The Forkhead Box F1 Transcription Factor in Disease and Development

A dissertation submitted to the Graduate School of the University of Cincinnati in partial fulfillment of the requirements to the degree of

Doctor of Philosophy (Ph.D.)

In the Department of Cancer and Cell Biology of the College of Medicine

Fall, 2018

Hannah Marie Flood

B.S. Allegheny College, 2010 M.S. Murray State University, 2012

Dissertation Committee: Vladimir V. Kalinichenko, M.D., Ph.D. (Chair) Chunying Du, PhD David Plas, PhD Nikolai Timchenko, PhD Chunyue Yin, PhD

Abstract

The forkhead box F1 transcription factor (FOXF1) is a mesenchymal-specific transcription factor and is expressed in mesenchyme-derived cells such as hepatic stellate cells and lung microvascular endothelial cells. FOXF1 functions are critical for transcriptional regulation during development of the liver, lung, and other organs, and has numerous roles in adult diseases. Here, we investigate FOXF1 and its role in hepatic fibrosis and its role in endothelial progenitor cells. Studying FOXF1 throughout different systems will enhance our knowledge base and can be used to develop further studies in a variety of systems.

FOXF1 is expressed in the collagen-producing cells of the liver, the hepatic stellate cells

(HSC), and HSC activation to myofibroblasts (MFs) requires FOXF1 presence. Therefore, our studies utilize a carbon tetrachloride-induced liver injury model to investigate the role of FOXF1 during liver fibrosis progression. We found that Foxf1 deletion increased collagen depositions and disrupted liver architecture. Timp2 expression was significantly increased in Foxf1-deficient mice while MMP9 activity was reduced. RNA sequencing of purified liver myofibroblasts demonstrated that FOXF1 inhibits expression of pro-fibrotic genes: Col1α2, Col5α2, and Mmp2 in fibrotic livers and binds to active repressors located in promotors and introns of these genes.

Overexpression of FOXF1 inhibits Col1a2, Col5a2, and MMP2 in primary murine HSCs in vitro. In summary, we found that FOXF1 prevents aberrant extracellular matrix depositions during hepatic fibrosis by repressing pro-fibrotic gene transcription in HSCs and MFs.

FOXF1 is additionally expressed in endothelial cells, which line blood vessels. FOXF1 in an important regulator of vascular formation and has roles in cellular proliferation. Therefore, our studies utilized endothelial colony forming cells (ECFC) to rescue mice with acute lung injury (ALI). We found a 53.8% increase in the survival of mice with ALI, which was associated

ii with a decrease in lung injury; however, further evaluation of the injected ECFCs revealed no fluorescent tracer or FOXF1 expression. We therefore developed a novel embryonic stem cell

(ESC) line with GFP:FOXF1 to both trace FOXF1 expression through differentiation to endothelial progenitor cells (EPC) in vitro and to trace the cell in vivo for future studies. Our novel differentiation method yields ~95.2% endothelial cells, 51.5% of which are FOXF1- positive. In summary, we have successfully developed a novel ESC line and a novel EPC differentiation protocol to be used in future studies.

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Acknowledgements

First and foremost, I need to thank my mentor, Vladimir Kalinichenko. You have been the most excellent mentor to me these past few years, and I have learned so many new things from working in your lab. I feel as though I was lucky to have landed in your office during that rotation meeting my first year, and I am so grateful you allowed me to join your lab. You have put together such an accomplished, phenomenal group of scientists who are also good people, and that has made all the difference in the encouragement I get in lab. Your support of my research ideas and professional goals remains unparalleled. I am thankful that you allowed me to take time away from research during more trying times in my personal life and that you allowed me time to recover once I returned to lab. Thank you.

To my labmates, thank you all for providing such a pleasant work environment for the past 5 years. I have enjoyed every one of you, but I would like to especially thank the person who trained me during my rotation, and then never stopped helping me, Craig Bolte. You answered every question and help me learn most of the experiments I did in the lab. I would additionally like to thank the people I sit between, Yufang Zhang and Arun Pradhan. You have not only helped me with lab work, but you have given me so much good life advice, including,

“Drinking coffee is like a puromycin selection: You start with 3mg, then maintain with 1mg,” –

Arun. To the graduate students in my lab, thank you for helping me with experiments, for discussing research and literature with me, for helping me blow off steam and listening to my rants, for making sure I remembered class times, and for being irreplaceable companions as we trudged through this long process together. Good luck to you all.

To my committee, you have been the kindest, most helpful committee a graduate student could have. When I describe you to other students, they get jealous of how amazing and

v supportive you have been and still are as I complete my studies and move on past graduate school. Thank you for everything.

To my class in the cancer biology program, you have truly been the most amazing group.

I know I will be able to count your support and friendships throughout my life and I am excited to see what the future holds for us all. Thank you Jordan, Nicole, Molly, and Sonya.

To the people I have met in Cincinnati, graduate students, scientists, muggles, no matter how we met, please know I have loved every moment. A place is only as good as the people you know in it, and you have all made Cincinnati pretty great.

To my friends outside of Cincinnati, you continue to amaze me with your friendships as I have gone through both personal and professional hardships. Thank you for your understanding and continuous support. I need to especially thank my best friends, Amanda Colvin Zielen and

Juanita Von Dwingelo, and additionally Emily D’Angelo and Mara Varvil. We are all geographically far apart but you have made it so easy to get together through the years.

I especially need to thank Josh, for your love and support and for dealing with my crazy.

I would like to dedicate this work to my family. I could not have done any of this without your support and understanding, and of course, your love. To Papa, Jacob, Grace, Alyssa, Victor,

Maxwell, and to the entire Flood Family, thank you for never asking when I would graduate.

The following dissertation is presented in memory of my grandfather, Gordon Arnett

Flood: the first to hold a doctorate in the Flood Family, and my mother, Patricia Elizabeth Flood: who loved me, who was so proud of me, who could solve all my problems in the simplest ways, and who I will always miss most at 10:20PM.

Table of Contents

Abstract ...... ii Acknowledgements ...... v List of Tables and Figures ...... x Chapter 1: Introduction ...... 1 Forkhead Box Family and FOXF1 ...... 2 FOXF1 during development ...... 4 FOXF1 in Diseases ...... 6 Alveolar capillary dysplasia with a misalignment of pulmonary veins ...... 6 Acute lung injury and acute respiratory distress syndrome ...... 8 Pulmonary and Hepatic Fibrosis ...... 9 Other diseases (Barret’s esophagus, Fanconia anemia) ...... 11 FOXF1 and Cancer ...... 12 Lung ...... 14 Gastrointestinal (Colorectal, gastric, liver) ...... 14 Breast ...... 16 Other cancers (Prostate, rhabdomyosarcoma) ...... 17 Summary ...... 18 Experimental Rationale ...... 19 Refereneces ...... 21 Chapter 2: The Role of FOXF1 in Hepatic Fibrosis ...... 29 Literature Review: The Liver and Hepatic Fibrosis ...... 30 Abstract ...... 38 Introduction ...... 39 Results ...... 41 Deletion of Foxf1 in αSMA-positive Cells Exacerbates CCl4-Induced Hepatic Fibrosis...... 41 FOXF1 Expression is Decreased in Hepatic Myofibroblasts of αSMACreER;Foxf1-/- Mice...... 44 Deletion of Foxf1 Reduces MMP9 Activity in CCl4-Injured Livers...... 44 Deletion of Foxf1 Does Not Influence Cellular Proliferation in Fibrotic Livers...... 48 RNA-seq Analysis Identified Direct FOXF1 Target Genes Critical for ECM Deposition and Hepatic Fibrosis...... 51 Discussion ...... 57 Methods...... 60

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References ...... 64 Supplementary Information ...... 69 Chapter 3: Mouse Embryonic Stem Cells and Endothelial Progenitor Cells ...... 87 Literature Review: Endothelial Cells, Endothelial Progenitor Cells, and Lung Regenerative Medicine ...... 88 References ...... 94 Chapter 3A: Transplantation of Endothelial Colony Forming Cells Improves Survival of FOXF1- deficient Mice ...... 96 Abstract ...... 97 Introduction ...... 98 Results ...... 100 Endothelial FOXF1 deletion-dependent ALI and death rescued by ECFCs...... 100 Pulmonary architecture is maintained and lung injury is attenuated after ECFC transplantation...... 100 No integration of transplanted ECFCs was observed after ALI...... 103 Endothelial architecture is maintained after ECFC transplantation with no prevention of macrophage infiltration...... 103 FOXF1 expression is decreased in PdgfbCreER;Foxf1-/- mice with or without ECFC transplantation...... 106 Discussion ...... 108 Methods...... 110 References ...... 113 Chapter 3B: Differentiation of Novel GFP:FOXF1 Embryonic Stem Cell Line into FOXF1-positive Endothelial Progenitor Cells ...... 116 Abstract ...... 117 Introduction ...... 118 Results ...... 120 Novel CRISPR/Cas9 embryonic stem cell line expresses GFP to track FOXF1 expression .... 120 Novel GFP:FOXF1 ESC line maintains stemness characteristics ...... 120 Differentiation of EPCs from ESCs requires activation of vascular progenitor cell pathways 120 Highly efficient EPC differentiation ...... 124 Discussion ...... 127 Methods...... 131 References ...... 134 Supplementary Information ...... 137

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Chapter 4: Discussion and Future Directions ...... 142 Liver ...... 143 ESCs ...... 146 References: ...... 151

List of Figures and Tables

Figure 1.1. FOXF1 protein ...... 3 Figure 2.0. FOXF1 activity during liver injury ...... 33 Figure 2.1. Hepatic fibrosis is increased after CCl4 injury in mice with FOXF1 deficiency ...... 43 Figure 2.2. αSMA-CreER effectively deletes Foxf1 from hepatic myofibroblasts ...... 45 Figure 2.3. Deletion of Foxf1 from myofibroblasts increases liver fibrosis and inhibits MMP9 activity ...... 47 Figure 2.4. Deletion of Foxf1 does not influence proliferation of hepatic myofibroblasts ...... 50 Figure 2.5. FOXF1 deletion alters expression of pro-fibrotic genes in hepatic myofibroblasts ...... 53 Figure 2.6. FOXF1 binds to DNA regulatory regions of Col1α2, Col5α2, and Mmp2 ..56 Supplemental Figure 2.1. FOXF1 expression in mouse livers ...... 71 Supplemental Figure 2.2. Treatment with tamoxifen alone does not induce hepatic fibrosis ...... 73 Supplemental Figure 2.3. Number and percentage of FOXF1+ myofibroblasts are reduced in αSMACreER;Foxf1-/- livers ...... 74 Supplemental Figure 2.4. Increased collagen deposition in Foxf1-deficient livers ...... 75 Supplemental Figure 2.5. Deletion of Foxf1 had no effect on serum protein or bilirubin levels ...... 76 Supplemental Figure 2.6. Collagen accumulation in Foxf1-deficient livers is time-dependent ...... 77 Supplemental Figure 2.7. Widespread hepatic fibrosis and appearance of liver tumor in αSMACreER;Foxf1-/- mouse after 18-weeks of CCl4 treatment ...... 78 Supplemental Figure 2.8. Deletion of Foxf1 does not affect Mmp8, Mmp9, Mmp13, Mmp16, Timp1, or Timp3 mRNAs in CCl4-treated livers ...... 79 Supplemental Figure 2.9. Deletion of Foxf1 does not affect proliferation of αSMA+ cells in CCl4-treated livers...... 80 Supplemental Figure 2.10. Purified stromal cells express Acta2 and Des ...... 81 Supplemental Figure 2.11. ChIP-seq shows FOXF1 binding sites in DNA regulatory regions of Col1α2, Col5α2, and Mmp2 ...... 82 Supplemental Figure 2.12. Full images for Western blot and zymography ...... 84 Supplemental Table 2.1. List of TaqMan probes used in qRT-PCR analysis ...... 85 Supplemental Table 2.2. FOXF1 binding sites as identified by ChIP-seq relative to the transcriptional start site ...... 86

Figure 3.0. Differences and Similarities between embryonic stem cells and induced pluripotent stem cells ...... 91 Figure 3.1. Introduction of ECFCs increases overall mouse survival after FOXF1- deficiency-induced acute lung injury ...... 101 Figure 3.2. ECFC transplantation attenuates pulmonary injury ...... 102 Figure 3.3. ECFCs integration was not observed in FOXF1-deficient lung ...... 104 Figure 3.4. ECFC transplantation allows for maintenance of endothelial architecture but did not reduce number of macrophages ...... 105 Figure 3.5. FOXF1 levels are not restored after ECFC transplantation ...... 107 Figure 3.6. Establishment of GFP:FOXF1 ESC line by CRISPR/Cas9-mediated knock-in ...... 121 Figure 3.7. Novel GFP:FOXF1 (A1) cell line expresses GFP ...... 122 Figure 3.8. Novel GFP:FOXF1 (A1) cell line is similar in morphology and stemness marker expression to parental (W4) cell line ...... 123 Figure 3.9. Generation of EPCs from ESCs through novel differentiation protocol ....125 Figure 3.10. High-yield differentiation of EPCs from ESCs ...... 126 Supplemental Figure 3.1. ECFC analysis of tdTomato tracker ...... 138 Supplemental Figure 3.2. PCR Screen for GFP:FOXF1 knock-in cell lines ...... 139 Supplemental Figure 3.3. Sequence of Non-GFP allele of heterozygous GFP:FOXF1 clones identified in PCR screen compared to Wild Type (WT) Foxf1 ...... 140 Supplemental Figure 3.4. Flow cytometry gating strategy for EPC analysis ...... 141 Figure 4.1. FOXF1 activity during the progression of liver injury ...... 144 Figure 4.2. Proposed studies to test GFP:FOXF1+ EPCs ...... 149

Chapter 1: Introduction

The Forkhead Box F1 Transcription Factor in Development and Disease

Forkhead Box Family and FOXF1

The forkhead box superfamily of transcription factors are characterized by their evolutionarily conserved DNA-binding domain [1-4]. The forkhead box (Fox) domain is approximately 100 amino acids in length [1, 2]. The Fox domain is also known as a winged-helix domain as it contains two wing-like loops and three α-helixes and are a subclass of the helix- turn-helix class of proteins [5, 6] (Fig. 1.1 A). Fox proteins transcriptionally regulate genes involved with biological and physiological processes such as immune response, embryogenesis, organogenesis, tissue repair, cell cycle progression, cell metabolism, survival, proliferation, and differentiation [7-12]. Fox proteins maintain cellular homeostasis and mutations or expression alterations result in disease and cancer development and progression [9, 13, 14].

The human FOXF1 gene encodes a homolog of the mouse Foxf1 transcription factor previously known as Freac-1 and Hfh-8 [15] (Fig. 1.1 B). At 379 amino acids, FOXF1 is one of the smallest members of the Fox family of transcription factors with only two exons, the first containing the winged helix DNA-binding domain [16]. Foxf1 is located at the 16q24.1 chromosomal region (Fig. 1.1 C) and is expressed in the mesenchyme of the developing and adult mouse [16, 17]. FOXF1 is a mesenchymal-specific transcription factor and is expressed in mesenchyme-derived cells such as lung microvascular endothelial cells, hepatic stellate cells, peribronchial smooth muscle cells, and fibroblasts [18] and its functions are additionally crucial for formation of mesoderm-derived tissues [19]. FOXF1 is an important transcriptional regulator during development of the lung, liver, gall bladder, esophagus, and trachea [18, 20, 21]. FOXF1 has numerous roles during development and in adult diseases. Mutations or haploinsufficiency of

FOXF1 is linked to a variety of pathologies such as alveolar capillary dysplasia/misalignment of

Figure 1.1. FOXF1 protein. (A) Schematic of two helix-turn-helix binding domains interacting with DNA. Image from . (B) Image showing the two-exon human FOXF1 gene. The green boxes indicate the open reading frame and the grey boxes indicate the untranslated region. (C) Chromosome 16 ideogram with the position of FOXF1 at 16q24.1 indicated with a green arrow. Image modified from .

pulmonary veins [22], lung edema [23], as well as abnormal repair after injury in both the lung

[24] and liver [25].

FOXF1 during development

FOXF1 expression initiates during mouse gastrulation on embryonic day e7 pc in the mesoderm, allantois, and lateral mesoderm that arises from the primitive streak region [11, 26].

Loss of FOXF1 is embryonic lethal due to defects in the yolk sac and allantois before the gastrulation stage when the body plan of the vertebrate embryo is established, and embryos are reabsorbed by e10 [19]. Humans heterozygous for FOXF1 are embryonic lethal; however, a subset of mice heterozygous for Foxf1 will survive to adulthood [19, 21, 27]. Perinatal mice heterozygous for Foxf1 with low levels of the protein display defects in pulmonary capillary and alveolar morphogenesis and total lung function whereas heterozygous mice with wild type levels of FOXF1 display normal lung morphology and function [11, 19, 21, 27]. Interestingly, transgenic Foxf1 overexpression results in perinatal lethality due to defects in lung development and vasculature [28]. FOXF1 is largely studied during development and is associated with developmental abnormalities in the lung and several other organ systems which were highlighted in several studies through use of heterozygous Foxf1 mouse models [19, 21, 27].

Inactivating FOXF1 mutations are known to cause congenital lung malformations [21,

27]. Heterozygous Foxf1 mice were found to exhibit lethal alveolar hemorrhaging at birth which correlated with lower mRNA levels of Foxf1 [27]. Additional lung abnormalities included disruption in alveolar formation and a disrupted formation of smaller pulmonary vasculature

[27]. Surviving Foxf1+/- mice exhibited near wild-type levels of Foxf1 mRNA [27]. Another study demonstrated that murine Foxf1 haploinsufficiency resulted in lung hypoplasia and right

lung lobe fusion [21]. When harvested at e18.5, perinatal Foxf1+/- lungs were found to be immature, with the right lobes fused, indicating a defect in lung branching morphogenesis. The lungs had a compact and undifferentiated mesenchyme and additionally were deficient of alveoli, which are established during the final step of lung development [29]. This lung hypoplasia phenotype demonstrated a delay in lung maturation for Foxf1+/- mice [21].

