molecule that modulates myeloid cell retention during the transition from wound healing to blastema formation. We further demonstrate that signaling through its receptor, cxcr-1/2, during early stages of regeneration is necessary for blastema formation. While others have nicely shown that myeloid cells including macrophages are necessary for limb regeneration (Godwin et al., 2013), our results newly suggest that bi-directional signaling from the blastemal progenitors is equally important. It is therefore likely that the continual communication between the cells that give rise to the blastema and supportive immune cells is crucial for building a permissive blastemal niche.

These results open up several intriguing lines of questioning for future studies.

Many have described that immune cells play important roles in promoting growth and healing in the contexts of tissue homeostasis, regeneration, and tumorigenesis (Aurora and Olson, 2014; Williams et al., 2016; Godwin et al., 2017b; Naik et al., 2018). Yet,

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others have described that immune cells play roles in promoting fibrotic scarring responses as opposed to regeneration (Lupher and Gallatin, 2006). It has therefore come to light in the last decade or so, that the response of the immune system is highly dependent on the micro-environment (Williams et al., 2016; Naik et al., 2018). Virtually all injuries in different organisms are immediately followed by the activation of common pro- inflammatory pathways from the immune system. The major differences lie in the response of the immune cells during wound healing resolution. Will they promote regeneration? Or will they promote fibrosis? Since immune cells are integral for blastema formation in a variety of natural organ and appendage regenerative contexts beyond just salamander limb regeneration (Aurora et al., 2014; Petrie et al., 2014; Godwin et al.,

2017a; Simkin et al., 2017a; Simkin et al., 2017b), it is conceivable that intrinsic differences may exist in the way that tissues from regenerating organisms respond to injury that in turn direct pro-regenerative immune responses. It would therefore be interesting to examine whether il-8 is similarly expressed in blastemal progenitors of regenerating organisms including zebrafish or the African spiny mouse and determine whether it may also direct the behaviors of cells within the immune system. Furthermore, comparing the inflammatory transcriptional profiles during late stages of wound healing between non-regenerative and regenerative organisms may provide further insights into inherent inflammation-related differences in the response of non-immune tissues to injury.

Finally, many questions remain about the overall regulation of early cxcr-1/2 signaling during early stages of limbs regeneration. Treatment with a cxcr-1/2 inhibitor completely prevented blastema formation, suggesting that the mechanisms modulating this pathway play an important role in regeneration. As CXCR-1/2 is known to bind many

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other inflammatory cytokines including IL-6 (Russo et al., 2014), it would be interesting to determine whether functionally perturbing these other ligands similarly affects limb regeneration. Additionally, future experiments identifying the transcriptional differences as a result of inhibition of CXCR-1/2 signaling will help us gain further insight into the down-stream processes that are affected in blastema formation.

6.5. Regulators of early wound epidermis to AEC maturation

The wound epidermis is a transient structure that matures from an early thin epithelial structure to the thicker AEC during blastema formation. While many have described the morphological steps occurring during this process (Campbell and Crews,

2008), little is known about the molecular regulation of wound epidermis maturation during regeneration. Our gain- and loss-of-function studies have collectively identified the pleiotropic growth factor cytokine mk as a regulator of wound epidermis maturation from the early stages to the AEC. We demonstrate that inhibition or genetic loss of mk impairs wound epidermis maturation to the AEC while overexpression during regeneration leads to aberrant overgrowth of the wound epidermis. Furthermore, the mk receptor sdc-1 is up-regulated and expressed within the wound epidermis during limb regeneration, suggesting that mk may likely directly modulate the wound epidermis. Altogether, these results suggest that mk is a critical regulator of wound epidermis maturation during limb regeneration.

However, many questions remain regarding the exact processes of wound epidermis maturation that mk affects. Our data suggest that mk may act as a survival factor, as inhibitor-treated limbs have higher levels of cell death in the wound epidermis.

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Furthermore, overexpression of mk is sufficient to induce abnormal expansion of the wound epidermis. As such, it would be interesting to determine whether mk modulates the cell growth/death rates of the wound epidermis during regeneration. Ongoing experiments are underway determining both whether mk mutants also exhibit increased cell death and whether there are differences specifically in wound epidermis proliferation.

We additionally identified two mk receptors that are expressed in the wound epidermis, sdc-1 and ptprz, that may mediate mk signaling. We have currently only validated the expression of sdc-1, which is expressed within both the wound epidermis and regenerating stump tissues. Interestingly, ptprz is expressed specifically within the wound epidermis (Leigh et al., 2018) and was down-regulated in the mk inhibitor-treated limbs, suggesting that it may be a wound epidermis specific mediator of mk signaling.

Current experiments are now underway to also validate the expression of ptprz and determine whether functional perturbation of ptprz and/or sdc-1 can phenocopy that of inhibitor-treated limbs or mk mutants.

Finally, while the most severe phenotype we observed when perturbing mk was pertaining to the wound epidermis, it likely plays other roles in limb regeneration based on its expression in other cell types including connective tissue blastemal progenitors in the regenerating stump tissues. In addition, the mk receptor sdc-1 is also expressed in stump tissues, lending additional supportive evidence to this hypothesis. Given the persistent inflammatory response and higher density of monocytes observed in regenerating limbs of inhibitor-treated and mk mutant animals, it is possible that mk may directly or indirectly modulate immune cell responses. Additionally, mk may play a role in bone resorption as well during early stages of regeneration as we observed that mk-

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overexpressing limbs exhibited morphological defects in bone histolysis and osteoclast presence. These are all interesting additional prospective processes that mk may participate in and future experimentation will help to delineate the potential significance of these roles in limb regeneration.

6.6. New frontiers and future perspectives for the field

Looking towards the future of the limb regeneration field, I believe there are many exciting potential lines of research that stem from our work presented here along with recent advances including the release of the axolotl genome (Nowoshilow et al., 2018;

Smith et al., 2019), the optimization of CRISPR targeting strategies (Fei et al., 2018), and the elucidation of cell-type specific markers (Gerber et al., 2018; Leigh et al., 2018).

Several of these potential directions are discussed in more detail in the following sections.

6.6.1. Exploring the roles of unannotated genes in salamander regeneration

While the focus of many studies in the limb regeneration field has been on identifying roles for genes that have known orthologs within other species, our dataset along with the publicly available datasets of others have uncovered that the majority of differentially expressed transcripts during early stages of regeneration are unannotated.

Oftentimes, these were the most highly differentially expressed transcripts, suggesting that these may be salamander specific genes that are critical regulators of limb regeneration. In addition, the axolotl genome is large (~32 gigabases), yet contains approximately the same number of genes as humans, further indicating that there may be important salamander-specific non-coding elements regulating limb regeneration.

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Therefore, an exciting future line of research would be to utilize CRISPR or morpholino knockdown strategies to perturb these unannotated genes and determine whether they play important roles in regeneration.

6.6.2. Cell-type specific roles in blastema formation

While we focused primarily on examining the functional roles of blastemal- progenitor enriched transcripts, there were many differentially expressed transcripts within the regenerating non-dividing stump tissues that potentially also play important paracrine roles in regeneration. Future studies investigating where these transcripts are expressed and functionally testing whether they are relevant could uncover novel roles that supportive cell-types play in limb regeneration. For example, one potential such cell- type could be the vasculature, as endothelial cells have been shown to play roles in induction of liver regeneration (Ding et al., 2010). In addition, it is interesting to speculate on whether these non-dividing cells, for example mature myofibers at the site of amputation, actually play functional paracrine roles in regeneration and this remains an unexplored area of limb regenerative biology.

Another more targeted approach to address the importance of different cell types on blastema formation would be to generate reporters for various cell types and transcriptionally profile a preferred cell type to ask how it responds during early stages of regeneration. Recent single cell studies have uncovered many potential cell-type specific markers that may be utilized for these studies (Leigh et al., 2018) and others have already begun to combine these approaches with single cell analysis to identify gene expression programs present within specific tissue lineages (Gerber et al., 2018).

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6.6.3. Genetic regulatory mechanisms of key genes

The release of the genome opens up the possibility of understanding genetic regulatory mechanisms of various genes in limb regeneration including key blastemal markers such as prrx-1 and kazd-1. In addition, the important roles that enhancer regulation plays in the regeneration of other organisms (Kang et al., 2016) have yet to be uncovered in the field of limb regeneration. Examining these elements may uncover important core regulatory circuits driven by key transcription factors (ex. sall4) (Erickson et al., 2016) for various cell types during the course of limb regeneration.

6.7. Concluding remarks

In all, my doctoral research began as an examination of the transcriptional programs in blastemal progenitors and the surrounding tissues during early stages of limb regeneration both in the presence and absence of the wound epidermis. In doing so, we uncovered the roles of the early wound epidermis and further identified blastemal progenitor-enriched as well as wound epidermis-dependent transcripts. Utilizing this resource, we further elucidated the important roles that two cytokines, il-8 and mk, play in regulating blastema formation. Altogether, these data provide the field with resources for new gene candidates that may regulate the initiation of blastema formation and in turn reveal new aspects of limb regenerative biology.

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Appendix

A.1. Supplemental materials for Chapter 2

Table A.1.1. Table of Ingenuity Pathway Analysis (IPA) Z-scores for Immune Signaling Pathways.