FOXF1 haploinsufficiency additionally causes defects in trachea, esophagus, gall bladder, and colon, and is important for ureter development [20, 21, 30, 31]. Foxf1+/- mice exhibit a narrowing of the esophageal lumen, with some mutant esophaguses merging with the trachea. In addition, some Foxf1 heterozygous mice displayed esophageal atresia, which is when the esophagus ends before reaching the pharynx [21]. Haploinsufficiency of Foxf1 also resulted in abnormal gall bladder development including either no gall bladder or a rudimentary gall bladder with structural abnormalities [20]. These abnormalities included malformation of the external smooth muscle layer, a reduced mesenchymal cell number, and in some heterozygous mice, the lack of a biliary epithelial layer [20]. It is noteworthy that the severity of the gall bladder phenotype was linked to mRNA levels of Foxf1; furthermore, mice with WT levels of

Foxf1 displayed a less severe phenotype [20].

A study looking at inactivation of a single Foxf1 allele along with inactivation of a single allele of another FOXF family member, Foxf2 (Foxf1+/-;Foxf2+/-), revealed developmental defects in the colon. Foxf1+/-;Foxf2+/- mice exhibited a dilated, thin-walled colon, known as megacolon [30]. Similar to the same condition in human patients [32], megacolon in Foxf1+/-

;Foxf2+/- mice was caused by innervation of distended parts of the distal colon with a reduction of enteric neurons [30]. FOXF1 has been shown to be expressed in ureteric mesenchyme at e14.5 and has recently been shown to be a crucial factor for ureter development [31].

Previously, SHH had been implicated in ureter development [33, 34]; however, the molecular mechanisms whereby SHH functioned were unknown. FOXF1 is known to be a direct target of HH signaling [35, 36], and recently, SHH was shown to activate FOXF1 to direct growth and differentiation of ureteric tissue, and it is possible that deregulation of the SHH signaling pathway could lead to congenital anomalies of the kidney and urinary tract (CAKUT) in humans [31]. Future studies could investigate FOXF1 as a therapeutic target for human patients with CAKUT.

The studies outlined above give insights to the role of FOXF1 during organogenesis and development. Altogether, wild type levels of Foxf1 are required for normal embryonic development and survival. While Foxf1+/- mice surviving to adulthood display normal organ morphology, murine haploinsufficiency of Foxf1 is associated with a delay in lung [24] and liver

[25] repair. In addition, FOXF1 is known to be important for proper lung barrier function and homeostasis [23]. While FOXF1 does play a major role during development, FOXF1 is also expressed in lung microvascular endothelial cells, hepatic stellate cells (HSCs) of the liver, and fibroblasts of the adult mouse [17, 37]; therefore, FOXF1 mutations are associated with a number of both neonatal and adult diseases.

FOXF1 in Diseases

Alveolar capillary dysplasia with a misalignment of pulmonary veins

Alveolar Capillary Dysplasia with a Misalignment of Pulmonary Veins (ACD/MPV) is a rare, fatal lung disorder affecting both parenchymal and pulmonary vasculature development [38,

39] and is typically diagnosed through autopsy [40]. ACD/MPV is characterized by a decrease in number and size of normally positioned capillaries [41, 42] with malpositioned veins [41, 43].

~80% of ACD/MPV patients have congenital abnormalities associated with ACD/MPV including malformations of the gastrointestinal track, cardiovascular system, and urogenital system [22, 41, 44-48]. While diagnosis of ACD/MPV commonly occurs at autopsy, extra- pulmonary abnormalities can alert physicians to the ACD/MPV condition. While the majority of

ACD/MPV patients are born full-term (95%) [41, 49], symptoms start to develop within 24-48 hours [49-52]. Infants will clinically present with respiratory distress and pulmonary hypertension resulting in cyanosis [22, 44, 49, 50, 52]. ACD/MPV patients typically succumb to hypoxemia respiratory failure within days or weeks of birth despite supportive care [43, 49, 52-

59] and will usually die within one month [48, 50, 52].

The majority of ACD/MPV cases are sporadic with ~10% being familial [43, 60-63].

Genetic studies have shown that FOXF1 is commonly mutated in ACD/MPV patients. Foxf1 heterozygous point mutations and genomic deletion copy number variants at 16q24.1 have been identified in most ACD/MPV patients [22, 59, 64-67]. More than 90% of the reported pathogenic deletions involving the upstream regulatory region of FOXF1 are maternally derived [66-68].

Treatments for ACD/MPV remain elusive. Current therapies include ventilation, nitric oxide, and extracorporeal membrane oxygenation, which are typical treatments for infants with other types of respiratory distress but have proven to be ineffective in treating ACD/MPV patients. As studies emerge which reveal more information regarding the underlying genetic factors leading to ACD/MPV, new avenues for treatment will develop. A recent study from our lab has linked ACD/MPV with a point mutation (p.S52F) in the DNA-binding domain of Foxf1 in which serine 52 is replaced with phenylalanine [69]. This study developed a S52F-Foxf1+/- knock-in mouse model which mimics features of ACD/MPV in humans including fused lung

lobes and hemorrhaging [69]. This novel mouse model will be an important tool for the development of desperately needed de novo treatment options for ACD/MPV patients.

Acute lung injury and acute respiratory distress syndrome

Acute lung injury (ALI) and the more severe acute respiratory distress syndrome (ARDS) are clinical syndromes characterized by an impairment in gas exchange that can result in respiratory failure [70]. Pathogenic mechanisms of lung injury include increase endothelial permeability, inflammatory response, and extracellular matrix remodeling, as well as pulmonary edema [71]. Our lab recently published a murine model of ALI whereby the deletion of FOXF1 deletion induced an inflammatory response and pulmonary edema with a 100% fatality rate [23].

FOXF1 is known to promote lung homeostasis and to aid in repair after injury [23, 24], and restoration of FOXF1 in FOXF1-deficiency-induced murine ALI promoted normal lung homeostasis and repair through S1P/S1PR1 signaling [23].

Management of ALI and ARDS is challenging and currently novel treatment approaches to increase patient survival for ALI and ARDS remain elusive. New therapeutic strategies that aim at the restoration of lung structure and function will be important for ALI patient survival, due to the increased inflammation and disruption of normal lung architecture during disease manifestation. Recent advances in treatment of human lung diseases have introduced cell replacement strategies to contribute to lung regeneration. Perhaps most promising is the use of embryonic stem cells (ESCs) with their ability to target sites of injury [72, 73] and their capability to differentiate into endothelial cells and generate de novo blood vessels [72]. Since

FOXF1 is important for proper lung barrier function, future treatments for ALI and ARDS

should consider development of a FOXF1-expressing endothelial progenitor cell as a therapy to restore lung structure and function.

Pulmonary and Hepatic Fibrosis

Fibrosis is a scaring process associated with excessive deposition of ECM components

[74]. While initially a normal wound-healing response to injury, fibrosis is associated with chronic diseases occurring in many organ systems such as the skin [75], liver [76, 77], heart [78,

79], kidney [80], and lung [79, 81]. The excessive scaring process disrupts organ morphology and ultimately impairs organ function [74]. Since fibroblasts are the main contributors to ECM deposition, they have been the recent target of numerous cellular and molecular studies regarding their role in fibrosis, including two from our lab regarding the role of FOXF1 in pulmonary [82] and hepatic [83] fibrosis. Both the lung and the liver act as filters with the lung filtering the air we breathe to the liver filtering our blood. Insults to each of these organs can result from a variety of infectious, toxic, and metabolic agents. Being a mesenchymal-specific transcription factor, FOXF1 is expressed in lung microvascular endothelial cells, hepatic stellate cells (HSCs) of the liver, and fibroblasts. Murine Foxf1 haploinsufficiency is associated with delayed repair in both the lung [24] and the liver [25]; however, the role of FOXF1 in chronic diseases such as lung and liver fibrosis remains poorly understood.

Idiopathic pulmonary fibrosis (IPF) is a common, lethal interstitial lung disease known to be triggered by recurrent epithelial cell injury. IPF is characterized by the scarring of the pulmonary parenchyma as a result of the accumulation of myofibroblasts (MFs) that secrete collagen and extracellular matrix (ECM) components. Resident lung stromal cells act as

progenitors for the majority of MFs during IPF pathogenesis; however, the transcriptional networks regulating MF function remain unclear. Studies in primary isolated fibroblasts have demonstrated that fibroblasts undergo a change in cadherin expression from CDH2 to CDH11 during MF transdifferentiation [84, 85]. FOXF1 is expressed in resident lung fibroblasts and has recently been shown to prevent cadherin switching of CDH2-CDH11, subsequently thwarting fibrosis progression [82].

FOXF1 is known to be important for lung homeostasis and repair after injury [23, 24]. A recent study demonstrated that the loss of FOXF1 in lung myofibroblasts promotes pulmonary fibrosis [82]. Conditional deletion of FOXF1 during the fibroblast to myofibroblast transition in bleomycin-induced lung fibrosis resulted in an increase in MF migration. This phenotype is associated with MF invasion of damaged basement membrane matrix where MFs accumulate and produce collagens [82]. Chromatin immunoprecipitation (ChIP) sequencing and ChIP assays demonstrated that FOXF1 bound to and transcriptionally regulated both CDH2 and CDH11, which were responsible for MF activation [82]. CDH2 and CDH11 promote fibroblast migration

[86], and both of these cadherins mediate migration and survival in other cell types [87].

Altogether, while FOXF1 expression is decreased in normal pulmonary fibrosis, complete loss of the transcription factor exacerbates the disease phenotype by allowing MF activation through the

CDH2 to CDH11 cadherin switching mechanism [82].

Hepatic fibrosis is the common end stage to a variety of liver diseases and injuries that ultimately results in disruption of hepatic architecture and leads to impairment of organ function.

During fibrogenesis, MFs differentiate from resident HSCs in response to live insult and secrete

ECM and collagen. The transcriptional regulators of MF activation and function remains poorly characterized; however, FOXF1 is known to be expressed in the HSCs of the liver [25] and has

been implicated as a key player in both activation of MFs from HSCs [25, 88, 89] and regulation of ECM deposition during chronic fibrosis [83].

Murine haploinsufficiency of Foxf1 resulted in diminished activation of HSCs and delayed or nonexistent liver repair [25]. Furthermore, siRNA-induced FOXF1 knock-down prevented activation of HSCs and subsequent collagen deposition after cholestatic liver injury

[88], reiterating the importance of FOXF1 in normal liver function and repair following hepatic injury. In a chronic hepatic injury model, conditional deletion of Foxf1 after MF activation resulted in exacerbated hepatic fibrosis with an increase in collagen deposition and liver architecture disruption due to aberrant MF accumulation [83]. Overexpression of FOXF1 in isolated hepatic myofibroblasts inhibited expression of pro-fibrotic genes: Col1α2, Col5α2, and

Mmp2 in vitro [83]. Taken together, this study demonstrated that normal FOXF1 expression in the liver repressed pro-fibrotic gene transcription in HSCs and MFs subsequently preventing MF accumulation and ECM and collagen deposition during hepatic fibrosis [83].

A mechanistic understanding of common fibrosis pathways can lead to the development of treatments that could be effective in multiple organs. FOXF1 has a clear role in the progression of both lung and liver fibrosis. Stabilization of FOXF1 levels may lessen collagen deposition in patients which could be a beneficial therapy for multiple disorders in the lung, liver, and other organs susceptible to fibrosis.

Other diseases (Barret’s esophagus, Fanconia anemia)

Barrett’s esophagus (BE) is a disease common in individuals suffering from long-term gastroesophageal reflux disease, and is characterized by a replacement of normal esophageal

squamous epithelium with tissue normally found in the intestine, metaplastic columnar epithelium [90]. Two separate genome-wide association studies identified single nucleotide polymorphisms near FOXF1 as significantly predisposing individuals to BE [91, 92]. In both of these studies, FOXF1 was highlighted due to its known role in esophageal development [21]; however, further mechanistic work will need to be done to elucidate FOXF1 has a role in the progression of BE.

A more promising study has identified a transcriptional role of FOXF1 in Fanconi anemia

(FA) [93], a rare pediatric disorder characterized by various congenital abnormalities, bone marrow failure, and increased susceptibility to cancers due to heterogeneous defects in the DNA damage repair pathway [94-96]. Through co-immunoprecipitation experiments, FOXF1 was found to interact with FA protein complexes [93]. The FA protein complex activates to repair

DNA damage. Foxf1-knock down studies decreased the stability of FA binding complexes, indicating that FOXF1 stabilizes the FA chromatin binding complex [93]. This study additionally demonstrated that FOXF1 was able to aid in DNA repair of HeLa tumor cells through association with FA complex proteins [93]. Based on these data, stabilization of FOXF1 could be a promising therapeutic strategy to treat FA as well as cancer.

FOXF1 and Cancer

Cancer occurs when a single cell changes and begins to grow unregulated [97]. These cell changes may result from an external agent such as a carcinogen, or from inherited genetic abnormalities. Tumor growth can preclude organs from functioning properly or can cause damage to adjacent normal tissue [97]. In 2018, cancer will be responsible for 9.6 million deaths worldwide [98]. This is up from 8.8 million people in 2015 and 8.2 million in 2012 [98]. The

five most deadly types of cancer worldwide, in order, are: lung, colorectal, gastric, liver, and breast [98]. Of these, lung and liver cancers have the lowest 5-year survival rates of 18.2% and

17.2%, respectively [99]. Approximately 70% of global cancer-related deaths occur in low or middle-income countries [98] although the number of cancer-related deaths in the United States has dropped in recent years with an increased number of survivors [100].

Cancer cells can invade normal tissues, which is known as metastasis and is the transition of a disease from one part of the body to another. Metastasis is a major cause of death from cancer [98, 101] and is responsible for approximately 90% of cancer-related mortalities

[102, 103]. In addition to lymph nodes and bone, the lung and liver are the most common places for metastatic cancers to settle [101, 104]. Cancer-related death can be prevented with early diagnosis and treatments. Treatments vary between cancer types and severity at the time of diagnosis but typically include chemotherapy, surgery, and targeted therapy [97]. While great advances have been made in cancer prevention and treatment, more work needs to be done to lower the devastating mortality numbers.

Deregulation of FOX transcription factors are known to lead to disease development and progression, including cancer [14]. The importance of FOXF1 in cancer has been supported by recent findings in multiple cancer models. FOXF1 has been shown to play a critical role in tumorigenesis, and expression levels of FOXF1 are known to determine the development and progression of cancer [105-107]. Depending on the specific type of cancer, FOXF1 has been shown to be either a tumor suppressor [107, 108] or an oncogene [106, 109] and additionally, high expression of FOXF1 has been associated with metastasis in multiple cancers [110, 111].

Altogether, current knowledge highlights the complexity of the role of FOXF1 in carcinogenesis.

Lung

FOXF1 is known to be expressed in both fetal and adult lung tissues [15, 17]. FOXF1 regulates fibroblast stimulation of lung cancer cell migration [105]. In lung cancer, FOXF1 also stimulates xenograft tumor growth and cancer cell migration through regulation of cancer- associated fibroblasts (CAFs) [105]. Previously, malignant lung cancer cells were shown to fuse with mesenchymal stem cells (MSCs) and the fusion cells had a less severe phenotype than the original malignant cells [109]. It was additionally determined through transcriptome profiling that FOXF1 contributed to the fusion reprogramming and contributed to tumor growth suppression [109].

While few studies have been done to look at the role of FOXF1 in the progression of lung cancer, recent data from our lab has shown that low levels of FOXF1 expression correlate with poor patient survival [112]. Additionally, FOXF1-overexpression in an orthotopic model of lung cancer in mice revealed an inhibition of lung tumor growth and metastasis [112]. Furthermore,

RNA-sequencing analysis revealed that Wnt/β-catenin signaling is regulated by FOXF1 in lung endothelial cells [112], confirming previous reports that FOXF1 controls mesenchymal Wnt expression and that loss of FOXF1 could lead to tumor susceptibility [30].

Taken together, these studies suggest that FOXF1 plays a tumor suppressive role in lung cancer. Future studies should look to elucidate the mechanisms whereby FOXF1 regulates CAFs, reprograms MSC-fusion tumor cells, and mediates Wnt/β-catenin signaling. These data will give insights to the development of novel therapeutics for FOXF1 stabilization to treat lung cancer.

Gastrointestinal (Colorectal, gastric, liver)

Colorectal.

In colorectal cancer (CRC), FOXF1 has been shown to act as a tumor suppressor that is epigenetically silenced during disease progression, leading to genomic instability [113].

Inactivating nuclear FOXF1 functions through mis-localization and overexpression of FOXF1 in the cytoplasm has been linked to a poor CRC prognosis [113]. Since FOXF1 is typically expressed in the cell nucleus, it is possible that in these separate compartments of the cell,

FOXF1 can act as either a tumor suppressor or an oncogene. Under-expression or loss of FOXF1 in tumor-associated stromal fibroblasts has been observed in CRC patients, suggesting that

FOXF1 expression in stromal fibroblasts is important in CRC progression [113]. Recently, genetic polymorphisms in FOXF1 have been linked to an increased risk of developing CRCs

[114]. Taken together, these data suggest that normally-functioning, nuclear FOXF1 acts as a tumor suppressor in CRCs.

Gastric.

In gastric cancer, FOXF1 has been shown to act as a tumor suppressor. A recent study identified methyl-CpG binding protein 2 (MeCP2), a known transcriptional regulator, as an oncogene in gastric cancer. MeCP2 was shown to inhibit Wnt5a/β-catenin signaling through inhibition of FOXF1. FOXF1 inhibition resulted in increased proliferation of gastric cancer cells

[115]. The Wnt5a/β-catenin signaling pathway is key in regulating cancer progression in liver

[116, 117], lung [118, 119], and colorectal cancers [120], and FOXF1 functions in this pathway though its interactions with bone morphogenetic protein 4 [30, 35].

Liver.

In the liver, FOXF1 is typically expressed in the hepatic stellate cells [25]. A recent paper demonstrated that a decrease in FOXF1 expression was associated with poor clinical outcomes for hepatocellular carcinoma (HCC) patients [121]; However, FOXF1 was shown to be expressed in the hepatocytes of the tumor cells. More work will need to be done to evaluate this alteration of FOXF1 expression from normal liver tissue to diseased tissue. Previously, a 2013 study performed a microarray analysis on RNA from HCC patients and submitted the dataset to the Gene Expression Omnibis (GEO) public database at the National Center for Biotechnological

Information (NCBI, Bethesda, MD; accession number GSE41804) [122]. Analysis of the dataset revealed that FOXF1 is significantly decreased (p<0.01) in HCC patients, leading to the possibility that FOXF1 acts as a tumor suppressor in HCC [122]. As previously described, in pre-cancerous hepatic fibrosis, low levels of FOXF1 expression lead to an increase in collagen depositions [83], which could indicate a protective role of FOXF1 during the progression of liver diseases and cancer.

Breast

FOXF1 may function as either a tumor suppressor or an oncogene in breast cancers. In breast cancer, some labs have identified FOXF1 as a tumor suppressor that is silenced during disease progression [107, 123]. Hypermethylation of the FOXF1 promoter epigenetically silences FOXF1 in breast cancer cell lines. Re-expression of FOXF1 was shown to lead to G1 cell cycle arrest, which halted tumor formation and growth [107]. Similar to colorectal cancer cell lines, FOXF1 was lost or under-expressed in breast cancer cell lines with inactive p53 which promoted genomic instability and tumorigenesis, consistent with a tumor suppressive role of

FOXF1 [123].

In contrast, other labs have demonstrated that FOXF1 acts as an oncogene during breast carcinogenesis [106, 124]. FOXF1 overexpression was shown to stimulate breast cancer xenograft tumor growth in vivo and cancer cell migration in vitro [106]. Overexpression of

FOXF1 in HC11 (normal mouse mammary epithelial cell line) and induced expression of

FOXF1 in HB2 cells (non-malignant human mammary epithelial cell line which does not endogenously express FOXF1) have been shown to upregulate the extracellular matrix protein, lysyl oxidase (LOX) [124]. LOX increases extracellular matrix stiffness and allows tumor progression and invasion [125] and is known to be upregulated in breast cancer cell lines and carcinomas [126, 127]. In these cell lines, FOXF1 was additionally shown to suppress SMAD2/3 signaling [124], which is striking since an imbalance between canonical and non-canonical TGF-

β signaling is thought to be responsible for the oncogenic activities of TGF-β in breast cancer

[128].

As a tumor suppressor, FOXF1 maintains genomic stability through its role in cell cycle progression. As an oncogene, overexpression of FOXF1 influences breast cancer progression invasion and metastasis through an imbalance of canonical and non-canonical TGF-β signaling.

Since FOXF1 seemingly has different roles in breast carcinogenesis depending on the specific tumor type. Future therapeutics could either stabilize or inhibit FOXF1 expression on an individual basis.

Other cancers (Prostate, rhabdomyosarcoma)

In the prostate, FOXF1 is expressed in the stroma and is thought to maintain normal prostate homeostasis through the stromal compartment in a similar way as during gut development and during embryogenesis [30]. A 2004 genomic analysis named FOXF1 (along

with one predicted and four known genes) as a candidate tumor suppressor gene in prostate cancer due to its downregulation in prostate tumors, as a result from the loss of the 16q24 chromosomal region [129]. Furthermore, FOXF1 has lower expression levels in tumor cells than in the normal prostate stromal compartment [130]. However, more work needs to be done to confirm the status of FOXF1 as a tumor suppressor in prostate cancer and to determine mechanistically how FOXF1 functions during prostate carcinogenesis.

FOXF1 has been shown to be highly expressed in alveolar rhabdomyosarcoma (RMS)

[131, 132], an aggressive pediatric soft tissue sarcoma [133]. Gene expression microarray data revealed that FOXF1 was more highly expressed in metastatic RMS tumors than non-metastatic

RMS samples [132]. This indicated an oncogenic role for FOXF1 in RMS. FOXF1 was additionally found to synergize with its most closely-related family member, FOXF2, to induce

RMS tumorigenesis in orthotropic tumor transplantation models [134], corroborating an oncogenic role for FOXF1 in RMS.

Summary

FOXF1 has many known roles throughout development and disease pathogenesis. Loss of this important transcription factor is embryonic lethal and mutations in the gene can result in fatal diseases. Depending on the tissue and type of cancer, FOXF1 has been shown to act as either a tumor suppressor or an oncogene. Despite our vast understanding of FOXF1, more work needs to be done to understand the transcriptional networks and signaling cascades influenced by

FOXF1 to identify novel therapies to increase patient survival.

Experimental Rationale

The studies detailed in the current dissertation were conducted to further our understanding of the FOXF1 transcription factor. Previous work has shown that FOXF1 is expressed in the collagen-producing cells of the liver, the hepatic stellate cells (HSC), and that

HSC activation requires FOXF1 presence. Chapter 2 utilizes a carbon tetrachloride-induced liver fibrosis model to investigate the role FOXF1 in accumulation of collagen during liver fibrosis. A more commonly known role for FOXF1 is expression in endothelial cells, the cells that line blood vessels. FOXF1 is an important regulator of vascular formation and has roles in cellular proliferation. Chapter 3 describes the utilization of endothelial colony forming cells for acute lung injury rescue and the development of embryonic stem cell (ESC)-derived, FOXF1- expressing endothelial progenitor cells.

In Chapter 2, we utilized a carbon-tetrachloride (CCl4) -induced model of liver injury in combination with a conditional FOXF1-knockout to understand the role of FOXF1 during hepatic fibrosis. While there are numerous types of hepatic injury that can lead to fibrosis, hepatoxicity induced by CCl4 mimics drug-induced liver damage which is a physiologically relevant model of this disease and will provide researchers with imperative details regarding hepatic fibrosis pathogenesis and progression. A better understanding of the cellular and molecular mechanisms underlying the progression of hepatic fibrosis will provide insights for the development of potential therapeutic treatments for patients. It is expected that the identified role of FOXF1 in hepatic fibrosis will lead to the development of novel therapies targeting FOXF1 for the treatments of liver diseases leading to fibrosis, fibrosis itself, and possibly hepatocellular carcinoma.

In Chapter 3A, we utilized endothelial colony forming cells, a type of endothelial progenitor cell (EPC), to rescue acute lung injury (ALI). Recent advances in the treatment of human lung diseases have described cell replacement strategies as novel therapies for lung tissue repair and regeneration. Since treatment options to increase ALI patient survival remain elusive, a better understanding of the molecular mechanisms of cell transplantation in murine models could provide insights for the development of novel treatment approaches for ALI and other severe lung disorders. In Chapter 3B, we utilized CRISPR/Cas9 to develop a novel GFP:FOXF1 embryonic stem cell (ESC) line. Since FOXF1 is expressed in lung endothelial cells and is known to be important for lung homeostasis and repair after acute lung injury, we developed a novel differentiation protocol that yielded FOXF1-expressing EPCs. It is expected that future studies will utilize these GFP:FOXF1-positive EPCs to mechanistically determine how cell replacement therapies work in murine models, which will lead to improvement of patient cell replacement therapies.

Although FOXF1 has been studied through embryonic development and in a myriad of diseases, there are many unanswered questions regarding different aspects of this transcription factor. Here in the current dissertation, we address the role of FOXF1 during the progression of hepatic fibrosis. We additionally develop a novel GFP:FOXF1 ESC line as a tool for future studies in lung regenerative medicine. Altogether, the work described here enhances our knowledge base on FOXF1 throughout different systems and can be used to develop future studies as described in the discussion in Chapter 4.

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Chapter 2: The Role of FOXF1 in Hepatic Fibrosis

Literature Review: The Liver and Hepatic Fibrosis

The liver is the largest organ of the human body and plays a vital role in whole body metabolism homeostasis [1]. It is composed of lobules made up of a central vein and sheets of hepatocytes, which are surrounded by portal triads. These portal triads contain portal veins, arteries, and bile ducts [2]. The liver is responsible for the production of bile required for food digestion [3], factors for blood clotting [4], amino acids for protein synthesis [5], and iron for transport [6, 7]. The liver is additionally responsible for storage of glycogen which can be converted into glucose when the body needs energy [8]. This organ aids in detoxifying chemical- laden blood from the digestive tract before reentering general circulation, and can eliminate various endogenous and exogenous molecules, such as metabolism waste in the form of urea, which is secreted from the body in urine [9].

The liver can be damaged in a number of ways, which can lead to a variety of chronic liver diseases such as hepatitis, cholestasis, steatosis, and alcoholic liver disease [10]. Hepatitis leads to cell inflammation and can cause immune-mediated liver damage. Hepatitis can be caused by viruses [11], ranging from the hepatitis virus to the varicella virus [12], or from drugs or toxins which lead to liver inflammation [13, 14]. Cholestasis is a disease which occurs when the normal flow of bile in the liver is obstructed, resulting in bile retention. This bile accumulation can damage hepatocytes and bile duct cells [15]. Steatosis, commonly referred to as fatty liver disease, is known to occur with an accumulation of cholesterol and triglycerides in the liver. The liver typically contains a small storage of fat; however, an excessive amount can lead to liver damage and inflammation [16, 17]. Finally, the most common form of liver damage in Western countries is alcoholic liver disease. Since the liver metabolizes alcohol, excessive

alcohol consumption over time can lead to hepatocyte necrosis, inflammatory cell infiltration, and portal hypertension [18].

Each of these liver diseases can independently lead to fibrosis, cirrhosis, and ultimately, hepatocellular carcinoma (HCC) [10, 19]. HCC has limited treatment options is the second leading cause of cancer mortality worldwide, with nearly 750,000 deaths reported in 2012. Since fibrosis is the common end stage of these and various additional hepatic diseases, it is important to understand the mechanisms involved in fibrosis, its regulation, and the reversal of its progression. Doing so could reveal new therapeutic targets to treat patients with these disorders, as well as to prevent the progression to non-reversible cirrhosis or HCC.

The liver is composed of many different cell types which contribute to normal liver homeostasis and proper function [20]. The three main liver cell types are hepatocytes, Kupffer cells (KCs), and hepatic stellate cells (HSCs). Hepatocytes are the main epithelial cells of the liver and make up 80% of the liver mass and approximately 70% of total cell numbers.

Hepatocytes are responsible for production of the factors necessary for proper liver functions; they are responsible for carbohydrate and lipid metabolism, as well as detoxification of substances taken in by the body to maintain homeostasis [21]. KCs represent an important component of the liver immune system as the resident macrophages [20]. They reside in the liver sinusoid lumen, located between the sheets of hepatocytes, which enables exposure to bacteria, endotoxins, and microbial debris directly from the gastrointestinal tract and portal vein [22].

HSCs reside in the space of Disse, an area within liver sinusoids between the sheets of hepatocytes and a layer of endothelial cells [23]. During the quiescent state, HSCs are characterized by their storage of lipids and are the body's main storage units of Vitamin A. After liver insult, HSCs can activate and differentiate into myofibroblasts (MFs) [1]. The forkhead box

F1 (Foxf1) transcription factor is expressed in both HSCs and MFs during embryonic development and in the adult liver [24]. While HSCs make up only a small number of cells in the total liver, they are the main contributors of MFs during liver repair [25, 26].

During normal liver injury, hepatocytes are damaged and immune cells infiltrate the microenvironment. Hepatocytes and KCs secrete cytokines such as TGF-β, TNF-α, and IL-63, which stimulate HSCs to undergo the transition from a quiescent to an activated state [27]. HSCs will lose quiescent features such as lipid storage, and will adopt new features such as contractility and chemotaxis, and will differentiate into MFs. MFs are the pathogenic cell type in fibrosis that express α-smooth muscle actin and produce extracellular matrix (ECM) and collagens [28]. Initially after liver insult, MFs secret ECM and collagens to encapsulate the site of injury and protect the liver (Fig. 2.0 A). Published studies have demonstrated that FOXF1 is essential for hepatic stellate cell activation and liver repair after acute liver injury (Fig. 2.0 B).

Foxf1 siRNA delivered to mice through nanoparticles prevented activation of HSCs and subsequent collagen deposition after cholestatic liver injury [29]. Murine haploinsufficiency of

Foxf1 (Foxf1+/-) was associated with diminished activation of HSCs and delayed liver repair after carbon tetrachloride-induced hepatic injury, which resulted in 100% mouse death by day 3 [24]

(Fig. 2.0 C). While we know that FOXF1 is required for HSC activation, the role of FOXF1 in differentiation of HSCs into MFs during the progression of hepatic fibrosis remains unknown.

HSCs and MFs, like KCs, synthesize TGF-β in the liver, which allows for TGF-β to play a key role in liver fibrosis initiation and maintenance in a positive feedback loop, promoting fibrosis progression [30]. TGF-β reduces cellular apoptosis and is a hepatocyte proliferation factor [23], suggesting TGF-β is required for cellular and organ homeostasis, as well as for cellular stabilization during liver injury and fibrogenesis. Initially the HSC injury response is

Figure 2.0. FOXF1 activity during liver injury. Schematic illustrations demonstrates (A) the normal wound healing response to hepatic injury. (B) FOXF1 is expressed in both quiescent hepatic stellate cells (HSCs) and activated myofibroblasts (MFs) during liver injury. (C) FOXF1 expression is necessary for proper activation of MFs to protect the liver from damage.

protective and facilitates repair (Fig. 2.0 A), however, prolonged activation of HSC and differentiation of MFs is pathogenic, driving hepatic fibrosis [31].

When scar tissue production is unregulated, fibrosis can occur [32]. Fibrosis is an overly exuberant wound healing process in which collagen and ECM components are overproduced, degraded deficiently, or a combination of both devices [33]. Fibrosis is the end stage of many types of chronic liver injury such as hepatitis, cholestasis, and steatosis [10]. It is additionally a critical factor important for the pathogenesis of HCC development with the majority of patients with HCC having fibrosis [19]. Hepatic fibrosis, before progression to cirrhosis or HCC, is known to be a reversible process [34]. Several factors are known to mediate the resolution of fibrosis and restoration of liver homeostasis. These include removal or treatment of the underlying insult, a shift in immune factor balance to facilitate repair, deactivation of MFs, and degradation of ECM and collagens [35]. This resolution allows for hepatocyte recovery, induction of restorative KCs, removal of the matrix-producing cells, and an alteration of the balance between matrix metalloproteinases (MMPs) and their inhibitors, tissue inhibitors of metalloproteinases (TIMPs), which work in a system which promotes remodeling of the ECM24.

MMPs and TIMPs help regulate liver repair homeostasis, and are secreted by HSCs and MFs.

MMPs and TIMPs work in a system which promotes remodeling of the ECM [36-38]. After liver insult, MMP expression decreases while TIMP expression increases to allow for ECM accumulation to encapsulate the site of injury and protect the liver [39].

While we know that FOXF1 is required for HSC activation, the role of FOXF1 in differentiation of HSCs into MFs and its role during the progression of hepatic fibrosis remains unknown. There are few published studies investigating FOXF1 in the liver, therefore, the current chapter reveals novel insights into the role of FOXF1 in the progression of fibrosis.

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The Forkhead Box F1 Transcription Factor Inhibits Collagen Deposition

and Accumulation of Myofibroblasts During Liver Fibrosis

Hannah M. Flood1, Craig Bolte1, Nupur Dasgupta2, Akanksha Sharma3, Yufang Zhang1, Chandrashekhar R. Gandhi1,3, Tanya V. Kalin1, Vladimir V. Kalinichenko1

1Department of Pediatrics, Cincinnati Children’s Research Foundation, Cincinnati, Ohio, USA. 2Division of Human Genetics, Cincinnati Children’s Research Foundation, Cincinnati, Ohio, USA. 3Division of Gastroenterology, Hepatology, and Nutrition, Cincinnati Children’s Research Foundation, Cincinnati, Ohio, USA.

The work presented in Chapter 2 has been published in the following peer-reviewed journal:

Flood HM, Bolte C, Dasgupta N, Sharma A, Zhang Y, Gandhi CR, Kalin TV, Kalinichenko VV. Forkhead Box F1 Transcription Factor Inhibits Collagen Deposition and Accumulation of Myofibroblasts During Liver Fibrosis. Biology Open. 2019;8(2).

Abstract

Hepatic fibrosis is the common end stage to a variety of chronic liver injuries and is characterized by an excessive deposition of extracellular matrix (ECM), which disrupts the liver architecture and impairs liver function. The fibrous lesions are produced by myofibroblasts, which differentiate from hepatic stellate cells (HSC). The myofibroblasts transcriptional networks remain poorly characterized. Previous studies have shown that the Forkhead box F1

(FOXF1) transcription factor is expressed in HSCs and stimulates their activation during acute liver injury; however, the role of FOXF1 in the progression of hepatic fibrosis is unknown. In the present study, we generated αSMACreER;Foxf1fl/fl mice to conditionally inactivate Foxf1 in myofibroblasts during carbon tetrachloride-mediated liver fibrosis. Foxf1 deletion increased collagen depositions and disrupted liver architecture. Timp2 expression was significantly increased in Foxf1-deficient mice while MMP9 activity was reduced. RNA sequencing of purified liver myofibroblasts demonstrated that FOXF1 inhibits expression of pro-fibrotic genes,

Col1α2, Col5α2, and Mmp2 in fibrotic livers and binds to active repressors located in promotors and introns of these genes. Overexpression of FOXF1 inhibits Col1a2, Col5a2, and MMP2 in primary murine HSCs in vitro. Altogether, FOXF1 prevents aberrant ECM depositions during hepatic fibrosis by repressing pro-fibrotic gene transcription in myofibroblasts and HSCs.