Pathway 4dpa 4N 5dpa 4N 4dpa 2N 5dpa 2N 4dpa WE 5dpa WE Activation of IRF 0.377964473 1 2.323790008 2.110579412 2.121320344 0.816496581 by Cytosolic Pattern Recognition Receptors Role of RIG1-like 0 0 2.333333333 1.889822365 2.449489743 1 Receptors in Antiviral Innate Immunity TREM1 Signaling 1.414213562 0 2.449489743 1.963961012 1.133893419 2.121320344 Interferon -1.807392228 -1.732050808 -1.876629727 -1.705605731 1.941450687 0 Signaling Inflammasome -1.807392228 -1.732050808 -1.876629727 -1.705605731 1.941450687 0 pathway Acute Phase 0.774596669 0.577350269 1.060660172 0.784464541 1.290994449 0.707106781 Response Signaling Oncostatin M 1.341640786 0 1.414213562 0.632455532 2.645751311 0 Signaling Complement 1.341640786 0 1.414213562 0.632455532 2.645751311 0 System Fc-gamma -0.25819889 -1.290994449 1.80838886 1.671258044 1.889822365 2.236067977 Receptor- mediated Phagocytosis in Macrophages and Monocytes Leukocyte -1.133893419 -0.962250449 1.832541665 1.483239697 2.523573073 1.732050808 Extravasation Signaling Role of Pattern -0.534522484 -2.333333333 2.057983022 1.616447718 2.840187787 2.333333333 Recognition Receptors in Recognition of Bacteria and Viruses Role of IL-17F in 0 0 1.666666667 1.889822365 0 0 Allergic Inflammatory Airway Diseases B Cell Activating 0 0 1.507556723 1.632993162 0 0 Factor Signaling April Mediated 0 0 1.507556723 1.133893419 -1 0 Signaling "Role of JAK1, 0 0 -1 -0.447213595 1 0 JAK2 and TYK2 in Interferon Signaling" IL-22 Signaling 0 0 0.707106781 1.133893419 0.447213595 0 Coagulation 0 -0.707106781 0.301511345 0.333333333 0 0 System TNFR1 Signaling 0 0 0 1.133893419 -1.264911064 -1.341640786 TNFR2 Signaling 0 0 0 0.816496581 -2.449489743 -2 STAT3 Pathway -3.16227766 -2.309401077 -3.528211425 -3.265986324 0 -0.447213595 Calcium-induced -3.16227766 -2.309401077 -3.528211425 -3.265986324 0 -0.447213595 T Lymphocyte Apoptosis iCOS-iCOSL -2.309401077 -0.707106781 -1.299867367 -0.816496581 1 0 Signaling in T Helper Cells

166 Table A.1.1. (Continued)

Pathway 4dpa 4N 5dpa 4N 4dpa 2N 5dpa 2N 4dpa WE 5dpa WE Role of NFAT in -2.840187787 -1.632993162 -1.299867367 -1.224744871 0.832050294 0 Regulation of the Immune Response Th1 Pathway -1.290994449 -0.904534034 -0.784464541 0.185695338 -0.904534034 -0.447213595 Th2 Pathway -1.290994449 -1.290994449 0.19245009 0 0.577350269 0 CD28 Signaling in -2.529822128 -0.707106781 -0.707106781 -0.39223227 0 -1.341640786 T Helper Cells IL-9 Signaling -1.133893419 -0.816496581 -1.069044968 -1.386750491 0 0 JAK/Stat -0.301511345 -0.301511345 -1.279204298 -1.788854382 0.25819889 -0.816496581 Signaling CD40 Signaling -1.414213562 -1.133893419 0 -0.5 -0.707106781 -0.447213595 PKC_ Signaling -3.050851079 -1.666666667 -0.755928946 -0.6 0 -1.632993162 in T Lymphocytes LPS-stimulated -3.050851079 -2.529822128 -0.942809042 -1 0 -1 MAPK Signaling NF-kB Activation -2.110579412 -2.110579412 -0.852802865 -1.091089451 1.154700538 0.377964473 by Viruses fMLP Signaling in -2.523573073 -1.807392228 -0.522232968 -1.347150628 2.110579412 1 Neutrophils Tec Kinase -1.697749375 -1.5 0 0 0.832050294 1.133893419 Signaling Role of PI3K/AKT -1 -0.377964473 -1.069044968 -1.941450687 1.133893419 0 Signaling in the Pathogenesis of Influenza GM-CSF -0.632455532 -0.301511345 -0.447213595 -0.654653671 1.154700538 1.341640786 Signaling Production of -1.632993162 -2.064741605 -1.154700538 -0.333333333 0 0.301511345 Nitric Oxide and Reactive Oxygen Species in Macrophages Chemokine -2.828427125 -0.816496581 0 -0.727606875 0.377964473 0 Signaling Macropinocytosis -2.529822128 -1.897366596 -1.414213562 -1.603567451 1.133893419 0 Signaling NF-kB Signaling -0.471404521 -1.290994449 -0.832050294 -0.589767825 1.527525232 1.264911064 IL-6 Signaling -1.697749375 -1.386750491 -0.648885685 -0.353553391 0.25819889 -0.377964473 Fc_RIIB Signaling -1.414213562 -1.341640786 -1.5 -1.807392228 1.666666667 0 in B Lymphocytes IL-2 Signaling -1.133893419 -1.632993162 -2.138089935 -1.941450687 0.632455532 -1 HMGB1 Signaling -1.414213562 -1.807392228 -1.767766953 -2.2 0 0

Table A.1.2. Table of Ingenuity Pathway Analysis (IPA) Z-scores for Developmental Growth Factor and Intracellular Signaling Pathways.

Pathway 4dpa 4N 5dpa 4N 4dpa 2N 5dpa 2N 4dpa WE 5dpa WE CXCR4 Signaling -3.299831646 -2.828427125 -2.030258905 -2.197401062 0.301511345 1 Ephrin Receptor -2.121320344 -3 0.755928946 0.208514414 1.133893419 1 Signaling HIPPO signaling 0.707106781 1 2.309401077 2.529822128 1.133893419 0 Wnt/Ca+ pathway -2.645751311 -2 -2.523573073 -2.110579412 0 0 Wnt/Beta-catenin 0.25819889 0.577350269 0.948683298 0.755928946 -2 -1 Signaling TGF-Beta 0.632455532 1.414213562 -1 -1.069044968 -0.447213595 0 Signaling CNTF Signaling -1 -1.414213562 -1.147078669 -1.414213562 1.154700538 0 NGF Signaling -1.941450687 -2.333333333 -1.347150628 -0.625543242 1.603567451 0 FGF Signaling -2.110579412 -1 -2.132007164 -2.523573073 0 0

167 Table A.1.2. (Continued)

Pathway 4dpa 4N 5dpa 4N 4dpa 2N 5dpa 2N 4dpa WE 5dpa WE VEGF Signaling -1.507556723 -1.133893419 -2.857738033 -3.299831646 1.414213562 0 ErbB4 Signaling -1.897366596 -2.121320344 -2.323790008 -2.713602101 1.133893419 0 Neuregulin -1.897366596 -2.828427125 -2.5 -3.207134903 1.341640786 0 Signaling Notch Signaling -2 -2 -0.816496581 -1.133893419 0 0 IGF-1 Signaling -0.904534034 -1.264911064 -2.293658555 -2.449489743 0 -0.447213595 Neurotrophin/TRK -1.507556723 -0.816496581 -2.132007164 -1.414213562 0.904534034 0 Signaling PDGF Signaling -0.577350269 -1.154700538 -1 -1.459600898 0.534522484 -0.816496581 HGF Signaling -2.713602101 -2.110579412 -1.732050808 -1.963961012 0.277350098 -0.816496581 ErbB2-ErbB3 -1.666666667 -1.632993162 -1.603567451 -2.323790008 1.133893419 0 Signaling EGF Signaling -0.904534034 -0.707106781 -1.459600898 -1.147078669 0.301511345 0 Growth Hormone -1.732050808 -1.507556723 -2.2 -1.885618083 0 -1 Signaling mTOR signaling -1.212678125 -2.182820625 -0.89802651 -0.6 1.414213562 0.377964473

Calcium signaling -1.507556723 0.447213595 -1.04257207 -0.447213595 0 N/A

Table A.1.3. Blastemal-progenitor enriched transcripts. Negative fold change indicates enrichment in blastemal progenitors.