Introduction

The liver is the body’s filter and insults can result from a variety of infectious, toxic, and metabolic agents. Hepatic fibrosis is the common end stage to a multitude of liver diseases [1] and is characterized by an excessive deposition of extracellular matrix (ECM) and collagen [2].

Novel animal models of hepatic fibrosis are greatly needed to identify molecular mechanisms responsible for the disease pathogenesis and development of therapeutic agents. Hepatic stellate cells (HSC) reside in the space of Disse and are characterized by their storage of lipids when in the quiescent state [3, 4]. During fibrogenesis, quiescent HSCs differentiate into myofibroblasts

(MF) in response to cytokine signaling from damaged hepatocytes and immune cells after liver insult. MFs secrete ECM and collagen to encapsulate the site of injury and shield the liver from plaguing insults [2]. While HSCs and MFs make up only a small number of cells in liver tissue, they are the main contributors of ECM and collagen during liver repair and fibrogenesis [5, 6].

The TGF-β and PDGF signaling pathways play key roles in hepatic fibrosis and HSC activation

[7]. TGF-β signaling stimulates cellular transdifferentiation of HSCs to MFs [8, 9], whereas

PDGF signaling induces cellular proliferation in fibrotic foci [10, 11].

The Forkhead Box F1 (FOXF1) transcription factor is expressed in human and murine

HSCs and is important in regulating stellate cell activation after acute liver injury [12]. In the advanced disease state of hepatocellular carcinoma (HCC), which is associated with significant fibrotic depositions, FOXF1 expression has been shown to be significantly decreased [13].

Foxf1-/- mice are embryonic lethal due to severe developmental abnormalities in the yolk sac and allantois [14]. Murine haploinsufficiency of Foxf1 causes lung hypoplasia, loss of alveolar capillaries in the lung, and gall bladder agenesis [15, 16], and was associated with delayed lung

+/- and liver repair. After acute liver injury by carbon tetrachloride (CCl4), Foxf1 mice exhibited

diminished activation of HSCs and delayed liver repair, indicating that FOXF1 is essential for liver repair after acute liver injury [12]. Foxf1 siRNA delivered to mice through nanoparticles prevented activation of HSCs and subsequent collagen deposition after cholestatic liver injury

[17]. While these studies have shown that FOXF1 is required for activation of HSCs after acute liver injury, the role of FOXF1 in MFs and in the progression of fibrotic responses remains unknown.

In the present study, we generated a novel genetic mouse model to conditionally delete

Foxf1 from MFs (αSMACreER;Foxf1fl/fl). During chronic liver injury, deletion of Foxf1 in MFs exacerbated hepatic fibrosis, increased collagen deposition, and stimulated expression of profibrotic genes in the liver tissue. Our studies indicate that Foxf1 expression in MFs is critical to prevent MF accumulation and collagen deposition during liver fibrosis.

Results

Deletion of Foxf1 in αSMA-positive Cells Exacerbates CCl4-Induced Hepatic Fibrosis. Previous studies demonstrated that FOXF1 is present in HSCs in murine developing and adult livers [12,

18]. Consistent with these studies, FOXF1 staining was detected in livers of e12.5-e17.5 mouse embryos as well as in mesenchyme of stomach and intestine (Suppl. Fig. 2.1). In adult mice,

FOXF1 is specifically expressed in the liver parenchyma but not in endothelial or smooth muscle cells surrounding the portal vein or hepatic artery [12] (Fig. 2.1 A, Suppl. Fig. 2.1), and FOXF1 staining co-localized with desmin (DES) (Fig. 2.1 A), a known marker of HSCs [19]. To investigate the role of Foxf1 in liver fibrosis, we utilized a conditional knockout approach.

Transgenic mice containing a tamoxifen-inducible αSMA-CreER transgene and two Foxf1-floxed alleles (αSMACreER;Foxf1fl/fl) were generated by breeding αSMA-CreER and Foxf1fl/fl mice (Fig.

2.1 B-C). Hepatic fibrosis was induced by chronic liver injury using multiple administrations of

CCl4, which is known to increase fibrotic depositions and disrupt liver architecture in experimental mice [20]. Tamoxifen was given 3 times per week to achieve a continuous deletion of Foxf1 in αSMA-positive MFs (Fig. 2.1 D) that derive from HSCs after liver injury [21].

Morphological analysis of liver sections revealed increased fibrotic deposition in CCl4-treated

αSMACreER;Foxf1-/- livers compared to controls as shown by H&E (Fig. 2.1 E, Suppl. Fig. 2.2) and Masson’s Trichrome staining (Fig. 2.1 F-G, Suppl. Fig. 2.2). Increased fibrosis in

αSMACreER;Foxf1-/- livers was confirmed by qRT-PCR for Col1α1 and Col3α1 mRNAs (Fig.

2.1 H) as well as significant increases in collagen levels by Sircol (Fig. 2.1 I) and hydroxyproline assays (Fig. 2.1 J). Treatment with tamoxifen alone (without CCl4) did not affect liver architecture or induce liver fibrosis (Suppl. Fig. 2.2). Thus, deletion of Foxf1 from MFs accelerates liver fibrosis after chronic liver injury.

Figure 2.1. Hepatic fibrosis is increased after CCl4 injury in mice with FOXF1 deficiency. (A) FOXF1 co-localizes with DES in hepatic stellate cells in adult mice. (B) Diagram demonstrates αSMA-CreER transgene with LoxP sites flanking the Foxf1 Exon 1 (encoding DNA-binding fl/fl fl/fl domain). (C) DNA gel shows genotypes of Foxf1 and αSMACreER;Foxf1 mice. (D)

Diagram illustrates CCl4 and tamoxifen (Tam) treatment protocol. (E-F) H&E and Masson’s trichrome staining show fibrotic depositions after 5-weeks of CCl4 treatment. Fibrosis was -/- increased in livers from αSMACreER;Foxf1 mice. White lines indicate fibrotic lesion boundaries. (G) ImageJ analysis of Masson’s trichrome images shows a significant increase in -/- collagen in livers from αSMACreER;Foxf1 mice. n=3 mice per group in week 0; n=5 mice per group in week 5. (H) qRT-PCR analysis demonstrates significant increases in Col1α1 and -/- Col3α1 mRNAs in livers from αSMACreER;Foxf1 mice. n=3 mice per group in week 0; n=5 fl/fl fl/fl mice per group in week 5. Untreated livers from Foxf1 and αSMACreER;Foxf1 mice were used as normal controls. mRNAs were normalized to Actb. (I-J) Collagen deposition was quantitated using Sircol and hyroxyproline assays. n=2 mice per group in week 0; n=4 mice per group in week 5. P<0.05 is indicated with *, P<0.01 is indicated with **.

FOXF1 Expression is Decreased in Hepatic Myofibroblasts of αSMACreER;Foxf1-/- Mice. Since

FOXF1 is expressed in HSCs in the liver [12], we examined the efficiency of Foxf1 deletion in our experimental model, using immunostaining for FOXF1 and DES. Without CCl4 treatment,

FOXF1 was observed in cell nuclei of DES-positive stellate cells in Foxf1fl/fl and

-/- αSMACreER;Foxf1 livers (Fig. 2.2 A). After CCl4 and Tam treatment, FOXF1 staining was reduced in DES-positive cells of αSMACreER;Foxf1-/- livers but not in Foxf1fl/fl livers (Fig. 2.2

A). We also immunostained liver sections for FOXF1 and αSMA, a marker of MFs [22]. While

αSMA was not detected in parenchyma of quiescent livers, αSMA staining was increased after

-/- CCl4 injury. FOXF1 was detected in MFs of control livers but not in αSMACreER;Foxf1 livers

(Fig. 2.2 B). Quantitative counts of FOXF1-expressing cells demonstrated that the number and percentage of FOXF1+ MFs (FOXF1+ αSMA+) were reduced whereas the number and percentage of FOXF1- MFs (FOXF1- αSMA+) were elevated in injured αSMACreER;Foxf1-/- livers compared to controls (Suppl. Fig. 2.3). FOXF1 protein and mRNA were increased in CCl4- treated Foxf1fl/fl livers and purified HSCs (Fig. 2.2 C-D, Suppl. Fig. 2.3) but not in the

αSMACreER;Foxf1-/- livers (Fig. 2.2 C-D). The loss of FOXF1 in αSMACreER;Foxf1-/- livers occurred in periportal regions while pericentral regions were unaffected (Fig. 2.2 E). Thus,

αSMA-CreER transgene effectively deletes Foxf1 from hepatic MFs during CCl4-mediated chronic liver injury.

Deletion of Foxf1 Reduces MMP9 Activity in CCl4-Injured Livers. Histological staining with

Sirius Red/Fast Green showed an increase in collagen accumulation in αSMACreER;Foxf1-/- livers after 5-weeks of CCl4 treatment (Fig. 2.3 A, Suppl. Fig. 2.4). Increased fibrosis in Foxf1- deficient livers was confirmed by immunostaining for DES and αSMA (Fig. 2.3 B-C). To

Figure 2.2. αSMA-CreER effectively deletes Foxf1 from hepatic myofibroblasts. (A-B) FOXF1 co-localizes with DES in HSCs before and after CCl4-induced injury. FOXF1 co-localizes with DES and αSMA in MFs after chronic liver injury. αSMA-CreER effectively deletes Foxf1 from MFs after Tam treatment. (C) Western blot shows total liver protein levels of FOXF1 are -/- decreased in αSMACreER;Foxf1 livers after CCl4 injury. Cropped blots are presented here with full length blots presented in Suppl. Fig. 12. (D) Quantification of Western blot revealed a -/- significant loss of FOXF1 in αSMACreER;Foxf1 livers. Quantification was averaged across two additional blots with n=3 mice per group in week 0 and n=5 mice per group in week 5. FOXF1 levels were internally normalized to ACTIN for each sample. P<0.05 is indicated with *. (E) FOXF1 staining is detected in liver parenchyma and fibrotic regions (yellow arrows). FOXF1 -/- staining is decreased in fibrotic regions of αSMACreER;Foxf1 livers (white arrows).

Figure 2.3. Deletion of Foxf1 from myofibroblasts increases liver fibrosis and inhibits MMP9 activity. (A) Sirius Red/Fast Green staining demonstrates increased collagen deposition between -/- portal triads inCCl4-treated αSMACreER;Foxf1 livers. (B-C) Immunohistochemistry shows -/- increased staining for DES and αSMA in CCl4-treated αSMACreER;Foxf1 livers. (D-E) Serum enzymatic analysis demonstrates increased AST and ALT levels after chronic CCl4-induced liver injury. Foxf1 deletion does not affect AST or ALT in blood serum. For AST levels: n=3 control mice and n=4 KO mice in week 0; n=5 control mice and n=5 KO mice in week 5; n=4 control mice and n=7 KO mice in week 18. For ALT levels: n=5 control mice and n=6 KO mice in week 0; n=7 control mice and n=8 KO mice in week 5; n=4 control mice and n=7 KO mice in week -/- 18. (F) Increased Timp2 mRNA in CCl4-treated αSMACreER;Foxf1 livers is found by qRT- PCR. (G) Representative zymography gel shows decreased MMP9 activity in CCl4-treated αSMACreER;Foxf1-/- livers. Cropped gel is presented here with full gel presented in Suppl. Fig. 12. (H) Quantification of zymography gels reveals a significant decrease in MMP9 activity in -/- CCl4-treated αSMACreER;Foxf1 livers. Quantification was averaged across 3 gels. P<0.05 is indicated with *, P<0.01 is indicated with **.

examine consequences of extended CCl4 treatment, we treated mice with CCl4 for 18-weeks.

While hepatic enzymes AST and ALT were increased in blood serum after 18-weeks of CCl4,

-/- there was no difference between CCl4 treated αSMACreER;Foxf1 and control mice (Fig. 2.3 D-

E). Blood serum protein (albumin, globulin) and bilirubin (direct, indirect) levels were not affected after deletion of Foxf1 (Suppl. Fig. 2.5). Collagen accumulation was time-dependent

(Suppl. Fig. 2.6), and after 18-weeks of CCl4 treatment, resulted in widespread liver fibrosis

(Suppl. Fig. 2.7) and in rare cases, appearance of visible tumors (Suppl. Fig. 2.7).

Since MMP9 plays an important role in collagen degradation after liver injury [23], we evaluated mRNA expression of Mmp9 and its inhibitor, Timp2, in liver tissue. Timp2 mRNA was

-/- increased in CCl4-injured αSMACreER;Foxf1 livers compared to controls (Fig. 2.3 F).

Although Mmp9 mRNA was unchanged (Suppl. Fig. 2.8), evaluation of MMP9 activity through zymography demonstrated a significant decrease in enzymatic activity of MMP9 in

-/- αSMACreER;Foxf1 livers after CCl4 treatment (Fig. 2.3 G-H). Mmp8, Mmp13, Mmp16, Timp1, and Timp3 mRNA levels were not affected in Foxf1-deficient livers (Suppl. Fig. 2.8). Thus,

Foxf1 deletion from MFs increases Timp2 mRNA and reduces MMP9 activity in fibrotic livers.

Deletion of Foxf1 Does Not Influence Cellular Proliferation in Fibrotic Livers. We evaluated proliferation markers to investigate if the increased fibrosis in αSMACreER;Foxf1-/- livers was due to an expansion of the stromal cells. While cellular proliferation was increased after CCl4 treatment, there were no significant differences in the number of proliferating hepatocytes or non-hepatocytes between Foxf1fl/fl and αSMACreER;Foxf1-/- livers (Fig. 2.4 A-C, Suppl. Fig.

2.9). Hepatocytes and non-hepatocytes were identified through distinct morphological appearances [24] from high magnification images. mRNAs of proliferation-specific genes

Figure 2.4. Deletion of Foxf1 does not influence proliferation of hepatic myofibroblasts. (A) Ki- 67 staining shows a significant increase in cell proliferation following CCl4-induced liver injury. No difference in Ki-67 staining is detected between Foxf1fl/fl and αSMACreER;Foxf1-/- livers. (B) PH3 staining shows no significant changes in mitotic rates between Foxf1fl/fl and αSMACreER;Foxf1-/- livers. (C) The number of Ki-67+ hepatocytes and non-hepatocytes in Foxf1fl/fl livers was similar to those in αSMACreER;Foxf1-/- livers. Numbers of Ki-67+ cells were counted in 20-25 random 200x microscope fields using n=3 mice per group in week 0 and n=7 control mice and n=6 KO mice in week 5. (D) qRT-PCR was used to measure mRNAs in whole liver RNA. mRNAs were normalized to Actb. n=3 mice per group in week 0; n=5 mice per group in week 5. (E) Co-localization of Ki-67 with DES shows the presence of Ki-67+ MFs in livers of CCl4-treated mice. (F) Quantification of co-localization of Ki-67 with DES shows no difference in the number of Ki-67+ DES+ cells in Foxf1fl/fl livers compared to αSMACreER;Foxf1-/- livers. (G) Western blot shows no significant difference in total liver protein levels of FOXM1 and CCND1 between Foxf1fl/fl and αSMACreER;Foxf1-/- livers. Cropped blots are presented here with full length blots presented in Suppl. Fig. 12. P<0.05 is indicated with *, P<0.01 is indicated with **.

Foxm1, Ccnb1, Ccnd1, and AurKB [25-27] were unchanged between Foxf1fl/fl and

-/- αSMACreER;Foxf1 livers (Fig. 2.4 D). Proliferating HSCs and MFs were detected in CCl4- treated livers by co-localization of Ki-67 with DES (Fig. 2.4 E) and αSMA (Suppl. Fig. 2.9); however, there were no changes in the number of Ki-67-positive HSCs and MFs after deletion of

Foxf1 (Fig. 2.4 F). Protein levels of proliferation-specific genes FOXM1 and CCND1 were unaltered in αSMACreER;Foxf1-/- livers compared to controls (Fig. 2.4 G). Thus, Foxf1 deletion does not affect proliferation of HSCs and MFs after chronic CCl4 liver injury.

RNA-seq Analysis Identified Direct FOXF1 Target Genes Critical for ECM Deposition and

Hepatic Fibrosis. In order to identify FOXF1 target genes, RNA-seq was performed on primary

fl/fl hepatic stromal cells (containing MFs and HSCs) isolated from CCl4-treated Foxf1 and

αSMACreER;Foxf1-/- livers. Purified cells expressed Des and Acta2, but lacked hepatocyte [28] and Kupffer Cell [29] markers (Suppl. Fig. 2.10). Foxf1 mRNA was lost in isolated

αSMACreER;Foxf1-/- stromal cells (Fig. 2.5 A), a finding consistent with efficient deletion of

Foxf1 from CCl4-treated livers. The RNA-seq was used to compare differential gene expression patterns between the Foxf1fl/fl and αSMACreER;Foxf1-/- stromal cells. The differential gene expression in the two groups are represented in a heat map (Fig. 2.5 B). Gene ontology demonstrated that increased functional pathways for the αSMACreER;Foxf1-/- mice were related to ECM regulation, while decreased functional pathways included normal liver functions and metabolism (Fig. 2.5 C). RNA-seq analysis was cross referenced with FOXF1 ChIP-seq analysis

(GEO Accession GSE100149). 905 genes were common between RNA-seq and ChIP-seq (Fig.

2.5 D), which include 74 genes related to ECM deposition and fibrosis. ChIP-seq proximity analysis revealed that 20 of these ECM genes had FOXF1 binding sites within 2KB of the transcription start site (Fig. 2.5 E).

Figure 2.5. FOXF1 deletion alters expression of pro-fibrotic genes in hepatic myofibroblasts. (A) qRT-PCR analysis of primary hepatic stromal cells submitted for RNA sequencing shows that Foxf1 mRNA is not detected (n.d.) in αSMACreER;Foxf1-/- livers. Samples were pooled for further analysis. (B) Heat map shows differentially expressed genes in stromal cells from fl/fl -/- Foxf1 and αSMACreER;Foxf1 livers after chronic CCl4-induced hepatic injury as identified by RNA-seq analysis. (C) Biological processes influenced by the deletion of Foxf1 were identified using ToppFunn. P-values and number of genes are listed for each classification. (D) 905 overlapping genes were identified between ChIP-seq (GEO accession GSE100149) and RNA-seq data, of which 74 were ECM-related genes. (E) Table shows 20 ECM-related genes identified by ChIP-seq and RNA-seq (FOXF1 binding was analyzed within 2 KB from transcriptional start site).

Interestingly, Col1α2, Col5α2, and Mmp2 were among the 20 ECM-related genes that had FOXF1 binding sites within the gene loci (Suppl. Fig. 2.11, Suppl. Table 2.1). COL1α2 and

COL5α2 are common ECM components in fibrotic livers [30], whereas MMP2 is a collagenase that is increased during liver fibrosis and associated with disease progression [31]. Expression of

-/- Col1α2, Col5α2, and Mmp2 mRNAs were increased in CCl4-treated αSMACreER;Foxf1 livers as shown by RNA-seq and qRT-PCR (Fig. 2.5 E, Fig. 2.6 D), suggesting a negative regulation by

FOXF1. The presence of gene silencing histone methylation marks H3K9me3 and H3K27me3

[32, 33] in FOXF1-binding promoter regions (Fig. 2.6 A-C) is consistent with negative regulation of these genes by FOXF1. In order to confirm the regulation of Col1α2, Col5α2, and

Mmp2 by FOXF1, we overexpressed FOXF1 in isolated murine HSCs (Fig. 2.6 E). Lentiviral- mediated overexpression of FOXF1 decreased Col1α2 and Mmp2 in vitro (Fig. 2.6 F). Thus,

FOXF1 negatively regulates expression of pro-fibrotic genes in MFs. Altogether, FOXF1 expression in myofibroblasts is essential to inhibit liver fibrosis after chronic liver injury (Fig.

2.6 G).

Figure 2.6. FOXF1 binds to DNA regulatory regions of Col1α2, Col5α2, and Mmp2. (A-C) ChIP-seq shows FOXF1 binding near the transcriptional start sites in Col1α2, Col5α2, and Mmp2 gene locuses. Histone modification marks of enhancers (H3K4me3, H3K9ac) and repressors (H3K9me3, H3K27me3) are aligned with FOXF1-binding regions. Significant areas of FOXF1 binding are marked with boxes, with blue boxes indicating the binding site is within gene promoter region. Gene transcriptional start sites are marked with a directional yellow arrow. (D) qRT-PCR analysis shows the significant increase of Col1α2, Col5α2, and Mmp2 -/- mRNAs in the isolated stromal cells of CCl4-treated αSMACreER;Foxf1 livers. For Col1α2 and Col5α2: n=3 mice per group in week 0; n=5 mice per group in week 5. For Mmp2: n=3 mice per group in week 0; n=6 control mice and n=4 KO mice in week 5. (E) Western blot shows increase in FOXF1 expression in isolated HSCs after FOXF1-overexpression. Cropped blots are presented here with full length blots presented in Suppl. Fig. 12. (F) qRT-PCR shows an increase of Foxf1 mRNA and a decrease of Col1α2, Col5α2, and Mmp2 mRNAs in isolated HSCs after FOXF1-overexpression. (G) Diagram of hepatic fibrosis in Foxf1-deficient mice shows that the loss of FOXF1 promotes ECM deposition and exacerbated fibrosis after CCl4-treatment. P<0.05 is indicated with *, P<0.01 is indicated with **, P<0.001 is indicated with ***.

Discussion

Myofibroblast activation is a key mechanism in the development of hepatic fibrosis.

However, transcriptional regulation of myofibroblasts during liver fibrogenesis remains poorly characterized. In the present study, we found that the deletion of Foxf1 in MFs during chronic

CCl4-mediated injury exacerbated liver fibrosis, increased collagen deposition, and stimulated expression of pro-fibrotic genes. ECM-related proteins were identified as novel FOXF1 transcriptional targets, suggesting that FOXF1 plays an important role in the regulation of ECM and collagen deposition during the progression of hepatic fibrosis.

Previous studies have focused on the role of FOXF1 in acute liver injury using a single

+/- CCl4 administration to Foxf1 mice. These studies demonstrated that FOXF1 is necessary for

+/- HSC activation to promote liver repair [12]. CCl4-treated Foxf1 mice exhibited diminished collagen depositions and increased mortality after the liver injury [12]. A recently published model of Foxf1-silencing using a lipid-based nanoparticle system to deliver Foxf1 siRNA to the liver demonstrated attenuated collagen deposition when Foxf1 siRNA was delivered 48 h prior to bile duct ligation [17]. It is likely that Foxf1 siRNA inhibited FOXF1 signaling in hepatic stellate cells, decreasing their activation and subsequent collagen depositions into the liver tissue, a finding consistent with previous studies using Foxf1+/- mice [12]. Recently, a model of chronic hepatic injury using CCl4-injections, similar to the present study, was unsuccessful in silencing

Foxf1 expression using the same lipid based system to deliver Foxf1 siRNA [17, 34]. This method involved four weeks of IP CCl4-injections before two weeks of treatment with Foxf1 siRNA [34]. It is possible that the lack of Foxf1 silencing was due to inability of nanoparticles to target hepatic MFs. In the current study, we utilized a conditional genetic mouse model to delete

Foxf1 in MFs during CCl4-induced hepatic fibrosis which shares multiple histological

similarities with human disease [35, 36]. Interestingly, the loss of Foxf1 in MFs resulted in increased collagen deposition, causing severe fibrotic lesions between hepatic portal triads in

αSMACreER;Foxf1-/- livers. Our studies suggest that FOXF1 inhibits production of collagen and

ECM during the progression of liver fibrosis. Increased fibrosis in Foxf1-deficient mice was associated with the appearance of liver tumors, a finding consistent with increased tumor formation in patients with liver cirrhosis [37]. Our studies suggest that maintaining Foxf1 expression can be beneficial in patients with advanced liver fibrosis to inhibit fibrotic responses and decrease the risk of liver tumorigenesis.

In the present study, collagens were significantly increased in αSMACreER;Foxf1-/- livers

-/- after chronic CCl4-treatment. Desmin and αSMA were both increased in αSMACreER;Foxf1 livers; however, there were no differences in the number of proliferating cells between Foxf1fl/fl and αSMACreER;Foxf1-/- livers. Previously, FOXF1 has been shown to stimulate cell proliferation in lung endothelial cells [38, 39] and in rhabdomyosarcoma tumor cells [40].

Surprisingly, we found that deletion of Foxf1 from MFs does not affect their proliferation during liver fibrogenesis. It is possible that FOXF1 requires additional co-activator or co-repressor proteins (that are not present in MFs) to regulate cellular proliferation. Additionally, we found an increase in Timp2 expression with a decrease in MMP9 activity in αSMACreER;Foxf1-/- livers.

Since it is well-known that TIMPs and MMPs regulate ECM depositions to balance the scaring and healing processes during fibrosis [23], it is possible that the loss of Foxf1 alters the

TIMP/MMP balance to allow accumulation of collagens without the degradation mechanisms necessary for proper wound healing. Interestingly, MMP9 has been implicated in HSC to MF transdifferentiation [41] in addition to its roles in ECM degradation [23, 42]. Therefore, decreased MMP9 activity can contribute to increased liver fibrosis in αSMACreER;Foxf1-/- mice.

Surprisingly, FOXF1 was increased in activated HSCs compared to quiescent HSCs. It is possible that FOXF1 is differentially regulated in HSCs compared to hepatic MFs, and that after liver injury, FOXF1 protects HSCs from differentiating into MFs through transcriptional repression of profibrotic genes.

Consistent with increased fibrosis in Foxf1-deficient livers, RNA-seq analysis revealed increased ECM-related functional pathways in a purified stromal cell population. Comparison with FOXF1 ChIP-seq data revealed 20 novel transcriptional targets of FOXF1, which include

Col1α2, Col5α2, and Mmp2, expression of which was increased in Foxf1-deficient cells.

COL1α2 is one of the most abundant ECM components in the liver along with COL1α1 and COL3α1 [43]. COL5α2 is highly expressed with Collagens 1 and 3 and is important in regulating the assembly and structure of these collagens in the fibrotic matrix [44]. MMP2 acts as a collagenase, known to be activated during hepatic fibrosis [31]. In addition to increased mRNA levels of the genes in FOXF1-deficient cells, we found multiple FOXF1 binding sites within their gene promoter region and introns, suggesting direct transcriptional repression. This hypothesis is supported by the presence of H3K4me3 and H3K9ac, histone modifications associated with transcriptional repression [32, 45], at FOXF1 binding sites. In summary, we have developed a novel genetic mouse model to study the role of FOXF1 in MFs during chronic liver injury. Using this model, we demonstrated that Foxf1 expression in MFs is necessary to inhibit hepatic fibrosis and maintain the balance of collagen depositions, through transcriptional repression of pro-fibrotic genes.

Methods

Mice.

The Foxf1fl/fl mouse line was previously generated and bred into the C57Bl/6 mouse background

[38, 46]. Foxf1fl/fl mice were bred with αSMA-CreER mice (Jackson Laboratory, 029925, [47] to generate αSMACreER;Foxf1fl/fl mice [48]. αSMACreER;Foxf1fl/fl mice were bred with Foxf1fl/fl mice and male pups were genotyped and used for all experiments at the age of 6-8 weeks. The following primers were used for genotyping: αSMA-CreER sense: 5’

TGCAACGAGTGATGAGGTTCGC 3’ and anti-sense: 5’

GATCCTGGCAATTTCGGCTATACG 3’; αSMA-WT sense 5’

GGTTTCTATTGCTACCAAGAGACAT 3’ and anti-sense: 5’

TGCACCAAACCCTGGACTAAGCAT 3’; Foxf1fl/fl sense: 5’ GCTTTGTCTCCAAGCGCTGC

3’ and anti-sense: 5’ TTCAGATCTGAGAGTGGCAGCTTC 3’. Foxf1fl/fl littermates were used as controls. To activate the conditional Foxf1 knockout, tamoxifen (Tam) was given via intraperitoneal injection (40 mg/kg of body weight; Sigma) three days in a row at the beginning of each week starting at week 2 over course of chronic liver injury period. Hepatic injury was induced by intraperitoneal injections of carbon tetrachloride (CCl4; 1 μl/g of body weight 20% v/v; Sigma; diluted in sunflower seed oil) three times a week every other day over the course of the chronic liver injury period. The levels of aminotransferases AST and ALT, proteins albumin and globulin, and direct and indirect bilirubin were determined by serological analysis of blood serum as previously described [49, 50]. All animal studies were approved by the Institutional

Animal Care and Use Committee (IACUC) of Cincinnati Children’s Research Foundation and the NIH IACUC Guidebook. All experiments were covered under our animal protocol

(IACUC2016-0038). The Cincinnati Children’s Research Foundation Institutional Animal Care and Use Committee is an AAALAC and NIH accredited institution (NIH Insurance #8310801).

Histology and immunohistochemistry.

Paraffin-embedded liver sections were used for H&E, immunohistochemistry (IHC), or immunofluorescence (IF) as previously described [12, 50, 51]. The following antibodies were used for immunostaining: FOXF1 [1:1000 IHC, 1:200 IF, R&D Systems], DES [1:500 IHC,

1:100 IF; (Santa Cruz Technologies)], αSMA [1:10,000 IHC, 1:5000 IF; (Sigma)], Ki-67 [1:1000

IHC, 1:200 IF; (Thermo Scientific)], Ki-67 [1:200 IF; (BD Biosciences)], and PH3 [1:10,000

IHC; (Santa Cruz Technologies)]. Antibody-antigen complexes were detected using biotinylated secondary antibodies followed by avidin-biotin-horseradish peroxidase complex and

3,3’diaminobenzidine substrate (Vector Labs) as previously described [12, 50, 52]. Sections were counterstained with nuclear fast red (Vector Labs). For immunofluorescence imaging, secondary antibodies conjugated with Alexa Fluor 488 or Alexa Fluor 594 (Invitrogen/Molecular

Probes) were used as described [53, 54]. Cell nuclei were counterstained with DAPI (Vector

Labs). Masson’s Trichrome (BD Biosciences) and Sirius Red/Fast Green (Chondrex, Inc.) specialty stains were performed according to manufacturer protocols. Brightfield images were obtained using a Zeiss AxioImage.A2 microscope. Fluorescent images were obtained using a

Zeiss AxioPlan 2 microscope. qRT-PCT, Western blot, and zymography.

The caudate lobe of the liver was halved and used for RNA and protein studies. RNA was isolated using RNA Stat-60 (Tel-Test, Inc.) according to manufacturer protocol and was reverse transcribed using the High Capacity Reverse Transcription Kit (Applied Biosystems) according to manufacturer protocol. mRNAs of specific genes were measured by qRT-PCR using TaqMan probes (Applied Biosystems; Supplementary Table 2.2) and the StepOnePlus Real-Time PCR system (Applied Biosystems) as described [54-57]. mRNAs were normalized to Actb. Protein

extracts were isolated using cell lysis buffer as previously described [58] and used for either

Western blot analysis with Pierce ECL Western blotting substrate (Thermo Scientific) or gel zymography (NOVEX) according to manufacturer protocols. The following antibodies were used for protein blots: FOXF1 [1:1000, R&D Systems] [38, 39, 48], ACTIN [1:2000; (Santa

Cruz)] [58], FOXM1 [1:3000; (Santa Cruz)] [58] [55], CCND1 [1:1000; (Cell Signaling)] [40].

Protein band intensities were determined by ImageJ software and were normalized to ACTIN.

Hepatic Stellate Cell Isolation, Transfection.

Hepatic Stellate Cells were isolated from male C57Bl/6-WT mice (40-50 g), purified using

Nycodenz gradient, and cultured as previously described [58-60]. Quiescent HSCs were harvested at day two after cell culture [61]. After ten days in culture, activated MFs were harvested [61]. The pMIEG3 bicistronic retroviral vector was used for FOXF1 protein overexpression as previously described [58]. The cells were transfected as previously described

[62]. mRNAs in isolated MFs were normalized to 18s [Eukaryotic 18S rRNA Endogenous

Control; (Applied Biosciences)]. Protein in isolated MFs were normalized to LAMIN AC

[1:10,000; (Santa Cruz)] [63].

RNA sequencing.

fl/fl RNA was isolated from HSC/MF population purified from CCl4-treated Foxf1 and

αSMACreER;Foxf1-/- livers using a differential plating method [64] which we modified. Briefly, liver cell suspension was plated on tissue culture dishes (Corning) and incubated for 2 hours at

37°C. Supernatant and non-adherent cells were washed off and the adherent cell population was collected for experiments. Samples were pooled to generate the libraries using the TruSeq RNA library preparation kit and were sequenced on an Illumina HiSeq 2000 sequencer, generating about 10 M high quality single end reads (75 base-long reads). Allignment was perfomed using

the Tophat/Cufflink pipeline [65, 66]. Finally, cuffmerge tool was used to generate Binary

Alignment/Map files (BAM files) [67]. BAM files of RNA-seq data were analyzed using

Avadis® NGS Version 1.3.0 software. Reads were filtered to remove: 1) duplicate reads, 2) non- primary-matched reads, and 3) reads with alignment scores <95. Quantification was performed on the filtered reads against the RefSeq annotation. Data normalization was performed with the

DESeq package. The sequencing depth was estimated by the read count of the gene with the median read count ratio across all genes. The method was based on the negative binomial distribution, which allows for less restrictive variance parameter assumptions than does the

Poisson distribution. The false discovery rate was calculated according to the Benjamini and

Hochberg algorithm [68]. Genes with expression altered by a factor of 1.5 and a false discovery rate of 0.05 in Foxf1fl/fl cells compared with αSMACreER;Foxf1-/- cells were selected for gene set enrichment analysis using ToppGene Suite. Hierarchical clustering was performed by Ward’s method using Euclidean distance metric. RNA-seq data were compared to previously published

ChIP-seq data (GEO accession GSE100149) using a two-way Venn diagram.

Statistical analysis.