Transcript ID/Blastx hit log2(FC) padj

c1081013_g5 TENA_HUMAN -1.62815405 8.49E-18

c1082828_g4 TTK_MACFA -1.964713877 7.57E-13

c1063668_g1 FHDC1_HUMAN -2.049134955 2.17E-12

c1088332_g1 TENA_CHICK -1.628989579 2.17E-12

c1088332_g1 TENA_CHICK -1.891231003 6.82E-10

c1085526_g1 SHD_HUMAN -1.842004233 2.00E-09

c1084875_g3 RSMN_RAT -1.480592796 2.22E-09

c1069546_g2 PRD16_HUMAN -3.235269881 2.88E-09

c1088917_g3 ASAP3_HUMAN -1.84283593 2.92E-09

c1054335_g2 CTK2_XENLA -3.352475926 4.13E-09

c1092373_g1 AIM1_HUMAN -1.219467611 1.33E-08

c1088332_g3 TENA_CHICK -1.421019508 2.23E-08

c1091712_g2 PKHH2_HUMAN -1.639626647 2.50E-08

c1086898_g3 GLI3_CHICK -1.585874373 2.95E-08

c1086150_g5 CO5A1_HUMAN -1.338292607 3.22E-08

c1031898_g1 HAS2_HUMAN -1.63755093 3.32E-08

c1067239_g7 RCN1_MOUSE -1.559643023 3.87E-08

c1088332_g1 TENA_CHICK -1.435037885 4.99E-08

c1070536_g3 CO5A1_MOUSE -2.123463542 5.11E-08

168 Table A.1.3. (Continued)

Transcript ID/Blastx hit log2(FC) padj

c1090290_g1 EMIL1_MOUSE -1.542007837 6.79E-08

c1076079_g1 GP1_CHLRE -1.201822658 7.06E-08

c1086236_g7 GLI2_HUMAN -2.896206816 9.30E-08

c1088332_g1 TENA_CHICK -1.894540705 1.09E-07

c1075841_g1 RABP2_HUMAN -2.003202312 1.37E-07

c1080761_g4 CO1A2_CHICK -1.544792872 1.53E-07

c1088664_g5 MAP1B_HUMAN -1.494007131 1.61E-07

c1088922_g4 MAP1B_HUMAN -1.556997542 1.62E-07

c1039642_g1 PTRF_RAT -1.555315423 1.62E-07

c1091175_g9 CD248_HUMAN -1.453305223 2.04E-07

c1062836_g4 LEP_SMICR -1.637221897 2.66E-07

c1032646_g1 SRAP_STAHJ -1.352291119 3.65E-07

c1077000_g1 SALL4_HUMAN -1.753022515 4.58E-07

c1089148_g2 G3ST4_BOVIN -1.384775945 5.46E-07

c1089394_g5 CO5A2_HUMAN -1.449369473 6.57E-07

c1090227_g2 FKBP9_HUMAN -2.083684231 6.63E-07

c1086257_g1 K0754_HUMAN -1.391754216 7.41E-07

c1091199_g1 CO5A2_MOUSE -1.27661545 7.41E-07

c1083713_g1 CALD1_CHICK -1.495252853 7.63E-07

c1060273_g2 SIA4B_PANTR -1.531992957 1.24E-06

c1092028_g1 KCP_XENLA -1.501640661 1.26E-06

c1082369_g3 TEN3_HUMAN -1.676491772 1.29E-06

c1086150_g5 CO5A1_HUMAN -1.205919238 1.33E-06

c1078376_g4 HTRA1_BOVIN -1.502864308 1.52E-06

c1055997_g1 SDC1_BOVIN -1.391595493 1.55E-06

c1078833_g1 INHBA_SHEEP -1.295455008 1.94E-06

c1086874_g10 CTHR1_RAT -1.707535666 2.19E-06

c1076723_g2 NFAC4_MOUSE -1.094884328 2.62E-06

c1060722_g1 ANGL2_HUMAN -1.188802927 3.49E-06

c1088743_g1 MXRA5_HUMAN -1.310755008 3.73E-06

c1088332_g1 TENA_CHICK -1.537700614 5.70E-06

c1080758_g1 RAI14_HUMAN -1.417754519 5.89E-06

c1066962_g3 SYNPO_MOUSE -1.386359684 6.01E-06

c1058708_g1 PTK7_CHICK -1.21218312 6.54E-06

c1075291_g1 IGS10_HUMAN -1.384157953 8.48E-06

169 Table A.1.3. (Continued)

Transcript ID/Blastx hit log2(FC) padj

c1090482_g1 TANC2_MOUSE -1.408588194 1.01E-05

c1068056_g1 MAMC2_HUMAN -1.729013746 1.04E-05

c1067239_g7 RCN1_HUMAN -1.558808307 1.09E-05

c1091370_g1 LTBP1_HUMAN -1.34600726 1.12E-05

c1064348_g8 FBN2_HUMAN -1.341511901 1.12E-05

c1088087_g4 GPC6_MOUSE -1.621957635 1.18E-05

c1070536_g3 CO5A1_CRILO -1.085109475 1.38E-05

c1070223_g2 CCD80_CHICK -1.174462926 1.41E-05

c1077695_g7 DDR2_HUMAN -1.540640808 1.46E-05

c1088518_g8 COCA1_HUMAN -1.591215323 1.73E-05

c1090227_g2 FKBP9_HUMAN -1.019973238 2.02E-05

c1087294_g1 DCTN1_XENLA -2.776940233 2.24E-05

c1068194_g2 LAMA1_MOUSE -1.522410716 2.43E-05

c1082616_g4 AEBP1_MOUSE -1.437518031 2.48E-05

c1057474_g1 MKA_XENLA -1.30457236 2.65E-05

c1089533_g3 CP1B1_HUMAN -1.755299095 2.72E-05

c1082616_g4 AEBP1_MOUSE -1.983425604 2.87E-05

c1090597_g4 ROA1_PANTR -1.027611182 3.18E-05

c1076561_g4 KAZD1_MOUSE -1.696488986 3.55E-05

c1080519_g6 STC1_RAT -1.735927259 3.57E-05

c1052564_g1 IPO4_HUMAN -1.062056765 3.80E-05

c1075231_g15 GLIS2_HUMAN -1.649615892 4.27E-05

c1092042_g1 HMCN1_HUMAN -1.112403764 4.41E-05

c1083713_g1 CALD1_CHICK -2.334999933 4.44E-05

c1069073_g1 GLIS3_HUMAN -1.712537 5.28E-05

c1078917_g3 TSP2_CHICK -1.305427235 5.47E-05

c1079712_g5 EFNB1_CHICK -1.703324035 5.50E-05

c1081670_g1 CO1A1_CYNPY -1.57164569 5.57E-05

c1079703_g4 MUC16_HUMAN -1.253184834 6.28E-05

c1042639_g3 EGR1_XENTR -1.041987165 6.41E-05

c1087063_g1 PEBP1_BOVIN -1.600667726 6.51E-05

c1064163_g1 LRRC4_RAT -1.436487075 6.60E-05

c1087698_g2 LIN41_MOUSE -1.510269801 7.12E-05

c1091365_g1 PTRF_RAT -1.548737242 7.30E-05

c1077160_g2 HIPL1_MOUSE -1.43063085 7.30E-05

170 Table A.1.3. (Continued)

Transcript ID/Blastx hit log2(FC) padj

c1057844_g1 HMGB3_CHICK -1.628060528 7.91E-05

c1086236_g7 GLI3_CHICK -1.473774244 9.48E-05

c1080793_g1 AOC3_MOUSE -1.524666452 9.67E-05

c1091081_g5 FLNA_MOUSE -1.128819808 9.79E-05

c1085559_g2 TPM2_CHICK -1.465327966 0.000100445

c1084310_g1 K1614_HUMAN -1.028435563 0.000102714

c1092253_g2 CRLD2_XENTR -1.147183901 0.000112876

c1067676_g1 KIRR1_HUMAN -2.58051131 0.000126195

c1092042_g2 HMCN1_HUMAN -1.002732419 0.000137448

c1086285_g2 K1C24_HUMAN -1.458029516 0.000151273

c1081277_g3 MDR1_CRIGR -1.357249752 0.000151307

c1088518_g9 COCA1_CHICK -1.295865793 0.000173138

c1070827_g2 PA24C_HUMAN -1.337615424 0.00017585

c1081345_g2 SYNE2_HUMAN -1.329285938 0.000188196

c1080761_g5 CO1A2_CHICK -1.107198956 0.000199213

c1092516_g4 K2C8_XENLA -1.059194038 0.000229186

c1088518_g10 COCA1_CHICK -1.365162947 0.000237354

c1074886_g4 STXB1_RAT -1.889097538 0.000245851

c1083301_g5 GPX7_HUMAN -1.436887108 0.000255386

c1089244_g1 FA76B_XENTR -2.491127615 0.000255386

c1086707_g4 IF4G1_HUMAN -1.15642966 0.000287863

c1086497_g1 CXB3_HUMAN -2.094376979 0.000297996

c1057320_g1 POL2_MOUSE -1.379174899 0.000358954

c1091395_g2 NOTC2_HUMAN -1.032055635 0.000369935

c1087193_g2 AF1L1_XENTR -1.59785317 0.000399431

c1080761_g7 CO1A2_CHICK -1.412147931 0.000404478

c1084248_g3 EPHB2_MOUSE -1.781330213 0.000404845

c1083151_g6 F18A1_HUMAN -1.187103625 0.000422371

c1085297_g4 CO3A1_CHICK -1.085065106 0.000437783

c1086150_g4 CO5A1_RAT -1.247982041 0.000449281

c1072542_g2 EHD3_RAT -1.562299354 0.000475464

c1089750_g2 CO6A2_CHICK -1.065074859 0.000475464

c1085057_g1 TSP3_HUMAN -1.215648117 0.000557248

c1080761_g5 APP1_SCHPO -1.08430152 0.000629249

c1318008_g1 YI31B_YEAST -1.573965262 0.00067379

171 Table A.1.3. (Continued)

Transcript ID/Blastx hit log2(FC) padj

c1070561_g6 KI67_HUMAN -1.526526937 0.000725726

c1084054_g3 TRIL_HUMAN -1.410432037 0.000776707

c1082616_g4 AEBP1_MOUSE -1.205965588 0.000848867

c1053631_g1 RHG10_HUMAN -1.142115997 0.000849154

c1076669_g1 P3H1_CHICK -1.060979131 0.000871103

c1072608_g3 TGFB2_CHICK -1.491070906 0.00107799

c1076995_g1 PGFRB_CANFA -1.086249241 0.00107865

c1087698_g2 LIN41_MOUSE -1.314847573 0.001083247

c779314_g2 SLIK6_HUMAN -1.351817869 0.001086423

c1092212_g1 GULP1_HUMAN -1.815522249 0.001143186

c1081287_g2 F198A_HUMAN -2.055296761 0.001146918

c1090747_g1 CR3L2_HUMAN -1.1380815 0.001194214

c1067690_g1 TPC1_RAT -1.319430595 0.001332518

c1091062_g2 K2C5_RAT -1.240429022 0.001358011

c1011988_g1 APCDL_HUMAN -1.118233255 0.001421737

c1075918_g14 FBN2_MOUSE -1.164880596 0.001487003

c1092110_g2 TEAD1_HUMAN -1.215462796 0.001502114

c1071771_g2 IL11_HUMAN -1.471694465 0.001583637

c1090056_g2 PGBM_HUMAN -1.045555121 0.001612064

c1080288_g3 LIN1_NYCCO -1.164300756 0.001719395

c1084795_g1 CEAM6_HUMAN -1.251503463 0.001876703

c1065468_g2 VKORL_RAT -1.146114231 0.001983796

c1086384_g12 IE2_HHV6Z -1.175657735 0.002008734

c1085297_g4 CO3A1_RAT -1.501636138 0.00206446

c1084565_g2 CTHR1_RAT -1.6966408 0.002095306

c1066349_g1 2A5B_RABIT -1.238348394 0.002098311

c1086150_g1 CO5A1_HUMAN -1.101897245 0.002144871

c1091062_g2 K2C5_RAT -1.245582316 0.002258554

c1079936_g2 MUC1_YEAST -1.43729237 0.002268624

c1090326_g2 CAD26_HUMAN -1.414996452 0.002320877

c1090048_g1 ASPH_BOVIN -1.336562405 0.002439065

c1053070_g2 HXDBA_TAKRU -1.210625764 0.002532294

c1086597_g2 ITA11_MOUSE -1.232702654 0.002537417

c1077260_g1 IL8_CHICK -1.445967548 0.002568126

c1088332_g1 TENA_CHICK -1.245028019 0.002685534

172 Table A.1.3. (Continued)

Transcript ID/Blastx hit log2(FC) padj

c1089394_g5 CO5A2_MOUSE -1.139360785 0.002845216

c1085414_g1 PERP_HUMAN -1.263960198 0.002918138

c1088743_g1 MXRA5_HUMAN -1.219450939 0.003025128

c1076392_g2 S5A2_MACFA -1.353615963 0.003142404

c1080761_g5 CO1A2_CHICK -1.389868532 0.003271336

c1068344_g3 CLCF1_HUMAN -1.553161092 0.003639362

c1073265_g1 PINLY_HUMAN -1.446070187 0.00365578

c1077640_g1 TM119_MOUSE -1.354265805 0.003711084

c1067201_g1 DKK3_CHICK -1.06951219 0.003735079

c1078588_g4 FZD2_MOUSE -1.169583636 0.003740425

c1082413_g5 ITB8_HUMAN -1.151239335 0.003810041

c1091311_g1 NCKX3_HUMAN -1.171755343 0.003946259

c1080550_g10 POSTN_HUMAN -1.049576007 0.003991044

c1080690_g6 WNT5B_AMBME -1.750254437 0.00404423

c1079703_g6 MUC16_HUMAN -1.460211479 0.004115024

c1087877_g3 BKRB2_RABIT -2.080319717 0.004123495

c1063943_g2 ANXA1_HUMAN -1.222079461 0.004148759

c1090220_g1 ANK2_MOUSE -1.751250542 0.004173209

c1075239_g2 PRIC2_HUMAN -1.173102687 0.004277638

c1091168_g1 NATT4_THANI -1.272113053 0.004312634

c1068090_g7 PICK1_RAT -2.057270373 0.004762796

c1080320_g1 CLIC3_HUMAN -1.572361944 0.004961711

c1067510_g1 EDNRA_HUMAN -1.084910964 0.005289689

c1079263_g1 ATN1_MOUSE -1.05283968 0.006292201

c1077896_g1 LRIG3_HUMAN -1.52298402 0.0063786

c1079594_g5 SOX7_XENTR -1.8931725 0.006434241

c1035197_g1 IOD3_LITCT -1.230023884 0.006480243

c964270_g2 SFRP5_BOVIN -1.557930273 0.006579446

c1081670_g1 CO1A1_CYNPY -1.276296739 0.006751355

c1067048_g2 PRRX1_RAT -1.176106213 0.006990471

c1077344_g4 ATS6_HUMAN -1.982280391 0.007264198

c1075751_g2 GOLI4_HUMAN -1.027959779 0.007314435

c1078917_g3 TSP2_CHICK -1.493058243 0.007443144

c1072351_g2 FKBP7_PONAB -1.14546987 0.007733289

c1064028_g3 CHRD_XENLA -1.466052856 0.007809942

c1076187_g1 PKD1_MOUSE -1.236770571 0.008174066

173 Table A.1.3. (Continued)

Transcript ID/Blastx hit log2(FC) padj

c1072068_g2 FAAA_HUMAN -1.255799793 0.008889508

c1089199_g1 PTPRS_HUMAN -1.02539146 0.009038492

c1080585_g1 TTC36_DANRE -1.272513258 0.009063231

c1077344_g4 ATS6_HUMAN -1.595138396 0.009777646

c1071990_g3 PEAK1_HUMAN -1.550446575 0.009786975

c1091343_g2 PCD16_HUMAN -1.218092115 0.010041898

c1060840_g1 TSN6_MOUSE -1.124805091 0.010418034

c1064516_g1 HXD10_CHICK -1.099147158 0.010446283

c1075159_g3 DCNL4_HUMAN -1.918577159 0.010571858

c1078136_g3 PLOD2_HUMAN -1.607874636 0.010631286

c1084384_g1 TF3A_HUMAN -1.913373809 0.010736089

c1059195_g1 UBE2C_XENLA -1.768332576 0.010783089

c1091062_g2 K2C5_RAT -1.032193232 0.010843835

c1068714_g1 TGFB1_HUMAN -1.065374877 0.011093989

c1072538_g10 ZFHX3_HUMAN -1.010426264 0.011157654

c1086374_g1 PALLD_HUMAN -1.569493276 0.01157875

c1090567_g1 HRPX_PLALO -1.231395804 0.011581652

c1080550_g10 POSTN_HUMAN -1.441992866 0.012119946

c1090056_g2 PGBM_HUMAN -1.070785001 0.012243491

c1085540_g2 LIN1_NYCCO -1.171341624 0.012749929

c1090567_g1 GTAN_DICDI -1.172953412 0.012975785

c1088267_g2 GIT1_HUMAN -1.862776613 0.013317992

c1087228_g1 PARD3_HUMAN -1.655498788 0.01355353

c1054023_g1 HOME1_HUMAN -1.655985123 0.014418856

c1088364_g2 POLR_DROME -1.107477007 0.014607394

c1562183_g1 Y8592_DICDI -1.72743958 0.014955866

c1057994_g4 S10A4_MOUSE -1.120691799 0.014975013

c1609543_g1 ZC3HD_HUMAN -1.360463358 0.015011296

c1084104_g1 ANKR1_XENLA -1.250347951 0.015936228

c1092516_g5 K2C5_PANTR -1.498116687 0.016009704

c1082602_g2 EMIL2_HUMAN -1.097912059 0.016038048

c1076922_g10 PXDN_HUMAN -1.184837546 0.016301542

c1081684_g3 RGS8_DANRE -1.822644086 0.016385821

c1091081_g5 FLNA_HUMAN -1.628838209 0.016803259

c512704_g1 POL2_MOUSE -1.490774095 0.016948713

c1085979_g1 TB15A_COTJA -1.004730221 0.017796681

174 Table A.1.3. (Continued)

Transcript ID/Blastx hit log2(FC) padj

c1089316_g1 KI26A_MOUSE -1.810912029 0.017848806

c1082235_g1 RFC2_BOVIN -1.003890612 0.017940271

c1086652_g2 MPP2_MOUSE -1.578352643 0.018425321

c1065609_g1 DEN_GAHVM -1.741577691 0.01925478

c1071270_g3 CERS2_BOVIN -1.544916924 0.019787062

c1090284_g2 INSR_XENLA -1.0219355 0.019949055

c1060102_g1 CA051_HUMAN -1.63541411 0.020226855

c1089919_g4 CP27B_HUMAN -1.061371127 0.020624008

c1082458_g3 CR3L1_HUMAN -1.039923496 0.020783986

c1069159_g1 HXA3_CHICK -1.237184226 0.021293166

c1089220_g1 NCAM1_CHICK -1.229882628 0.02168676

c1090056_g1 PGBM_HUMAN -1.208347529 0.021919659

c1088272_g1 BMP1_HUMAN -1.396991405 0.023210412

c1071958_g2 ANXA6_HUMAN -1.710590635 0.023765726

c1081900_g1 S100P_HUMAN -1.03261438 0.023944888

c1089272_g1 SH3B4_HUMAN -1.013793913 0.023967299

c1076561_g4 KAZD1_MOUSE -1.381228821 0.024519327

c1091343_g3 PCD16_HUMAN -1.2809168 0.024611682

c1070469_g3 CAMP_GORGO -1.188002015 0.025303941

c1071878_g3 MAP6_HUMAN -1.665448719 0.025303941

c1068736_g1 CKAP4_HUMAN -1.12741589 0.025590395

c1087705_g1 ELF3_MOUSE -1.380699729 0.026930308

c1070593_g11 COPZ2_MOUSE -1.756403551 0.02693712

c1075841_g1 RABP2_RAT -1.473641701 0.027043261

c1070839_g1 TBX4_MOUSE -1.146268465 0.027327461

c1092545_g3 GT251_HUMAN -1.055627508 0.028028006

c1075851_g1 SPICE_BOVIN -1.392611027 0.028299796

c1077247_g1 SPTN2_HUMAN -1.295101686 0.028299796

c1239849_g1 RBM25_HUMAN -1.064340795 0.02931604

c1741796_g1 POL_GALV -1.192998664 0.030794864

c1088332_g2 TENA_CHICK -1.043152377 0.031542868

c1079613_g7 MATN2_HUMAN -1.083938907 0.031904477

c1085540_g1 COBA1_HUMAN -1.075106813 0.031973336

c1075197_g2 MRCKA_HUMAN -1.031965297 0.032400105

c1506666_g1 LIN1_NYCCO -1.613582534 0.032853189

c1079879_g3 DESP_HUMAN -1.164657262 0.032935412

175 Table A.1.3. (Continued)

Transcript ID/Blastx hit log2(FC) padj

c958434_g1 LIN1_NYCCO -1.241811913 0.032986956

c1087929_g5 ZMIZ1_MOUSE -1.644034935 0.03343453

c1086762_g2 PRDM1_MOUSE -1.037159595 0.033468811

c1082499_g1 TT21B_HUMAN -1.657809837 0.033873062

c1083541_g2 FLIP1_HUMAN -1.57123845 0.035281919

c1086999_g4 BNC1_HUMAN -1.688718942 0.035404541

c1086859_g1 EPHA2_HUMAN -1.504068199 0.035864674

c1089895_g1 DREB_HUMAN -1.564965924 0.0375888

c1079879_g3 DESP_MOUSE -1.033903421 0.037974347

c1090571_g2 TBX5_CHICK -1.581788487 0.038612838

c874462_g1 VTCN1_MOUSE -1.462808989 0.038779132

c1072235_g1 K118A_XENLA -1.421259515 0.040136708

c959265_g1 CYGB2_DANRE -1.456092206 0.040343855

c1089316_g1 KI26A_MOUSE -1.30167432 0.04074388

c1075231_g16 GLIS2_XENLA -1.597640513 0.041175045

c1076361_g2 UPK2_MOUSE -1.145928797 0.041343099

c1064174_g2 ATLA2_MOUSE -1.625825936 0.041405604

c1091668_g2 ANR35_HUMAN -1.060149427 0.042035152

c1248822_g1 CSP_PLARE -1.15171914 0.042573582

c1086285_g2 K1C24_HUMAN -1.043333024 0.042818789

c1088042_g1 MMP13_XENLA -1.454817898 0.042956529

c1078665_g1 MB212_MOUSE -1.1239959 0.045511497

c1087883_g1 IGSF3_HUMAN -1.102264552 0.045539183

c1083946_g1 PMEPA_HUMAN -1.053759928 0.045719696

c1084637_g1 SARG_RAT -1.367127717 0.047513781

c1074252_g1 BORG4_HUMAN -1.405534456 0.048348856

c1080374_g3 XBP1_MOUSE -1.401671465 0.048881237

c568518_g1 LIN1_NYCCO -1.445656027 0.04910916

c1089812_g1 NAV2_HUMAN -1.427049397 0.049180296

c1060685_g1 PRRX2_HUMAN -1.18884741 0.049512451

c1076625_g1 CSRN2_HUMAN -1.269708601 0.049918573

176

Figure A.1.1. Immune signaling pathways are predominantly activated in the wound epidermis and non-dividing cells. (A) Heatmap depicting the predicted activation or inhibition of many immune signaling pathways detected using the IPA software. The isolated dividing cellular fraction exhibits predicted activation of very few and inhibition of a majority of immune signaling pathways.