Student’s t-test was used to determine statistical significance. P<0.05 was considered to be significant, with P<0.05 indicated with *, P<0.01 indicated with **, and P<0.001 indicated with

***. Values for all measurements were expressed as mean ± standard error of mean.

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Supplementary Information For Chapter 2

Supplemental Figure 2.1. FOXF1 expression in mouse livers. Supplemental Figure 2.2. Treatment with tamoxifen alone does not induce hepatic fibrosis. Supplemental Figure 2.3. Number and percentage of FOXF1+ myofibroblasts are reduced in -/- αSMACreER;Foxf1 livers. Supplemental Figure 2.4. Increased collagen deposition in Foxf1-deficient livers. Supplemental Figure 2.5. Deletion of Foxf1 had no effect on serum protein or bilirubin levels. Supplemental Figure 2.6. Collagen accumulation in Foxf1-deficient livers is time-dependent. Supplemental Figure 2.7. Widespread hepatic fibrosis and appearance of liver tumors in -/- αSMACreER;Foxf1 mice after 18-weeks of CCl4 treatment. Supplemental Figure 2.8. Deletion of Foxf1 does not affect Mmp8, Mmp13, Mmp16, Timp1, or

Timp3 mRNAs in CCl4-treated livers. + Supplemental Figure 2.9. Deletion of Foxf1 does not affect proliferation of αSMA cells in CCl4- treated livers. Supplemental Figure 2.10. Purified stromal cells express Acta2 and Des. Supplemental Figure 2.11. ChIP-seq shows FOXF1 binding sites in DNA regulatory regions of Col1α2, Col5α2, and Mmp2. Supplemental Figure 2.12. Full images for Western blot and zymography.

Supplemental Table 2.1. List of TaqMan probes used in qRT-PCR analysis. Supplemental Table 2.2. FOXF1 binding sites as identified by ChIP-seq relative to the transcriptional start site.

Supplemental Figure 2.1. FOXF1 expression in mouse livers. (A) Immunostaining shows FOXF1 protein (dark brown) in nuclei of mesenchymal cells surrounding stomach (st) and intestine (int) of e12.5-17.5 mouse embryos. FOXF1 is also detected in mesothelial and stellate cells of the liver (li). Slides were counterstained with nuclear fast red (red). (B) Immunostaining shows FOXF1 expression in HSCs of uninjured livers. Lung sections were used as positive control for FOXF1 staining. (C) Immunostaining shows that FOXF1 is expressed in liver parenchyme but not in endothelial cells lining the portal vein (PV) and hepatic artery (HA). Bile ducts are shown as BD. (D) Co- localization of FOXF1 with αSMA shows that FOXF1 is not expressed in smooth muscle cells fl/fl surrounding the portal vein (V) in uninjured Foxf1 livers.

Supplemental Figure 2.2. Treatment with tamoxifen alone does not induce hepatic fibrosis. (A) fl/fl fl/fl H&E and (B) Masson’s trichrome staining of Foxf1 and αSMACreER;Foxf1 livers after 5- weeks of CCl4 and Tam treatment show widespread hepatic fibrosis. (C) H&E and Masson’s fl/fl fl/fl trichrome staining of Foxf1 and αSMACreER;Foxf1 livers from mice treated with Tam alone fl/fl or in combination with CCl4. Tam treatment does not cause hepatic fibrosis in either Foxf1 or fl/fl αSMACreER;Foxf1 mice. Tam-mediated deletion of Foxf1 exacerbates the fibrotic phenotype -/- observed in CCl4-treated αSMACreER;Foxf1 livers.

Supplemental Figure 2.3. Number and percentage of FOXF1+ myofibroblasts are reduced in αSMACreER;Foxf1-/- livers. (A) Immunostaining shows FOXF1 expression in HSCs of uninjured livers. Lung sections were used as positive control for FOXF1 staining. (B) Co- localization of FOXF1 with αSMA in Foxf1fl/fl and αSMACreER;Foxf1-/- livers. Scale bars 200 μm. (C) Counts of cells double stained for FOXF1 and αSMA show an increased number and -/- percentage of αSMA+ FOXF1- MFs in CCl4-treated αSMACreER;Foxf1 livers compared to controls. (D) qRT-PCR shows a significant increase in Foxf1 mRNA in HSCs isolated from C57Bl/6-WT mice at the activated stage compared to uninjured livers.

Supplemental Figure 2.4. Increased collagen deposition in Foxf1-deficient livers. Sirius Red/Fast -/- Green staining shows widespread collagen accumulation in αSMACreER;Foxf1 livers fl/fl compared to Foxf1 livers.

Supplemental Figure 2.5. Deletion of Foxf1 had no effect on serum protein or bilirubin levels. The loss of Foxf1 did not change serum albumin levels between control and Foxf1-deficient mice at 0, 5, or 18 weeks of CCl4-treatment. Globulin levels were significantly increased at week 18 between Foxf1fl/fl and αSMACreER;Foxf1-/- livers, as were total protein levels. n=5 mice per group in weeks 0 and 5; n=3 control mice and n=5 KO mice in week 18. Deletion of Foxf1 had no effect on bilirubin levels in blood serum. Data are shown as mean ± SEM.

Supplemental Figure 2.6. Collagen accumulation in Foxf1-deficient livers is time-dependent.

H&E, Masson’s trichrome, and Sirius Red/Fast green staining show a timecourse of CCl4- -/- induced hepatic fibrosis. Collagen depositions were greater in CCl4-treated αSMACreER;Foxf1 livers.

Supplemental Figure 2.7. Widespread hepatic fibrosis and appearance of liver tumor in -/- αSMACreER;Foxf1 mouse after 18-weeks of CCl4 treatment. Representative liver sections stained for (A) H&E, (B) Masson’s Trichrome, and (C) Sirius Red/Fast Green demonstrate that -/- severe fibrotic lesions occur in αSMACreER;Foxf1 livers after 18-weeks of chronic hepatic -/- injury. (D) H&E images show a hepatic tumor (Tu) in an αSMACreER;Foxf1 liver (Li) after 18- weeks of CCl4-treatment. Tumors were found in 14.29% of mice (1 mouse out of n=7). Tumor boundaries are shown with arrows.

Supplemental Figure 2.8. Deletion of Foxf1 does not affect Mmp8, Mmp9, Mmp13, Mmp16,

Timp1, or Timp3 mRNAs in CCl4-treated livers. qRT-PCR was used to measure mRNAs in whole liver extract. Mmp16 was not detected (n.d.) in any sample tested. mRNAs were normalized to Actb. For Mmp8, Mmp13, Mmp16, Timp1, and Timp3: n=3 mice per group in week 0; n=5 mice per group in week 5. P<0.05 is indicated with *, P<0.01 is indicated with **.

+ Supplemental Figure 2.9. Deletion of Foxf1 does not affect proliferation of αSMA cells in CCl4- fl/fl treated livers. (A) Immunostaining for PH3 and Ki-67 shows no difference between Foxf1 and -/- αSMACreER;Foxf1 livers. (B) Co-localization of Ki-67 with αSMA shows the presence of Ki- + 67 MFs in livers of CCl4-treateed mice. Co-localization of Ki-67 with FOXF1 shows that + proliferating FOXF1 cells were uncommon in CCl4-treated livers.

Supplemental Figure 2.10. Purified stromal cells express Acta2 and Des. qRT-PCR analysis shows the presence of Acta2 and Des mRNAs. Hepatocyte marker Alb and Kupffer cell marker -/- Clec4f were not detected in purified stromal cells (one αSMACreER;Foxf1 sample expressed Clec4f ). mRNAs were normalized to Actb.

Supplemental Figure 2.11. ChIP-seq shows FOXF1 binding sites in DNA regulatory regions of Col1α2, Col5α2, and Mmp2. Schematics of FOXF1 binding across entire genes: Col1α2, Col5α2, and Mmp2. ChIP-seq for FOXF1 shows significant binding in multiple DNA regions (indicated with *) of Col1α2 (a peak binding height of 2.04425), Col5α2 ( a peak binding height of 6.81417), and Mmp2 (a peak binding height of 6.98452).

Supplemental Figure 2.12. Full images for Western blot and zymography. (A) Full Western blots for FOXF1 (40 kDa) and corresponding ACTIN (42 kDa). Samples derive from the same experiment and blots were processed in parallel. Cropped blots are shown in Fig. 2C. (B) Full zymography gel used for analysis of MMP9 activity (92 kDa). Cropped gel shown in Fig. 3G. (C) Full Western blots for CCND1 (33 kDa), FOXM1 (84 kDa), and corresponding ACTIN (42 kDa). Samples derive from the same experiment and blots were processed in parallel. Cropped blots shown in Fig. 4G. (D) Full Western blots for FOXF1 and corresponding LAMIN AC (70 kDa). The FOXF1 bands are higher than the predicted 40 kDa due to the tags on the vector. Cropped blots shown in Fig. 6E.

Supplemental Table 2.1. List of TaqMan probes used in qRT-PCR analysis.

Supplemental Table 2.2. FOXF1 binding sites as identified by ChIP-seq relative to the transcriptional start site.

Chapter 3: Mouse Embryonic Stem Cells and Endothelial Progenitor Cells

3A: Transplantation of Endothelial Colony Forming Cells Improves Survival of FOXF1-deficient Mice

3B: Differentiation of Novel GFP:FOXF1 Embryonic Stem Cell Line into FOXF1-positive Endothelial Progenitor Cells

Literature Review: Endothelial Cells, Endothelial Progenitor Cells, and Lung Regenerative Medicine

Endothelial cells (ECs) are found throughout all tissues of the body. ECs form a one-cell thick layer called the endothelium and line the walls of three main systems: the arterial, venous, and lymphatic systems. The arterial system supplies tissues with oxygen and nutrients throughout the body. The venous system removes deoxygenated blood and waste from the tissues, and the lymphatic system is involved in the regulation of interstitial fluid balance. The endothelium maintains vascular structure and regulates vascular homeostasis. Additionally, as the interface between blood and tissue, ECs interact with cells circulating in blood on one side, and with cells of the vascular wall on the other side. ECs regulate vascular permeability by acting as a barrier for molecule transport between the blood or lymph and tissues. FOXF1 is known to be critical for lung angiogenesis during embryonic development [1-3] and is important for lung endothelial barrier function in the adult mouse [4].

An early differentiation study demonstrated that endothelial cells matured through distinct steps by following EC differentiation from embryoid bodies [5], which contain derivatives from all three germ layers and are to be considered pluripotent [6]. This study was the first step in determining how researchers can differentiate their own ECs from stem cells.

Advances in the treatment of human lung diseases have introduced cell replacement strategies to contribute to lung tissue repair and regeneration. Since ECs play a critical role in the regulation of vascular permeability, angiogenesis, and tissue regeneration, development of clinically- relevant numbers of ECs ex vivo could lead to novel treatment approaches for the restoration of endothelial function in patients with vascular diseases.

Recent studies have focused on the generation of endothelial progenitor cells (EPCs), to varying degrees of success. EPCs, in all forms, are a promising target for regenerative medicine research. EPCs are able to regenerate ECs in vivo [7, 8], are able to home to injury sites [9, 10], form de novo blood vessels [9], and engraft into the host lung in human patients [11]. EPC transplantation therapy has recently emerged as beneficial treatment for many chronic lung diseases. Introduction of EPCs were shown to improve patient health in a phase I pulmonary arterial hypertension clinical trial [12]. Neonatal patients with respiratory distress syndrome were more likely to survive if they had higher numbers of EPC concentrations circulating in their blood, suggesting that EPCs may be involved in the regeneration of neonatal lung after injury

[13]. Together, these preliminary studies highlight the potential importance of EPC-based therapies in clinical treatment of pulmonary diseases.

Since many labs and researchers have been working to understand EPCs, three main populations of adult EPCs have emerged from the literature [14]. These putative adult EPC populations are the colony form unit-Hill (CFU-Hill) cell, the circulating angiogenic cell (CAC), and the endothelial colony forming cell (ECFC) [14, 15]. CFU-Hill cells were isolated by plating peripheral blood mononuclear cells and isolating “discrete” colonies [16, 17]. CACs were isolated in a similar manner as CFU-Hill cells but were cultured with endothelial growth medium

[10, 18]. While both CFU-Hill and CAC cells display endothelial characteristics, CAC cells do not form colonies as do CFU-Hill cells. To isolate ECFCs, the peripheral blood mononuclear fraction was plated and cultured for weeks until colonies form with a distinctive cobblestone morphology [19]. These cells are phenotypically akin to ECs and form tubes in vitro and de novo vessels in vivo [14]. While these three populations were initially isolated directly from the

peripheral blood, researchers have been utilizing in vitro techniques to differentiate embryonic stem cells (ESCs) into EPCs with unique or similar properties to these three main types of EPC.

EPCs are currently being derived from multiple types of ESCs in many studies. Mouse

ESCs (mESCs) were the first to be identified, isolated, and cultured [20]. These mESCs were derived from the mouse blastocyst stage of embryonic development and cultures could clonally derive from single cells [20]. When injected back into mice, these cells would form teratocarcinomas [20], which contain derivatives from all three germ layers [21]. ESCs are considered to be pluripotent and can give rise to any mature cell type (Fig. 3.0 A). mESCs are not the only type of stem cell utilized in research today. Controversy infiltrated the stem cell field when in 1998, Thompson [22] and Gearhert [23] separately isolated human ESCs (hESCs) which were derived from either donated in vitro fertilized human embryos grown to the blastocyst stage [22] or from primordial germ cells from aborted fetuses 5-9 week post- fertilization [23]. The repercussions from these experiments are still felt today with these experiments considered controversial. While hESC are still in use today from these early studies, researchers in the stem cell field have developed a new types of human stem cell which can be derived from adult somatic cells.

Since stem cells derived from humans are more clinically relevant than those derived from mice, researches needed to re-evaluate century-old dogma that once cell fate was decided, it was permanent. In 2006, Yamanaka successfully reprogrammed patient skin fibroblasts into pluripotent stem cells by altering the expression of four genes: OCT3/4, SOX2, C-MYC, and

KLF4 [24]. These are now known as “Yamanaka Factors.” These stem cells are known as induced pluripotent stem cells (iPSCs) and can be derived from adult somatic cells [24]. Similar

Figure 3.0. Differences and Similarities between embryonic stem cells and induced pluripotent stem cells. (A) Diagram represents the totipotent zygote maturing into the pluripotent blastocyst. Cells are isolated from the blastocyst and can be cultured as pluripotent embryonic stem (ESCs). (B) Diagram represents the isolation of a somatic cell from a patient which is then exposed to reprogramming factors. The reprogrammed cell is known as an induced pluripotent stem cell (iPSC) and can be cultured ex vivo.

to ESCs, iPSCs are pluripotent, capable of self-renewal, and can differentiate into any cell of the body (Fig. 3.0 B).

The first proof-of-principle therapeutic application for reprogramming iPSCs to cure diseases occurred in mice with sickle cell anemia [25]. Researchers utilized skin cells from the diseased mouse and reverted the somatic cells back to a stem-like state as in previous reports [24,

25]. The mouse-derived iPSCs were differentiated into bone marrow stem cells and were transplanted back into the mouse [25]. Final analysis revealed disease-free mice with blood and kidney measurements indistinguishable from wild-type mice [25].

iPSCs are becoming an attractive resource to generate ECs which can be used in regenerative medicine to engineer de novo blood vessels revascularize ischemic tissues. Many studies have successfully differentiated EPCs from iPSCs in vitro. As mentioned previously, development of clinically-relevant numbers of EPCs ex vivo could lead to novel treatment approaches for the restoration of endothelial function in patients with vascular diseases.

Peripheral blood studies have shown that improved survival in patients with acute lung injury is associated with an increase in numbers of circulating EPCs [26]. A recent pulmonary hypertension phase I trial demonstrated that the introduction of EPCs into the pulmonary artery resulted in an improvement of the 6-minute walk capacity of patients, reflecting a physiological response involving the pulmonary, cardiovascular, circulatory, and neuromuscular systems [12].

Excitingly, a study of female patients who had received stem cell transplants from male donors revealed 37.5-42.3% endothelial chimerism in lung biopsies [11]. Altogether, these studies have highlighted the therapeutic potential of iPSC-derived EPCs for treatment of lung diseases.

Besides utilization of iPSCs and ESCs for tissue repair and regeneration, in vitro differentiation to EPCs and ECs will provide opportunities to elucidate the molecular

mechanisms which govern endothelial fate. These mechanistic studies will aid in our understanding of vascular disease pathogenesis and will give us insights into the development of novel therapeutic interventions for disease treatments.

References

1. Ren X, Ustiyan V, Pradhan A, Cai Y, Havrilak JA, Bolte CS, et al. FOXF1 transcription factor is required for formation of embryonic vasculature by regulating VEGF signaling in endothelial cells. Circ Res. 2014;115(8):709-20. 2. Kalinichenko VV, Lim L, Stolz DB, Shin B, Rausa FM, Clark J, et al. Defects in pulmonary vasculature and perinatal lung hemorrhage in mice heterozygous null for the Forkhead Box f1 transcription factor. Dev Biol. 2001;235(2):489-506. 3. Mahlapuu M, Ormestad M, Enerback S, Carlsson P. The forkhead transcription factor Foxf1 is required for differentiation of extra-embryonic and lateral plate mesoderm. Development. 2001;128(2):155-66. 4. Cai Y, Bolte C, Le T, Goda C, Xu Y, Kalin TV, et al. FOXF1 maintains endothelial barrier function and prevents edema after lung injury. Sci Signal. 2016;9(424):ra40. 5. Vittet D, Prandini MH, Berthier R, Schweitzer A, Martin-Sisteron H, Uzan G, et al. Embryonic stem cells differentiate in vitro to endothelial cells through successive maturation steps. Blood. 1996;88(9):3424-31. 6. Smith AG. Mouse embryo stem cells: their identification, propagation and manipulation. Semin Cell Biol. 1992;3(6):385-99. 7. Asahara T, Masuda H, Takahashi T, Kalka C, Pastore C, Silver M, et al. Bone marrow origin of endothelial progenitor cells responsible for postnatal vasculogenesis in physiological and pathological neovascularization. Circ Res. 1999;85(3):221-8. 8. Asahara T, Takahashi T, Masuda H, Kalka C, Chen D, Iwaguro H, et al. VEGF contributes to postnatal neovascularization by mobilizing bone marrow-derived endothelial progenitor cells. EMBO J. 1999;18(14):3964-72. 9. Hristov M, Erl W, Weber PC. Endothelial progenitor cells: mobilization, differentiation, and homing. Arterioscler Thromb Vasc Biol. 2003;23(7):1185-9. 10. Kalka C, Masuda H, Takahashi T, Kalka-Moll WM, Silver M, Kearney M, et al. Transplantation of ex vivo expanded endothelial progenitor cells for therapeutic neovascularization. Proc Natl Acad Sci U S A. 2000;97(7):3422-7. 11. Suratt BT, Cool CD, Serls AE, Chen L, Varella-Garcia M, Shpall EJ, et al. Human pulmonary chimerism after hematopoietic stem cell transplantation. Am J Respir Crit Care Med. 2003;168(3):318-22. 12. Granton J, Langleben D, Kutryk MB, Camack N, Galipeau J, Courtman DW, et al. Endothelial NO-Synthase Gene-Enhanced Progenitor Cell Therapy for Pulmonary Arterial Hypertension: The PHACeT Trial. Circ Res. 2015;117(7):645-54. 13. Qi Y, Qian L, Sun B, Chen C, Cao Y. Circulating CD34(+) cells are elevated in neonates with respiratory distress syndrome. Inflamm Res. 2010;59(10):889-95. 14. Basile DP, Yoder MC. Circulating and tissue resident endothelial progenitor cells. J Cell Physiol. 2014;229(1):10-6. 15. Critser PJ, Yoder MC. Endothelial colony-forming cell role in neoangiogenesis and tissue repair. Curr Opin Organ Transplant. 2010;15(1):68-72. 16. Asahara T, Murohara T, Sullivan A, Silver M, van der Zee R, Li T, et al. Isolation of putative progenitor endothelial cells for angiogenesis. Science. 1997;275(5302):964-7. 17. Hill JM, Zalos G, Halcox JP, Schenke WH, Waclawiw MA, Quyyumi AA, et al. Circulating endothelial progenitor cells, vascular function, and cardiovascular risk. N Engl J Med. 2003;348(7):593-600.