177 A.2. Supplemental materials for Chapter 3

Figure A.2.1. Validation of the il-8 overexpression construct and il-8 morpholino knockdown. (A) Western blot validation of cells transfected with either the il-8 overexpression vector or control vector (T2A-tdTomato). Cell lysate and media was collected 48 hpt. Myc-staining is specific to 293T cell lysate and media. (B-E) Immunostaining of 293T cells transfected with the il-8 overexpression vector (IL8myc- T2A-tdT) 48 hours post-transfection (hpt). (F) Western blot validation of knockdown of IL- 8 expression in limbs injected with il-8 translation blocking morpholino (MO) vs. control MO. All four limbs were taken from one animal. FL, forelimb, HL, hindlimb.

178

Figure A.2.2. IL-8 morpholino knockdown does not affect blastemal cell or wound epidermis proliferation. (A-B) Representative images of EdU-stained sections of control or il-8 morpholino treated limbs. Scale bars represent 200 µm and white dotted line demarcates the amputation plane. (C-D) Quantification of the percentage of EdU+ cells in either the blastema (C) or apical epithelial cap (AEC) (D).

179 A.3. Supplemental materials for Chapter 4

Table A.3.1. Top 100 differentially expressed transcripts in stump-derived dividing cells of full skin flap sutured limbs. Negative fold change indicates enrichment in full skin flap sutured limbs.

transcript id blastx hit logFC padj

c1087956_g9_i4 EWS_HUMAN 5.599435134 4.00E-11

c1090300_g1_i1 CNN2_MOUSE 10.17192719 2.07E-10

c1090687_g2_i3 PTBP1_BOVIN 10.4881593 5.69E-10

c1091001_g2_i3 CDCA2_HUMAN 8.62454394 1.60E-09

c1083454_g6_i2 ZBTB3_HUMAN 9.266583269 4.48E-08

c1086809_g3_i3 BI1_BOVIN 9.377086079 4.48E-08

c1082127_g2_i6 HERC1_HUMAN 7.825040137 6.60E-08

c1090706_g8_i5 STAM2_HUMAN 9.331682098 2.98E-07

c1049691_g1_i1 MPI1B_XENLA -9.111045299 3.28E-07

c1091340_g1_i3 K0317_MOUSE 8.893277374 5.17E-07

c1091444_g2_i5 MPP5_HUMAN -9.410160283 1.24E-06

c1083468_g1_i3 ELYS_HUMAN 8.934284289 1.46E-06

c1078658_g1_i1 RS10_HUMAN 8.959774488 1.60E-06

c1075582_g1_i3 NU155_HUMAN -9.187643789 2.42E-06

c1089782_g7_i2 PPARD_CANFA 8.784970234 2.66E-06

c1070268_g1_i5 ZC3HF_HUMAN 7.374405656 2.92E-06

c1088530_g5_i1 WNK1_HUMAN 8.4471637 3.17E-06

c1074527_g1_i10 EXOC3_MOUSE -8.752858773 3.17E-06

c1083410_g1_i2 PDCD4_MOUSE 8.499494561 3.17E-06

c1080761_g5_i1 CO1A2_CHICK 7.998227333 4.01E-06

c1077444_g4_i5 UNC5B_XENLA 8.428399454 4.01E-06

c1066045_g1_i3 DPOD1_RAT 8.36478719 4.92E-06

c1091174_g4_i4 LIN1_NYCCO -9.244637259 4.92E-06

c1079168_g1_i1 POL2_MOUSE -8.398669414 4.92E-06

c1080282_g1_i3 OSBL5_HUMAN 8.415537213 5.43E-06

c1089852_g4_i6 ADCY2_HUMAN 8.240339587 6.72E-06

c1065086_g1_i1 SPAG1_HUMAN -8.766342421 6.86E-06

c1088974_g4_i1 DSEL_HUMAN -8.526652263 6.87E-06

c1046138_g1_i1 Y8359_DICDI -3.260881536 7.83E-06

c1082570_g3_i2 CCNL2_HUMAN 8.274528644 8.83E-06

c1075167_g1_i4 MOT2_HUMAN -8.287112996 9.45E-06

180 Table A.3.1. (Continued)

transcript id blastx hit logFC padj

c1089216_g3_i4 SBNO2_BOVIN 9.001641319 1.08E-05

c1068424_g1_i4 EP15R_HUMAN 8.610848855 1.14E-05

c1077195_g2_i9 KC1A_XENLA 8.160667469 1.30E-05

c1067524_g1_i2 TNKS2_HUMAN 8.324102287 1.30E-05

c1090731_g2_i1 SHC1_XENLA 8.635804203 1.30E-05

c1081414_g1_i1 RB11A_RAT 8.132589277 1.39E-05

c1087547_g1_i1 DDR1_HUMAN 8.126206952 1.50E-05

c1083389_g1_i4 KLH11_MOUSE 6.35467149 1.59E-05

c1082905_g1_i4 PAR1_XENLA 8.272930557 1.62E-05

c1082925_g1_i6 S43A3_HUMAN -8.226900742 1.63E-05

c1070268_g1_i4 ZC3HF_HUMAN -6.145289331 1.72E-05

c1078260_g1_i5 PSS_HELPY -8.540663466 2.20E-05

c1086010_g2_i2 RAB4B_DANRE 8.225840078 2.33E-05

c1087417_g3_i1 IGF1R_HUMAN 8.496420151 2.71E-05

c1083067_g2_i1 CTR3_HUMAN 7.929698021 2.72E-05

c1074527_g1_i5 EXOC3_MOUSE 7.972740647 2.75E-05

c1071883_g2_i2 ITCH_MOUSE 8.012113684 2.75E-05

c1069151_g3_i1 MTMR5_MOUSE 8.62794095 3.20E-05

c1065062_g2_i2 S39AB_HUMAN 8.194534133 3.26E-05

c1086079_g1_i3 OBRG_RAT 8.10255092 3.36E-05

c1091771_g2_i1 LPIN2_HUMAN 8.203973099 3.39E-05

c1088042_g1_i3 MMP13_XENLA 8.432317627 3.66E-05

c1082633_g1_i2 LCAP_HUMAN 7.871333047 3.80E-05

c1081108_g2_i7 DDX5_PANTR 7.891088664 4.60E-05

c1055766_g1_i1 SEC13_BOVIN 8.524152917 6.15E-05

c1078442_g1_i2 LMCD1_HUMAN 7.987588424 6.31E-05

c1092570_g1_i1 HBA_TARGR -2.795007509 6.31E-05

c1080139_g3_i3 LIPA1_HUMAN 8.105917572 6.31E-05

c1078109_g9_i1 MEX3A_HUMAN 8.268070969 7.18E-05

c1055916_g2_i2 IKBZ_HUMAN 7.779741488 9.45E-05

c1078671_g2_i2 ALDOC_MACFA 8.521236252 9.47E-05

c1089493_g4_i2 ACK1_BOVIN 7.65273966 9.78E-05

c1077442_g1_i5 LYST_HUMAN 8.606270434 9.96E-05

c1083480_g1_i4 RGPA2_HUMAN -8.006862207 0.000101

c1080217_g1_i4 F172B_HUMAN 7.877170173 0.000104099

c1061180_g1_i2 IDE_BOVIN 7.673554829 0.000104099

181 Table A.3.1. (Continued)

transcript id blastx hit logFC padj

c1060498_g3_i1 NSL1_HUMAN -7.720681223 0.000104099

c1084240_g1_i4 S35B2_HUMAN 7.645495842 0.000105373

c1086935_g5_i1 SV422_XENLA -8.6390291 0.000108966

c1092563_g1_i3 YRD6_CAEEL 8.225508631 0.000126173

c1048841_g2_i1 LATS1_HUMAN 7.997389594 0.000129076

c1077756_g2_i7 PB1_HUMAN 7.289072999 0.000159256

c1080236_g3_i1 UBP4_HUMAN 7.79291751 0.000159256

c1089887_g2_i7 CJ012_HUMAN -8.160417673 0.000159256

c1089781_g1_i1 SPTC3_HUMAN 5.257520628 0.000163181

c1088438_g6_i2 DBF4B_HUMAN 8.028075621 0.000169023

c1089051_g1_i2 FUS_MOUSE 8.053062297 0.000171054

c1077796_g2_i2 CUL1_PONAB -8.547478627 0.000175025

c1089919_g4_i6 CP27B_HUMAN 5.65582147 0.000179832

c1085398_g1_i3 THEM4_XENLA 8.343035603 0.000187545

c1081108_g2_i8 DDX5_MACFA 8.264902784 0.000201884

c1089782_g7_i1 PPARD_CANFA 7.782377673 0.000203002

c1088063_g1_i2 RTJK_DROME 8.287448625 0.00021605

c1077943_g7_i1 NCOAT_HUMAN 7.457875205 0.000251284

c1073023_g3_i1 2A5E_HUMAN 8.033322571 0.000280192

c1088647_g1_i3 CAMP2_HUMAN 7.810268326 0.00030595

c1088902_g3_i1 LIN1_NYCCO -8.143726303 0.000319549

c1090534_g5_i2 ANR50_HUMAN 8.007365897 0.000327285

c1087639_g3_i1 SNX25_HUMAN 7.781326171 0.000327285

c1081183_g1_i1 SYRC_BOVIN -8.084362594 0.000338457

c1089348_g4_i3 KCC1A_RAT 7.542828134 0.000366421

c1083426_g7_i1 PRS35_MACMU -7.922666539 0.00037533

c1085289_g4_i3 TRI39_RAT 8.014620313 0.000393626

c1087211_g2_i3 MARF1_CHICK 8.422928762 0.000395116

c1073562_g2_i1 OSBL7_HUMAN 5.335409111 0.000404568

c1077195_g2_i1 KC1A_XENLA 7.338799444 0.000414958

c1084191_g3_i2 SMBT2_HUMAN 7.283901338 0.00058118

c1046138_g1_i2 GTAN_DICDI -3.035935157 0.000708374

c1086426_g1_i2 PO2F1_XENTR -7.385910348 0.000710994

182 Table A.3.2. Top 100 differentially expressed transcripts in stump-derived non- dividing cells of full skin flap sutured limbs. Negative fold change indicates enrichment in full skin flap sutured limbs.

transcript id blastx hit logFC padj

c1092764_g1_i2 RTJK_DROFU -5.539683228 1.07E-21

c1078994_g2_i6 CATK_RABIT 5.224015138 1.23E-20

c1088639_g4_i1 RRT15_YEAST -8.318969996 3.08E-20

c1078994_g2_i3 CATK_HUMAN 4.597527523 1.46E-19

c1680103_g1_i1 GP1_CHLRE -6.476805339 5.17E-18

c1082906_g1_i2 POL2_MOUSE 11.00237365 6.48E-15

c1057214_g1_i1 SN_HUMAN 3.32015022 2.91E-14

c1090300_g1_i1 CNN2_MOUSE 10.00028264 5.73E-12

c1092690_g1_i4 SACS_MOUSE 3.431589474 6.27E-12

c1082885_g1_i2 NSF_MOUSE 9.582646445 1.86E-10

c1081515_g3_i1 WNK3_HUMAN 9.292561444 4.09E-10

c1070626_g1_i3 ANO6_HUMAN 9.272540715 4.11E-10

c1065747_g3_i2 KCRB_CHICK 2.211294235 4.80E-10

c1086010_g2_i2 RAB4B_DANRE 9.425222703 4.80E-10

c1092570_g1_i1 HBA_TARGR -2.861455759 7.58E-10

c1053472_g1_i1 GBP1_CHLAE 2.242707772 8.03E-10

c1091277_g1_i5 LIN1_NYCCO 3.234902668 1.76E-09

c1052471_g1_i1 ELP1_HUMAN 8.993267195 1.76E-09

c1072152_g3_i2 HERC3_HUMAN 5.891029939 2.18E-09

c1078351_g1_i1 TP53B_HUMAN 5.722295876 3.28E-09

c1086318_g1_i3 SOAT1_HUMAN 9.226651468 3.36E-09

c1084683_g5_i2 N42L2_HUMAN 8.992956384 4.67E-09

c1090167_g1_i3 S38A3_HUMAN 8.922998113 4.88E-09

c1088436_g1_i3 BPHL_MOUSE -2.509414817 6.42E-09

c1079951_g1_i3 CL16A_HUMAN 9.064970347 6.60E-09

c1082738_g2_i2 IF2_KINRD -5.601338329 7.84E-09

c1076922_g10_i1 PXDN_HUMAN 9.024373475 8.91E-09

c1086644_g3_i11 CNOT1_HUMAN 8.914952716 9.34E-09

c1088054_g3_i1 LAMB2_HUMAN -8.743758423 1.68E-08

c1082693_g1_i3 PTPRE_HUMAN 9.126244302 1.70E-08

c1072251_g4_i1 NGAP_HUMAN 8.418655407 2.30E-08

c1078580_g2_i2 TT39A_XENTR 2.331938095 2.36E-08

c1077756_g2_i6 PB1_HUMAN 8.56969585 2.72E-08

183 Table A.3.2. (Continued)

transcript id blastx hit logFC padj

c1089216_g3_i4 SBNO2_BOVIN 8.983518514 2.72E-08

c1086263_g1_i1 LEMD2_MOUSE 8.696770124 4.56E-08

c1052471_g1_i3 ELP1_HUMAN -8.599629492 5.26E-08

c1089211_g9_i2 NDUS1_HUMAN 8.487125145 5.39E-08

c1085700_g1_i4 F107B_HUMAN 8.503133082 8.20E-08

c1085723_g3_i4 VATA_HUMAN 8.479739414 8.78E-08

c1075582_g1_i1 NU155_HUMAN 8.539617693 1.45E-07

c1086079_g1_i3 OBRG_RAT 5.712703792 1.57E-07

c1085524_g4_i1 SPTB2_HUMAN -8.575696084 1.57E-07

c1088042_g1_i2 MMP13_XENLA 3.849037515 1.74E-07

c1085700_g1_i1 F107B_HUMAN -8.480194288 1.74E-07

c1088299_g3_i11 HNRPU_HUMAN 8.35027298 1.75E-07

c1089084_g5_i1 TRHY_HUMAN 3.10660932 1.84E-07

c1090758_g5_i4 WDR82_CHICK 8.240547603 1.91E-07

c1064056_g1_i1 GP350_EBVB9 -8.414324826 2.56E-07

c1089084_g4_i1 PAR14_MOUSE 3.357960967 2.58E-07

c1090897_g2_i5 POLG_PVCV2 8.280234978 3.03E-07

c1081415_g2_i1 HERC5_HUMAN 2.719380072 3.14E-07

c1086799_g1_i3 SPT2_XENTR 8.399002755 3.30E-07

c1597296_g1_i1 POLY_DROME 3.287948164 3.31E-07

c1042145_g2_i2 UBL7_MOUSE -8.317234547 3.57E-07

c1085065_g1_i1 SETX_HUMAN -7.607067309 3.90E-07

c1081540_g4_i1 SPX2_XENLA 1.91882721 4.16E-07

c1084184_g1_i4 GMPBA_XENLA 8.356331549 4.17E-07

c1083484_g4_i1 SYWC_PONAB -6.427921108 4.28E-07

c1082127_g2_i4 HERC1_HUMAN 8.144953545 4.65E-07

c1089402_g2_i3 IRF3_CHICK 3.467059588 4.87E-07

c1084654_g3_i1 2AAA_PIG 8.33089139 5.35E-07

c1090770_g1_i3 HARB1_HUMAN -4.684961605 6.50E-07

c1085660_g2_i4 RXRBA_DANRE 8.586893266 6.54E-07

c1084500_g2_i5 PRAM_HUMAN 8.361954934 6.59E-07

c1084053_g1_i4 LRRK1_HUMAN 8.205707988 7.06E-07

c1090476_g2_i1 HCDH_PIG 8.052051755 8.20E-07

c979896_g1_i1 043R_FRG3G -8.693499665 8.45E-07

c1052055_g2_i1 RTJK_DROME 2.662018488 8.85E-07

c1085660_g2_i5 RXRBA_DANRE 7.173467873 8.85E-07

184 Table A.3.2. (Continued)

transcript id blastx hit logFC padj

c1080270_g3_i4 ZBT46_HUMAN 8.590389418 9.17E-07

c1078218_g3_i2 H4_XENTR -4.220526388 9.25E-07

c1089828_g2_i1 SMTN_HUMAN 6.234509963 9.25E-07

c1071943_g2_i1 MX1_MACMU 8.215122108 9.30E-07

c1091771_g2_i3 LPIN2_HUMAN 7.831749739 9.54E-07

c1092844_g1_i2 RTBS_DROME -2.102780869 1.03E-06

c1082664_g1_i4 ZO1_CANFA 8.392339556 1.12E-06

c1071771_g2_i1 IL11_HUMAN 2.919327428 1.13E-06

c1082749_g2_i5 RHG23_MOUSE -8.098960826 1.23E-06

c1085800_g1_i3 FA96A_HUMAN 7.954996382 1.33E-06

c1081560_g1_i1 EPDR1_MACFA 6.008417753 1.59E-06

c1064539_g2_i1 CCD93_CHICK 8.037942279 1.65E-06

c1042145_g2_i1 UBL7_MOUSE 7.862532113 1.65E-06

c1085431_g1_i4 DOCK9_HUMAN 6.382680704 1.75E-06

c1090666_g1_i1 PLEC_MOUSE 7.857563979 1.87E-06

c1089338_g4_i2 ST38L_HUMAN 7.987019409 2.35E-06

c1075965_g1_i3 GPR17_HUMAN 8.029837418 2.47E-06

c1081290_g1_i4 IF4G3_HUMAN -8.402370312 2.55E-06

c1072497_g2_i2 DEFI8_HUMAN 8.012857777 2.69E-06

c1084774_g1_i3 SEPT7_XENLA 7.957131692 2.69E-06

c1067485_g1_i1 HARB1_HUMAN -2.304221293 2.85E-06

c1082633_g1_i2 LCAP_HUMAN 7.539546147 2.86E-06

c1063882_g2_i1 MAST3_XENLA 7.967450769 2.87E-06

c1078350_g2_i1 SC31A_HUMAN 8.172470312 2.89E-06

c1086389_g2_i1 ARF1_RAT 7.836321346 3.59E-06

c1061972_g1_i1 ACTZ_RAT 7.886894718 3.83E-06

c1084682_g4_i3 ZN326_PONAB 7.724498639 3.94E-06

c1087610_g1_i1 DIA1_HUMAN 8.071491831 4.35E-06

c1073648_g4_i6 UBP2L_MOUSE 7.946903484 4.35E-06

c1085800_g1_i1 FA96A_HUMAN -8.062908379 4.64E-06

c1052724_g1_i2 CDK12_RAT -7.806816947 4.69E-06

185 Table A.3.3. Top 100 differentially expressed transcripts in epithelial cells of full skin flap sutured limbs. Negative fold change indicates enrichment in full skin flap sutured limbs.