18. Vasa M, Fichtlscherer S, Aicher A, Adler K, Urbich C, Martin H, et al. Number and migratory activity of circulating endothelial progenitor cells inversely correlate with risk factors for coronary artery disease. Circ Res. 2001;89(1):E1-7. 19. Prater DN, Case J, Ingram DA, Yoder MC. Working hypothesis to redefine endothelial progenitor cells. Leukemia. 2007;21(6):1141-9. 20. Evans MJ, Kaufman MH. Establishment in culture of pluripotential cells from mouse embryos. Nature. 1981;292(5819):154-6. 21. Bulic-Jakus F, Ulamec M, Vlahovic M, Sincic N, Katusic A, Juric-Lekc G, et al. Of mice and men: teratomas and teratocarcinomas. Coll Antropol. 2006;30(4):921-4. 22. Thomson JA, Itskovitz-Eldor J, Shapiro SS, Waknitz MA, Swiergiel JJ, Marshall VS, et al. Embryonic stem cell lines derived from human blastocysts. Science. 1998;282(5391):1145-7. 23. Shamblott MJ, Axelman J, Wang S, Bugg EM, Littlefield JW, Donovan PJ, et al. Derivation of pluripotent stem cells from cultured human primordial germ cells. Proc Natl Acad Sci U S A. 1998;95(23):13726-31. 24. Takahashi K, Yamanaka S. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell. 2006;126(4):663-76. 25. Hanna J, Wernig M, Markoulaki S, Sun CW, Meissner A, Cassady JP, et al. Treatment of sickle cell anemia mouse model with iPS cells generated from autologous skin. Science. 2007;318(5858):1920-3. 26. Burnham EL, Taylor WR, Quyyumi AA, Rojas M, Brigham KL, Moss M. Increased circulating endothelial progenitor cells are associated with survival in acute lung injury. Am J Respir Crit Care Med. 2005;172(7):854-60.

Chapter 3A: Transplantation of Endothelial Colony Forming Cells Improves Survival of FOXF1- deficient Mice

Hannah M. Flood1,2, Yufang Zhang1,2, Xiaomeng Ren1,2, Yang Lin3, Mervin Yoder3, and Vladimir V. Kalinichenko1,2

1Department of Pediatrics, Cincinnati Children’s Research Foundation, Cincinnati, Ohio, USA. 2Center for Lung Regenerative Medicine, Cincinnati Children’s Research Foundation, Cincinnati, Ohio, USA. 2Department of Pediatrics, Indiana University School of Medicine, Indianapolis, Indiana, USA.

The work presented in Chapter 3, Part A was published as an abstract at the following conference:

Flood HM, Zhang Y, Ren X, Lin Y, Yoder M, Kalinichenko VV. Transplantation of Endothelial Progenitor Cells Improves Survival of FOXF1-deficient Mice. Abstract: Regenerative Mechanisms and Therapeutic Intervention Meeting. Indianapolis, IN. 2016.

Abstract

During lung homeostasis, the endothelial barrier regulates the passage of nutrients and fluids.

After acute lung injury (ALI), endothelial permeability is increased which causes edema and inflammation. Our lab recently demonstrated that the transcription factor FOXF1 promotes normal lung homeostasis and repair through S1P/S1PR1 signaling, which regulates endothelial barrier function. However, novel ALI treatment approaches to increase patient survival rates remain elusive. In the present study, we utilized a recently published murine model of ALI that develops after endothelial-specific deletion of Foxf1. We then introduced endothelial colony forming cells (ECFCs) generated in vitro into the bloodstream of FOXF1-deficient mice with the purpose of improving endothelial barrier function. Inactivation of Foxf1 in lung endothelial cells resulted in lethality of 100% of mice by day 24 post-tamoxifen administration. Introduction of

ECFCs into tail vein of mice resulted in a 53.8% increase in survival of FOXF1-deficient mice, which was associated with a marked decrease in lung injury. Immunohistochemical analysis demonstrated a maintenance of normal endothelial architecture after ECFC-injection.

Surprisingly, the introduction of ECFCs did not decreased inflammatory response. Altogether,

ECFCs protect FOXF1-deficient mice from lung inflammation and improve mouse survival. We were unable to determine if ECFCs integrated into pulmonary blood vessels or lung parenchyma, therefore, further studies need to determine the precise mechanism by which the introduction of

ECFCs reduced the inflammatory response, lung injury, and death of FOXF1-deficient mice.

Introduction

During lung homeostasis, the endothelial barrier regulates the passage of nutrients and fluids and is additionally responsible for alveolar gas exchange. After acute lung injury (ALI), endothelial permeability is increased which causes edema and inflammation and results in respiratory failure [1]. ALI, with the complication of life-threatening acute respiratory distress syndrome (ARDS), is the annual cause of approximately 75,000 deaths and 3.5 million hospitalization days [2, 3]. ARDS is additionally the cause of death in more than 35% of ALI patients [4, 5]. ALI and ARDS management is difficult, and treatments remain elusive [4, 6].

FOXF1 is a transcription factor expressed in lung endothelial cells and known to be critical for endothelial barrier function [7]. Endothelial cells are known to aid in lung repair after ALI. We recently published that endothelial loss of FOXF1 resulted in the loss of endothelial integrity and disruption of endothelial barrier function and that restoration of FOXF1 promotes normal lung homeostasis and repair through S1P/S1PR1 signaling [7].

FOXF1 is critical for lung angiogenesis during embryonic development [8-10]. In mice, global deletion of FOXF1 is embryonic lethal [10], but a subset of mice heterozygous for FOXF1 will survive with a variety of lung developmental defects [9, 11, 12]. Mutations in FOXF1 are found in patients with the rare congenital disorder alveolar capillary dysplasia with misalignment of pulmonary veins (ACD/MPV) [13-15]. ACD/MPV patient mortality typically occurs within the first month of life due to the severity of lung developmental defects and respiratory insufficiency along with developmental defects of the alveolar capillary network, vein positioning, and the resulting respiratory insufficiency [16-18]. Current ACD/MPV treatment options are limited to supportive therapies, rendering a need for more-effective treatments [19].

For more severe lung disorders such as chronic obstructive pulmonary disease, an inflammatory

lung disorder resulting in obstructed airflow from the lungs, the most promising treatment for patients is lung transplant, which is limited by lung availability and patient matches [20]. While life-extending and with a 5-year survival rate of 54% [21], this risky treatment approach is not always a solution. It is, therefore, prudent to investigate new avenues for lung disease treatments.

Peripheral blood studies have shown that improved survival in ALI patients is associated with an increase in numbers of circulating endothelial progenitor cells (EPCs) [22]. Furthermore, neonatal patients with respiratory distress syndrome were more likely to survive if they had higher EPC concentrations circulating in their blood, suggesting that EPCs may be involved in regeneration of neonatal lung after injury [23].

In the current study, we utilized a recently published murine model of lung injury that develops after endothelial-specific deletion of FOXF1 [7]. We introduced endothelial colony forming cells (ECFCs) generated in vitro into the bloodstream of FOXF1-deficient mice with the purpose of improving endothelial barrier function. ECFCs are a subset of EPCs that have been cultured for an extended period of time until colonies emerge and display a distinctive cobblestone morphology [24]. Phenotypically similar to endothelial cells, ECFCs can form de novo vessels in vivo and can form tubes in vitro [25]. Our hypothesis was that ECFCs would integrate into the lung endothelium of injured mice to reduce the incidence of pulmonary injury and prevent mortality in FOXF1-deficient mice. The goal of our study was to determine novel mechanistic avenues that could lead to new ALI and ARDS treatment approaches to increase patient survival rates through the promotion of endothelial regeneration by ECFC transplantation.

Results

Endothelial FOXF1 deletion-dependent ALI and death rescued by ECFCs. Recent studies have shown that intraperitoneal injection of bone-marrow derived cell populations with EPC characteristics improves the extent of alveolar disruption after hyperoxia treatment in neonatal mice [26]. To investigate if ECFCs could have similar therapeutic results in adult mice with acute lung injury, we utilized collaborator-derived ECFCs and a recently published mouse model of ALI [7]. Transgenic mice containing a tamoxifen-inducible Pdgfb-CreER transgene and two

Foxf1-floxed alleles (PdgfbCreER;Foxf1fl/fl) were generated by breeding Pdgfb-CreER and

Foxf1fl/fl mice (Fig. 3.1A). ALI was induced with the deletion of Foxf1 through subsequent administrations of tamoxifen on days -6 and -5 (Fig. 3.1A-B). As previously reported [7], 100% mortality occurred by day 24 post-tamoxifen administration in PdgfbCreER;Foxf1-/- mice (Fig.

3.1C). After ECFC transplantation, mouse survival was increased by 53.8% with 7/13 healthy mice surviving to study completion. Thus, introduction of ECFCs post ALI was sufficient to increase mouse survival.

Pulmonary architecture is maintained and lung injury is attenuated after ECFC transplantation.

While no ECFC integration was observed, after transplantation, we investigated the 53.8% increase in mouse survival. Morphological analysis of lung sections revealed a reduction in injury after ECFC transplantation as observed through H&E staining (Fig. 3.2A). Masson’s trichrome staining revealed that collagen depositions coincided with areas of lung injury but were absent in normal lung portions of ECFC transplanted mice (Fig. 3.2B). In all, transplantation of the ECFCs attenuated lung injury caused by loss of FOXF1.

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Figure 3.1. Introduction of ECFCs increases overall mouse survival after FOXF1-deficiency- induced acute lung injury. (A) Diagram demonstrates Pdgfb-CreER transgene with LoxP sites flanking Foxf1 Exon 1 (encoding DNA-binding domain). (B) Schematic illustrates tamoxifen and ECFC treatment protocols. (C) Kaplan-Meyer curve shows uniform mortality in PdgfbCreER;Foxf1-/- mice by day 24 post-initial tamoxifen injection. After ECFC transplantation, Foxf1-deficient mice demonstrated a 53.8% increase in survival.

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Figure 3.2. ECFC transplantation attenuates pulmonary injury. (A) H&E staining demonstrates a decrease in lung injury in ECFC-transplanted, Foxf1-deficient mice. (B) Masson’s trichrome staining demonstrates that collagen deposition is restricted to injured regions in PdgfbCreER;Foxf1-/- mice after ECFC transplantation.

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No integration of transplanted ECFCs was observed after ALI. Since our hypothesis was that

ECFCs would integrate into the lung endothelium of injured mice, the ECFCs utilized were developed with a tdTomato tag so we could track the cells in vivo. To investigate if the transplanted ECFCs integrated into the host lung endothelial system, we evaluated co- localization of endothelial marker endomucin (EMCN) with RFP to detect the tdTomato tag

(Suppl. Fig. 3.3). No integration of ECFCs was observed in pulmonary architecture (Fig. 3.3A).

Immunohistochemical staining was performed to check if ECFCs lost tdTomato fluorescence during tissue processing; however, no positive cells were detected in lung tissue (Fig. 3.3B).

Since mouse survival was increased but ECFC integration was not observed, we evaluated mRNA expression of endothelial markers Kdr (Vegfr2, Flk-1, Cd309) and Cdh5 (Ve-cadherin,

Cd144). Surprisingly, Kdr and Cdh5 mRNA levels were not rescued in ALI lungs after ECFC transplantation (Fig. 3.3C-D). Altogether, further studies are needed to determine if the ECFCs integrate into the lungs.

Endothelial architecture is maintained after ECFC transplantation with no prevention of macrophage infiltration. Since pulmonary architecture was maintained after ECFC transplantation, we wanted to examine endothelial marker expression. Endothelial cells were detected in lung sections by PECAM staining (Fig. 3.4A). PECAM is expressed in the injured areas of lungs after loss of Foxf1; however, in uninjured areas of ECFC transplanted lungs, endothelial architecture is maintained, though this expression is reduced compared to non-injured lungs as shown by Pecam1 mRNA levels (Fig. 3.4B). Since recruitment of inflammatory cells to lung is characteristic of ALI [4, 27] and macrophages are a major component of the

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Figure 3.3. ECFCs integration was not observed in FOXF1-deficient lung. (A) Co- immunofluorescence of endothelial marker endomucin (EMCN), and endogenous RFP (tdTomato-tagged ECFCs) demonstrate no ECFC integration into pulmonary architecture. (B) tdTomato immunohistochemical staining shows that cells were not detected in lung parenchyma. Positive control inset is a representative tdTomato-tagged tumor cell line injected into mouse lung. (C-D) qRT-PCR analysis demonstrated a decrease in Kdr (Vegfr2, Flk-1, Cd309) and Cdh5 (Ve-cadherin, Cd144) mRNAs in PdgfbCreER;Foxf1-/- lungs with or without ECFC transplantation. mRNAs were normalized to Actb. P-values for Kdr mRNA levels: WT+ECFC to KO+Con: P=0.2032; WT+ECFC to KO+ECFC: P=0.2681; KO+Con to KO+ECFC: P=0.2667. P-values for Cdh5 mRNA levels: WT+ECFC to KO+Con: P=0.1848; WT+ECFC to KO+ECFC: P=0.2053; KO+Con to KO+ECFC: P=0.4820.

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Figure 3.4. ECFC transplantation allows for maintenance of endothelial architecture but did not reduce number of macrophages. (A) PECAM staining highlights a maintenance of lung endothelial architecture in ECFC-transplanted, Foxf1-deficient mice. (B) qRT-PCR analysis demonstrated a decrease in Pecam1 (Cd31) mRNAs with or without ECFC transplantation. mRNAs were normalized to Actb. P-values for Pecam1 mRNA levels: WT+ECFC to KO+Con: P=0.1927; WT+ECFC to KO+ECFC: P=0.2250; KO+Con to KO+ECFC: P=0.4647. (C-D) MAC3 cell counts and MAC3 staining shows a severe increase in lung inflammatory cells in PDGFbCreER;Foxf1-/- mice with no reduction in number of inflammatory cells after ECFC transplantation. P-values for MAC3 cell counts: WT+ECFC to KO+Con: P=0.0740; WT+ECFC to KO+ECFC: P=0.1590; KO+Con to KO+ECFC: P=0.9472.

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inflammatory process [28], we evaluated macrophage marker MAC3. MAC3 staining revealed that even with ECFC transplantation, the number of observed inflammatory cells was unchanged

(Fig. 3.4C-D). This may be due to the incomplete rescue observed in our morphological analysis.

FOXF1 expression is decreased in PdgfbCreER;Foxf1-/- mice with or without ECFC transplantation. Although pulmonary architecture is maintained after ECFC transplantation, we observed neither cell integration nor reduced numbers of inflammatory cells. Since FOXF1 is known to be expressed in lung endothelial cells and is important for normal lung homeostasis and repair, we examined FOXF1 in the ECFC. FOXF1 protein and Foxf1 mRNA were decreased in PdgfbCreER;Foxf1-/- lungs even after ECFC transplantation (Fig. 3.5A-C). In order to analyze

FOXF1 expression in the injected ECFC, we performed flow cytometry on ECFCs and compared them to endothelial cells from WT mouse lung. We found that FOXF1 was not expressed in the

ECFCs (Fig. 3.5D). Unpublished lab data has suggested that C-KIT expression is necessary for lung integration. Therefore, we additionally checked lung mRNA levels for Kit and found that they remained decreased after ECFC transplantation (Fig. 3.5E), suggesting that a progenitor cell population was not permanently restored to the lung after injury and ECFC transplantation.

Altogether, these data indicate that the cells utilized for transplantation were FOXF1-C-KIT- and were unlikely unable to integrate; however, more work will need to be done to confirm this hypothesis.