transcript id blastx hit logFC padj

c1090257_g1_i2 TSN15_MOUSE 10.65357538 4.18E-12

c1090300_g1_i1 CNN2_MOUSE 10.72497754 1.18E-11

c1086285_g2_i3 K1C24_HUMAN 10.4413052 1.89E-11

c1092690_g1_i3 SACS_MOUSE 9.00189591 1.89E-11

c1088639_g4_i1 RRT15_YEAST -4.508234683 7.17E-11

c1084101_g1_i2 TB182_MOUSE 11.34428408 1.69E-10

c1090687_g2_i3 PTBP1_BOVIN 11.51147751 3.14E-10

c1058430_g3_i1 K1C15_RAT 3.821635653 3.90E-10

c1090600_g1_i6 GCSH_CHICK 5.628028962 3.42E-09

c1090064_g1_i3 RN5A_HUMAN 2.473571913 3.54E-09

c1088810_g1_i3 CO7A1_HUMAN 9.872322957 3.92E-09

c1068399_g1_i2 LAMB3_HUMAN 9.327438602 4.22E-09

c1084191_g3_i2 SMBT2_HUMAN 7.030765253 1.18E-08

c1075471_g2_i4 TOP2A_CHICK 2.78171849 1.92E-08

c1086457_g2_i4 PKN2_MOUSE 9.674315484 2.29E-08

c972758_g1_i1 POLR_DROME 9.317001622 3.06E-08

c1083705_g1_i2 UN13D_HUMAN -9.054962123 3.79E-08

c1091613_g1_i2 TNR6A_HUMAN 8.994633953 5.36E-08

c1086872_g4_i3 CYTC_RABIT -9.452696071 6.09E-08

c1092079_g1_i2 PANX1_MOUSE -9.668498735 8.97E-08

c1461347_g1_i1 MUC5A_HUMAN -4.201898825 1.03E-07

c1066879_g2_i1 TRI33_HUMAN 8.731716605 1.11E-07

c1087857_g1_i1 MYO1C_CHICK 9.293334538 1.13E-07

c1082467_g1_i2 ZBT11_HUMAN 8.730748092 1.15E-07

c1087476_g1_i1 MUC5B_HUMAN -3.227012641 1.22E-07

c1081555_g1_i5 ETUD1_HUMAN -8.823319198 1.22E-07

c1091235_g1_i5 PRRT3_HUMAN 8.90666019 1.57E-07

c1090681_g1_i3 FREM2_HUMAN 5.254949056 1.60E-07

c1085139_g1_i4 PRDX4_HUMAN 8.727004412 2.18E-07

c1090393_g1_i4 HMGN2_PONAB 1.820350319 2.18E-07

c1092564_g1_i4 JADE2_MOUSE -9.225665615 2.24E-07

c1079364_g6_i4 BRD7_CHICK -9.498102575 2.24E-07

c1091539_g2_i2 POL_WDSV 9.100413462 3.02E-07

186 Table A.3.3. (Continued)

transcript id blastx hit logFC padj

c1086995_g1_i2 FDFT_PONAB -8.895810454 5.76E-07

c1075095_g5_i2 MCATL_HUMAN 8.831980129 7.18E-07

c1090795_g1_i1 MSPD1_MOUSE 8.813490009 9.40E-07

c1088527_g2_i2 NOC2L_BOVIN 8.329759162 1.04E-06

c1082467_g1_i1 ZBT11_HUMAN -8.569030977 1.28E-06

c1086350_g2_i3 LMA2L_HUMAN 8.681389279 1.56E-06

c1092813_g3_i1 ANXA1_PANTR 6.826599362 1.70E-06

c1085797_g1_i1 PKH4B_HUMAN 8.441934763 2.97E-06

c1070684_g6_i4 TM100_BOVIN -9.407323215 3.22E-06

c1091969_g1_i2 ATPD_MOUSE 9.182073868 3.23E-06

c1085994_g1_i2 SMAP2_CHICK -8.670961134 3.45E-06

c1082222_g1_i3 RBM14_PONAB -8.422111345 3.51E-06

c1073835_g3_i1 PLEK2_MOUSE 2.672156028 3.59E-06

c1081608_g4_i3 CO2_BOVIN 4.629448203 3.63E-06

c1074983_g4_i1 HID1_HUMAN 8.108459398 5.23E-06

c1083368_g1_i1 RENBP_PIG 8.173149933 8.67E-06

c1088538_g2_i2 PCDH9_HUMAN 7.921943784 9.61E-06

c1049724_g1_i1 DYL1_RAT 3.453071342 1.04E-05

c1081071_g2_i3 CNTLN_HUMAN 7.984604595 1.44E-05

c1090670_g1_i1 GPD1L_XENTR 9.3193441 1.59E-05

c1082738_g2_i2 IF2_KINRD -4.807542884 2.02E-05

c1079696_g1_i2 GAPD1_HUMAN 8.199232592 2.11E-05

c1085156_g5_i2 TGM3_MOUSE 5.789571912 2.14E-05

c1064453_g2_i3 CSKP_HUMAN 8.024329933 2.14E-05

c1078208_g3_i7 AF17_HUMAN 8.662035048 2.21E-05

c1064716_g4_i2 P66A_HUMAN 9.047977398 2.26E-05

c1079507_g1_i2 STAT1_HUMAN 8.728821489 2.70E-05

c1090558_g1_i3 SZT2_HUMAN 8.458554452 2.86E-05

c1089406_g2_i1 SOGA2_MOUSE 5.810787769 2.87E-05

c1081498_g2_i1 HOOK2_XENLA 8.075631981 2.90E-05

c1088192_g4_i4 OASL2_RAT 2.150590569 3.02E-05

c1091613_g1_i3 TNR6A_HUMAN -2.628152433 3.02E-05

c1089983_g3_i1 I20RA_HUMAN 8.317687135 3.08E-05

c1075955_g1_i3 FBX28_HUMAN 7.833857986 3.48E-05

c1078595_g4_i3 DRAM1_MOUSE 5.349510356 4.25E-05

c1082870_g1_i3 COR2A_HUMAN 8.101613637 4.36E-05

187 Table A.3.3. (Continued)

transcript id blastx hit logFC padj

c1080135_g1_i1 IBP2_HUMAN 3.279577401 4.45E-05

c1083265_g7_i3 CAV1_DIDVI -8.159187434 4.74E-05

c1080732_g6_i1 TPC11_CHICK 8.131283847 5.46E-05

c1041958_g1_i2 LIN1_NYCCO 8.510829891 5.94E-05

c1085634_g3_i5 DYN2_MOUSE 8.010511729 6.54E-05

c1084825_g1_i7 RFXK_HUMAN -7.753124731 6.73E-05

c1045890_g1_i3 CPE1B_XENLA 8.051298363 6.94E-05

c1066879_g2_i2 TRI33_HUMAN -6.891767024 7.58E-05

c1086414_g4_i3 RBL2_HUMAN -7.893663368 7.68E-05

c1087059_g7_i1 PP1B_XENTR 8.272181411 8.00E-05

c1065332_g1_i4 MUC1_YEAST 8.21653022 8.00E-05

c1091000_g1_i3 RAD17_HUMAN 8.057187637 8.00E-05

c1085670_g3_i4 C1GLC_HUMAN 8.709107806 8.15E-05

c1072053_g1_i1 UH1BL_XENLA 7.79363608 8.75E-05

c1091852_g2_i2 LAMA5_HUMAN 3.302416849 9.10E-05

c1062596_g1_i1 CP135_HUMAN -7.709965269 9.19E-05

c1081740_g1_i2 AGAP3_HUMAN 7.198650411 9.40E-05

c1043485_g1_i1 BCL3_HUMAN 7.974986889 9.60E-05

c1074926_g1_i1 HTF4_MOUSE 7.989655032 9.71E-05

c1080285_g4_i4 TXIP1_MOUSE 7.934141155 0.000100146

c1079147_g1_i1 ENPP4_HUMAN 7.907390328 0.000101543

c1068313_g1_i1 AT7L3_HUMAN 8.237068526 0.000101597

c1075036_g3_i9 PKN1_THETN 7.689978039 0.000103994

c1069508_g1_i1 CAN15_MOUSE 8.101987358 0.000118447

c1079985_g4_i3 GFPT1_RAT 9.059151021 0.000122688

c1092764_g1_i2 RTJK_DROFU -3.598426273 0.0001239

c1076698_g3_i9 HNRH3_HUMAN 1.745882183 0.000126679

c1057236_g1_i2 IF2M_BOVIN 6.340511482 0.000128845

c1086934_g2_i4 CTL1_XENLA 1.927666798 0.000151422

c1069931_g5_i2 SORT_HUMAN 2.810402884 0.000161274

c1082163_g1_i1 SIAE_HUMAN 2.341915853 0.000162676

188 A.4. Supplemental materials for Chapter 5

Table A.4.1. Top 100 differentially expressed genes in DMSO/iMDK treatments. Negative fold change indicates overexpression in iMDK, while positive fold change indicates overexpression in DMSO.

transcript id blastx hit logFC padj

c1069619_g1_i1 UROM_BOVIN -3.219397053 3.78156E-63

c1067192_g2_i1 GILT_BOVIN -10.19916006 9.33083E-60

c1068735_g2_i3 S10AA_CHICK -10.77421892 3.56393E-52

c1070122_g1_i1 CP2J6_MOUSE -6.199446518 2.68918E-41

c1088042_g1_i4 MMP13_XENLA -9.385904463 1.80146E-38

c1074092_g3_i8 FRI1_LITCT -5.871716127 1.39528E-35

c1078497_g1_i1 CTNA1_HUMAN 2.071273079 4.98319E-35

c1078879_g1_i2 PORIM_HUMAN -2.634626841 4.05581E-32

c1091411_g5_i2 MMP18_XENLA -14.95597225 4.37216E-31

c1083466_g1_i2 C560_HUMAN 1.632854145 5.6542E-28

c1053558_g1_i2 LYSC_RABIT 4.610939561 1.58403E-25

c1076999_g1_i4 SC61G_MOUSE 10.35400803 1.06845E-24

c1090676_g2_i2 TSN_CHICK 13.39554876 1.06845E-24

c1079944_g4_i2 CS052_HUMAN 3.867207416 1.03408E-23

c1067192_g2_i2 GILT_BOVIN 12.71681457 1.45775E-22

c1084825_g1_i8 RFXK_HUMAN -6.347781905 1.78437E-22

c1083302_g7_i1 DCNL3_XENTR -2.035811211 6.34729E-22

c1076874_g1_i2 RTBS_DROME 8.987582433 1.30276E-21

c1083265_g7_i2 CAV1_DIDVI -13.37500174 3.28467E-21

c1084989_g2_i2 M1IP1_HUMAN 12.33780997 1.23983E-20

c1073457_g2_i1 EMD_MOUSE 12.0649769 4.90699E-20

c1085679_g4_i1 MSS4_HUMAN 6.42551468 2.68107E-19

c1089923_g1_i2 SNED1_MOUSE 5.665457024 8.04772E-19

c1089890_g4_i3 MMP18_XENLA -3.711999065 1.0831E-18

c1075788_g1_i2 XPO5_HUMAN 11.41163528 1.55131E-18

c1077954_g1_i2 ABCB9_HUMAN -1.034214966 3.00419E-18

c1086055_g1_i6 DHE3_HUMAN -11.86856518 7.84699E-18

c1080065_g1_i3 LEG8_HUMAN -11.36980653 1.92933E-17

c1088757_g3_i5 TM131_HUMAN -11.45019448 6.8566E-17

c1085204_g1_i3 KDEL2_HUMAN -11.20444188 9.65964E-17

189 Table A.4.1. (Continued)

transcript id blastx hit logFC padj

c1091725_g1_i1 URP2_BOVIN -2.053239703 9.75928E-17

c1089244_g1_i4 FA76B_XENTR -10.90772373 1.01134E-16

c1081121_g1_i1 FA98A_RAT -10.98267709 1.19553E-16

c1082824_g3_i4 ITA6_CHICK 11.36884087 1.35983E-16

c1068880_g1_i2 MYPT1_HUMAN 11.44610513 1.35983E-16

c1071947_g1_i1 CDN1A_HUMAN -1.98396188 2.6498E-16

c1088286_g4_i10 PAQR3_HUMAN -10.84476358 2.76186E-16

c1083894_g1_i1 M17L2_XENTR 1.931352877 4.38619E-16

c1089291_g1_i2 WAP1_PHIOL -10.81864107 4.38619E-16

c1079790_g3_i6 CCNC_XENLA -10.87727809 4.98414E-16

c1084825_g1_i11 RFXK_HUMAN -7.557935763 4.98414E-16

c1090907_g1_i7 NCPR_HUMAN -10.61632445 8.8038E-16

c1084227_g2_i6 BIRC5_BOVIN -10.76925458 2.49203E-15

c1085389_g6_i4 LECT2_BOVIN -10.54025637 2.77683E-15

c1089123_g1_i3 IF44L_HUMAN -11.02802577 3.53209E-15

c1077347_g1_i3 CYSP2_HOMAM -10.74070124 4.0995E-15

c1083836_g2_i1 CATK_PIG 10.48632805 4.10968E-15

c1085967_g1_i3 SELN_XENTR 11.12599873 6.71498E-15

c1085376_g2_i1 CNFN_DANRE -10.23908817 8.94011E-15

c1081919_g1_i2 YAP1_HUMAN -10.34615329 9.42223E-15

c1088286_g4_i11 PAQR3_HUMAN 10.32537503 9.63016E-15

c1075330_g2_i1 PSB6_BOVIN 8.060955837 1.08778E-14

c1086071_g1_i5 CT027_XENTR 11.10453444 1.08778E-14

c1088175_g1_i2 PIGS_HUMAN 10.39762999 1.15038E-14

c1083925_g7_i1 UVS2_XENLA -1.94637579 1.18334E-14

c1070847_g1_i7 TM154_HUMAN -10.39651616 1.49751E-14

c1089890_g2_i1 MMP3_RABIT -4.621387099 1.61894E-14

c1090167_g1_i2 S38A3_HUMAN -10.69160638 1.90694E-14

c1080762_g13_i1 CWC15_PONAB -10.29209096 2.58901E-14

c1088267_g2_i14 GIT1_HUMAN -10.14817849 2.85131E-14

c1053012_g1_i6 NR2CA_XENLA -10.31585894 3.17019E-14

c1071002_g3_i4 JAK2_PIG -10.38363307 3.53309E-14

c1068761_g1_i1 AKA10_HUMAN -10.70341903 3.549E-14

c1081431_g2_i2 GPBL1_PONAB -3.034868233 4.68409E-14

c1053012_g1_i1 NR2CA_XENLA 10.46561074 7.51441E-14

190 Table A.4.1. (Continued)

transcript id blastx hit logFC padj

c1088059_g1_i6 EIF1_PONAB -10.53258588 8.90044E-14

c1089890_g5_i1 MMP18_XENLA -3.916075813 1.13595E-13

c1084227_g2_i2 BIRC5_BOVIN 9.938323162 1.94034E-13

c1078066_g1_i2 DPOE3_MOUSE -10.09161797 2.16255E-13

c1088852_g2_i3 NO66_DANRE 1.495251756 2.31134E-13

c1088814_g1_i5 AP3D1_MOUSE 10.39222766 2.95368E-13

c1069931_g5_i1 SORT_HUMAN -10.75369787 3.53546E-13

c1077291_g1_i1 CMLO_XENLA 10.12525054 4.05188E-13

c1091849_g1_i5 HBA2_TRICR 9.708006092 5.77383E-13

c1076139_g2_i1 SPT5H_HUMAN 5.691303182 6.57351E-13

c1067457_g2_i1 RCD1_RAT 10.5666141 6.83102E-13

c1081877_g1_i2 VASP_BOVIN -9.776314241 8.51358E-13

c1084443_g4_i3 S10AD_HUMAN -1.863345078 9.01461E-13

c1084825_g1_i3 MF2NB_DANRE 9.642926766 1.07715E-12

c1075515_g1_i8 NRBP_MACFA -9.766074109 1.21711E-12

c1084478_g4_i7 FANCL_HUMAN -9.669065633 1.61304E-12

c1072722_g1_i8 HYEP_RABIT -11.65799646 1.81998E-12

c1075002_g1_i2 DDAH1_MOUSE -9.647187847 2.02255E-12

c1071544_g1_i2 MOT7_MOUSE 9.809284959 3.11762E-12

c1089890_g4_i1 MMP18_XENLA -1.824314992 3.162E-12

c1065650_g1_i3 WDR3_HUMAN -11.00469068 3.87903E-12

c1090907_g1_i8 NCPR_HUMAN 1.081226169 4.43617E-12

c1067048_g2_i2 PRRX1_RAT -1.942357961 5.51735E-12

c1089079_g2_i2 ADPRH_MOUSE -1.513490616 6.32262E-12

c1085444_g1_i1 ZBT34_HUMAN 2.203222094 7.08975E-12

c1089402_g2_i3 IRF3_CHICK 10.07652743 8.62822E-12

c1076882_g1_i6 CV039_XENLA 2.927742586 8.91177E-12

c1088124_g1_i2 RS21_PIG 10.48571745 9.9563E-12

c1075515_g1_i4 NRBP_MACFA 9.464209578 1.03413E-11

c1088097_g2_i7 TX1B3_HUMAN -9.379024741 1.14552E-11

c1077595_g1_i2 FUND2_HUMAN 2.941866219 1.25903E-11

c1076557_g4_i1 FMO3_CANFA -9.588270973 1.53042E-11

c1081781_g1_i3 HYES_PIG -9.334678354 1.81735E-11

c1089079_g2_i1 ADPRH_MOUSE 9.666861008 1.9689E-11

c1075822_g5_i3 CP2C8_HUMAN -4.056441733 2.43533E-11

191 Table A.4.2. Select enriched pathways in iMDK-treated limbs. Enriched pathways -log(p-value) HIF1α Signaling 4.51 IL-8 Signaling 3.79 Leukocyte Extravasation Signaling 3.61 Agranulocyte Adhesion and Diapedesis 3.47 Granulocyte Adhesion and Diapedesis 3.14 Integrin Signaling 2.93 Th1 Pathway 2.89 TREM1 Signaling 2.51 Th1 and Th2 Activation Pathway 2.48 STAT3 Pathway 2.41 Production of Nitric Oxide and Reactive Oxygen Species in 2.34 Macrophages Oncostatin M Signaling 2.16 T Cell Exhaustion Signaling Pathway 2.07 Toll-like Receptor Signaling 1.82

Table A.4.3. Select enriched pathways in DMSO-treated limbs. Enriched pathways -log(p-value) β-alanine Degradation I 3.54 Valine Degradation I 2.48 RhoA Signaling 2.29 Protein Kinase A Signaling 2.06 Actin Cytoskeleton Signaling 1.75 Regulation of Actin-based Motility by Rho 1.75 Calcium Signaling 1.6 Cell Cycle: G1/S Checkpoint Regulation 1.57 Thrombin Signaling 1.55 RhoGDI Signaling 1.52

192

Figure A.4.1. Mk is not expressed in monocytes. (A) Double RNAscope in situ hybridization of mk and csf1r at 5 dpa reveals discrete expression patterns. Arrowheads denote mk+ only cells and arrows denote csf1r+ only cells. (B) Quantification of mk+ only, csf1r+ only, and co-positive cells. Scale bar in A denotes 50 µm.

Figure A.4.2. Validation of custom polyclonal rabbit anti-MK antibody. Western blotting was performed with increasing volumetric ratios of MK antibody: blocking peptide ratios (1:0, 1:0.5, 1:1, and 1:10) on 10 dpa extracts revealed lower levels of staining with increased levels of peptide.

193

Figure A.4.3. PCA analysis of DMSO and iMDK samples.

Figure A.4.4. Mk targeting strategy for mk mutant generation. The targeting gRNA is shown in pink. The exon is denoted in yellow. An example genotyping result from deep sequencing of PCR amplicons of an mk null mutant reveals no wildtype alleles present.

194

Figure A.4.5. Validation of pCAG-MK overexpression construct. Western blots of either pCAG-tdTomato or pCAG-MK transfected cells reveals the overexpression and secretion of axolotl MK in transfected cell lysates and media.

195