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Figure 3.5. FOXF1 levels are not restored after ECFC transplantation. (A-B) Western blot shows total lung protein levels of FOXF1 are decreased in PdgfbCreER;Foxf1-/- mice with or without ECFC transplantation. FOXF1 levels were internally normalized to ACTIN for each sample. P-values for FOXF1 protein levels: WT+ECFC to KO+Con: P=0.2509; WT+ECFC to KO+ECFC: P=0.2098; KO+Con to KO+ECFC: P=0.3192. (C) qRT-PCR analysis demonstrated a decrease in Foxf1 mRNAs in PdgfbCreER;Foxf1-/- lungs with or without ECFC transplantation. mRNAs were normalized to Actb. P-values for Foxf1 mRNA levels: WT+ECFC to KO+Con: P=0.1941; WT+ECFC to KO+ECFC: P=0.1951; KO+Con to KO+ECFC: P=0.5776. (D) Flow cytometry was used to demonstrate that transplanted ECFC were negative for FOXF1. Wild Type (WT) lung endothelial cells from C57Bl/6 mice were used as a control. (E) qRT-PCR analysis demonstrated a decrease in Kit mRNAs in PdgfbCreER;Foxf1-/- lungs with or without ECFC transplantation. mRNAs were normalized to Actb. P-values for Kit mRNA levels: WT+ECFC to KO+Con: P=0.2111; WT+ECFC to KO+ECFC: P=0.2111; KO+Con to KO+ECFC: P=0.9808.

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Discussion

During ALI, endothelial permeability is increased, resulting in pulmonary edema and inflammation. FOXF1 is known to be critical for endothelial barrier function and loss of the transcription factor leads to ALI and death in a recently published murine model [7]. Treatment options for ALI remain elusive. In the present study, we showed that transplantation of ECFCs attenuates ALI in FOXF1-deficient mice and improves mouse survival. Morphological analysis demonstrated a decrease in injured areas in PdgfbCreER;Foxf1-/- + ECFC lungs compared to control-treated PdgfbCreER;Foxf1-/- lungs. Our studies additionally demonstrated a decrease in collagen deposits, which were correlated with injured areas in PdgfbCreER;Foxf1-/- + ECFC lungs. Altogether, ECFC transplantation was able to prevent extensive lung injury, representing a possible novel treatment approach for ALI, ARDS, and other lung disorders.

After ECFC transplantation, it was found that the transplanted cells did not integrate into the existing lung structure. Injected ECFCs did not co-express with EMCN staining and were not observed in immunohistochemical analysis. On this point, either the ECFCs did not express factors that would allow for lung integration or alternatively the tools used were insufficient to trace the cell. Flow analysis of the transplanted ECFCs revealed only 6.5% RFP expression, suggesting that these cells were untraceable in vivo. Additionally, endothelial cell markers Kdr and Cdh5 were not increased after ECFC transplantation, even with the 53.8% improvement of mouse survival. It is possible that the increase in survival may have been due to secreted factors.

Surprisingly, immunohistochemical analysis of endothelial marker PECAM revealed a maintenance of endothelial morphology after ECFC transplantation in PdgfbCreER;Foxf1-/- lungs; however, Pecam1 mRNA levels were not improved after ECFC transplantation.

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Interestingly, while morphology was improved in PdgfbCreER;Foxf1-/- lungs after ECFC transplantation, the number of macrophages identified through immunohistochemical staining of

MAC3 was unaltered. Since inflammation is a hallmark of ALI and macrophages are a major component of the inflammatory process [28], it seems that ECFC transplantation is not sufficient to reverse all effects of the disorder. Since the surviving mice in the ALI and ECFC-transplanted cohort were healthy at the conclusion of the study at day 20, it would be interesting to monitor the response over a longer period of time to see if and how quickly the lungs were able to clear the macrophages.

Altogether, the transplanted ECFCs were either untraceable or did not integrate. They did not express FOXF1, which is a factor necessary for proper endothelial barrier function [7] and may account for the infiltration of macrophages in PdgfbCreER;Foxf1-/- + ECFC lungs. While the precise mechanism by which the introduction of ECFCs reduced the inflammatory response, lung injury, and death of FOXF1-deficient mice is currently unknown, the present study was able to improve lung morphology and increase mouse survival after ALI. Endothelial progenitor cell transplantation may be an interesting treatment option for patients with lung disorders. Future studies will need to first focus on the development of novel tools to overcome the obstacles outlined here with a traceable, FOXF1-expressing endothelial progenitor cell line to utilize in further experiments, and second, to determine how the transplanted cells are able to improve lung morphology and increase survival.

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Methods

Mice.

The Foxf1fl/fl mouse line was previously generated and bred into the C57Bl/6 mouse background

[29]. Foxf1fl/fl mice were bred with Pdgfb-CreER mice to generate PdgfbCreER;Foxf1fl/fl mice

[30]. PdgfbCreER;Foxf1fl/fl mice were bred with Foxf1fl/fl mice and adult mice were used for all experiments. Foxf1fl/fl littermates were used as controls. To activate the conditional Foxf1 knockout, tamoxifen (Tam) was given via intraperitoneal injection (40 mg/kg of body weight;

Sigma) at days -6 and -5 before ECFC administration. tdTomato-labeled ECFCs or control media were injected into tail vein of PdgfbCreER;Foxf1-/- or Foxf1fl/fl mice at days 0 and 3. To collect survival data, mice were observed daily. All animal studies were approved by the Institutional

Animal Care and Use Committee (IACUC) of Cincinnati Children’s Research Foundation and performed in accordance with the NIH IACUC Guidebook. All experiments were covered under our animal protocol (IACUC2016-0038). The Cincinnati Children’s Research Foundation

Institutional Animal Care and Use Committee is an AAALAC and NIH accredited institution

(NIH Insurance #8310801).

Histology and immunohistochemistry.

Paraffin-embedded liver sections were used for H&E, or immunohistochemistry (IHC) as previously described [31-33]. Frozen sections were used for immunofluorescence (IF). The following antibodies were used for immunostaining: EMCN [1:200 IF, (Abcam)], tdTomato

[1:1500 IHC, (LS Bio)], PECAM [1:500 IHC, (BD Pharmingen)], and MAC3 [1:3000 IHC; (BD

Pharmingen)]. Antibody-antigen complexes were detected using biotinylated secondary antibodies followed by avidin-biotin-horseradish peroxidase complex and 3,3’diaminobenzidine substrate (Vector Labs) as previously described [31, 32, 34]. Sections were counterstained with

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nuclear fast red (Vector Labs). For immunofluorescence imaging, secondary antibody conjugated with Alexa Fluor 488 (Invitrogen/Molecular Probes) was used as described [35, 36]. Cell nuclei were counterstained with DAPI (Vector Labs). Masson’s trichrome (BD Biosciences) staining was performed according to manufacturer’s protocol. Brightfield images were obtained using a

Zeiss AxioImage.A2 microscope. Fluorescent images were obtained using a Zeiss AxioPlan 2 microscope. qRT-PCR and protein.

The left lobe of the lung was halved and used for RNA and protein studies. RNA was isolated using RNA Stat-60 (Tel-Test, Inc.) according to manufacturer protocol and was reverse transcribed using the High Capacity Reverse Transcription Kit (Applied Biosystems) according to manufacturer protocol. mRNAs of specific genes were measured by qRT-PCR using TaqMan probes (Applied Biosystems) and the StepOnePlus Real-Time PCR system (Applied Biosystems) as described [36-39]. The following Taqman probes were used for qRT-PCR: Kdr

(Mm01222421_m1), Cdh5 (Mm00486938_m1), Pecam1 (Mm01242576_m1), Foxf1

(Mm00487497_m1), and Kit (Mm00445212_m1). mRNAs were normalized to Actb

(Mm00607939_s1). Protein extracts were isolated using cell lysis buffer as previously described

[40] and used for Western blot analysis with Pierce ECL Western blotting substrate (Thermo

Scientific) according to manufacturer protocol. The following antibodies were used for protein blots: FOXF1 (1:1000, R&D Systems) [30, 41, 42] and ACTIN (1:2000; Santa Cruz) [43].

Protein band intensities were determined by ImageJ software and were normalized to ACTIN.

Flow cytometry.

Flow cytometry experiments were performed as previously described [44]. Briefly, cells were filtered using a 70μm nylon mesh (Corning) and washed with cell sorting buffer (CSB; Recipe:

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50mL phosphate buffered saline (Corning), 500μL fetal bovine serum (BDBiosciences), 500μL

0.5M EDTA (Fisher)). Cells were re-suspended in CSB and analyzed for RFP expression using

FACSAria II. FlowJo software was used to analyze data. Viability dye (BioLegend) was used to label dead cells.

Statistical analysis.

Student’s t-test was used to determine statistical significance. P<0.05 was considered to be significant. Values for all measurements were expressed as mean ± standard error of mean.

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Chapter 3B: Differentiation of Novel GFP:FOXF1 Embryonic Stem Cell Line into FOXF1- positive Endothelial Progenitor Cells

Hannah M. Flood1,2, Olena Kolesnichenko1,2, Craig Bolte1,2, and Vladimir V. Kalinichenko1,2

The work presented in Chapter 3, Part B was published as an abstract at the following conference:

Flood HM and Kalinichenko VV. Differentiation of Novel GFP:FOXF1 Embryonic Stem Cell Line into FOXF1-positive Endothelial Progenitor Cells. Abstract: Jensen Symposium. Cincinnati, OH. 2018.

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Abstract

Pulmonary endothelial cells are critical for endothelial barrier maintenance and lung homeostasis. Acute lung injury (ALI) increases endothelial permeability, which results in edema and inflammation. ALI can develop into the life-threatening complication acute respiratory distress syndrome. Recent advances in treatment of human lung diseases have introduced cell replacement strategies to contribute to lung regeneration. Preliminary studies in our lab have shown that the introduction of endothelial colony forming cells prior to ALI in mice reduces the extent of injury and improves survival. However, the cells utilized in the preliminary studies did not express a fluorescent tracer for monitoring in vivo integration, and were additionally negative for FOXF1, a transcription factor that has been shown to be expressed in lung endothelial cells and important for proper lung function. The current study sought to first develop a traceable cell line to determine if the transplanted cells integrate into the lung barrier to rescue function and phenotype, and second to differentiate into a sub-population of endothelial progenitor cells that expresses FOXF1. Using the CRISPR/Cas9 system, we successfully generated a novel embryonic stem cell line where we have knocked GFP in to the Foxf1 locus. We have reviewed previously published endothelial cell differentiation protocols to develop a novel differentiation method which yields ~95% endothelial cells. The majority of these endothelial cells are positive for stemness marker C-KIT, suggesting that they have successfully differentiated into an endothelial progenitor cell (EPC) population. Further analysis demonstrated that our EPC population is ~51.5% FOXF1-positive as detected by GFP:FOXF1. Altogether, we have successfully developed a differentiation protocol which yields ~51.5% FOXF1-positive endothelial progenitor cells.

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Introduction

Pulmonary endothelial cells (ECs) are critical for endothelial barrier maintenance and lung homeostasis, and are known to aid in lung repair after injury. Acute lung injury (ALI) increases endothelial permeability, which results in edema and inflammation [1]. ALI can develop into the life-threatening complication acute respiratory distress syndrome [2, 3].

Improved survival of patients with acute lung injury is associated with increased numbers of circulating endothelial progenitor cells (EPCs) in peripheral blood studies [4]. Moreover, EPCs may be involved in regeneration of neonatal lung after injury as indicated by increased survival of neonatal respiratory distress syndrome patients with higher concentrations of EPC circulating in their blood [5]. For severe lung disorders such as chronic obstructive pulmonary disease, an inflammatory lung disorder resulting in obstructed air flow from the lungs, treatment often requires lung transplantation; however, a major impediment is the limited availability of lungs and patient matches [6]. In the few cases where there is a genetic match, this procedure carries high risk, despite a 5-year survival rate of 54% [7]. Therefore, it is necessary to explore novel treatments for lung disease.

Advances in treatment of human lung diseases have introduced cell replacement strategies to contribute to lung tissue repair and regeneration. EPC transplantation therapy has recently emerged as a beneficial treatment for many chronic lung diseases due to the ability of embryonic stem cells (ESC) to home to injury sites [8, 9] and their capability to differentiate into endothelial cells and generate de novo blood vessels [8]. A retrospective study of female patients who received hematopoietic stem cell transplants from male donors revealed 37.5-42.3% endothelial chimerism in lung biopsies [10], highlighting the therapeutic potential of both ESCs and EPCs for treatment of lung diseases. A recent phase I trial for pulmonary hypertension treatment demonstrated that introduction of EPCs into the pulmonary artery resulted in

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improvement of 6-minute walk capacity of patients, which reflects a complex physiological response involving the pulmonary, cardiovascular, circulatory, and neuromuscular systems [11].

Preliminary studies in our lab have shown that the introduction of endothelial colony forming cells prior to ALI in mice reduces the extent of injury and improves survival (Chapter 3,

Part A). However, the cells utilized in the preliminary studies did not express a fluorescent tracer for monitoring in vivo integration, and were additionally negative for FOXF1, a transcription factor that has been shown to be expressed in lung endothelial cells and important for proper lung function [12]. Therefore, the first objective of our study was to establish an ESC line with a

GFP:FOXF1 knock-in reporter for investigating the transcriptional role of FOXF1.

Since restoration of endothelial function in patients with vascular diseases may be possible if clinically-relevant numbers of vessel-forming cells can be derived in vitro, we next sought to differentiate our newly-developed GFP:FOXF1 ESC line into a sub-population of

EPCs that express FOXF1. In 1996, Vittet et al. demonstrated that endothelial cells matured through distinct steps and that endothelial expression markers could be detected at specific stages of cell maturation [13]. This study followed EC differentiation from embryoid bodies, which are spontaneously-differentiated cyst-like structures containing derivatives from all three germ layers [14] to identify EC markers during cell maturation, demonstrating that EC commitment follows sequential maturation stages [13]. We utilized previously published methods to develop a novel differentiation protocol that yielded FOXF1+ endothelial progenitor cells.

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Results

Novel CRISPR/Cas9 embryonic stem cell line expresses GFP to track FOXF1 expression. It has been previously established that FOXF1 is an important factor for embryonic development and lung organogenesis [1-3]. FOXF1 is additionally important for maintenance of lung barrier function after acute lung injury [12]. Therefore, to monitor expression of this important transcription factor, we utilized the CRISPR/Cas9 system to knock GFP into the Foxf1 locus of an established ESC line (Fig. 3.6 A). The W4/129S6 ESC line (hereafter referred to as W4 or parental line) is a wild type ESC line derived from 129s6 mice at the blastocyst stage. We confirmed by PCR the site-specific integration of GFP into the Foxf1 locus. The PCR screen revealed two homozygous clones and three heterozygous clones (Fig. 3.6 B, Suppl. Fig. 3.2,

Suppl. Fig. 3.3). One homozygous clone (G1) was found to not express detectable GFP (data not shown); however, the other homozygous clone (A1) was found to express low levels of GFP

(Fig. 3.7 A-C) and was utilized for all experiments (hereafter referred to as A1 or GFP:FOXF1).

Novel GFP:FOXF1 ESC line maintains stemness characteristics. Since our goal was to differentiate the A1 cells into FOXF1+ endothelial cells, it was important to determine if the

CRISPR/Cas9 insertion altered their stem cell characteristics. The GFP:FOXF1 cell line grew indistinguishably from the parental cell line and was morphologically similar in appearance (Fig.

3.8A). Evaluation of mRNA expression of known stemness markers Nanog [15] and Sox2 [16] showed no difference between the GFP:FOXF1 knock-in ESC line and the parental ESC line

(Fig. 3.8 B). We have, therefore, generated a novel ESC line that acts similar to the established parental line with the addition of GFP:FOXF1.

Differentiation of EPCs from ESCs requires activation of vascular progenitor cell pathways.

During lung organogenesis, pulmonary vascular specification requires a variety of specific

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Figure 3.6. Establishment of GFP:FOXF1 ESC line by CRISPR/Cas9-mediated knock-in. (A) Schematic illustration of the CRISPR/Cas9 strategy to knock 2AGFP (highlighted in yellow) into the FOXF1 locus. The upper diagram shows the gRNA target site (underlined) with the Protospacer Adjacent Motif (PAM) sequence (highlighted in blue). The position of the 2A-GFP reporter is indicated with an arrow. The middle diagram shows the targeted exon (exon 2) of Foxf1 with the construct knock-in location indicated (dashed lines). The bottom diagram represents the GFP knock-in allele. (B) The cells were screened by PCR (full length gel shown) and A1 was identified as a homozygous clone. The expected fragment sizes are indicated for WT (317 bp; yellow *) and knock-in (162 bp; green *). Primers are described in Suppl. Fig. 3.2 with additional PCR screens. Additional clones identified are described in Suppl. Tab. 3.1.

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Figure 3.7. Novel GFP:FOXF1 (A1) cell line expresses GFP. (A) Flow cytometry shows representative dot plots and (B) quantification of the percentage of endogenous GFP levels in the A1 cell line. Cells were not stained for live/dead markers; only endogenous GFP levels were tested. (C) Image of undifferentiated GFP:FOXF1 (A1) cells showing GFP-positive cells within the cell colony after 9 days in culture as detected through fluorescent imaging. P-values for GFP expression levels: W4 to A1 in GFP-Low: P=0.0509; GFP-Low to GFP-High for A1: P=0.0929.

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Figure 3.8. Novel GFP:FOXF1 (A1) cell line is similar in morphology and stemness marker expression to parental (W4) cell line. (A) Bright field images of the parental (W4) cell line and the GFP:FOXF1 (A1) cell line show a similar appearance after 3 days in culture. (B) qRT-PCR analysis demonstrated no difference in mRNA levels of stemness markers Nanog or Sox2 between parental and GFP:FOXF1 cell lines. mRNA levels were normalized to Gapdh. P-values for mRNA expression levels: Nanog W4 to A1: P=0.3366; Sox2 W4 to A1: P=0.3793.

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signals. Endothelial cell commitment follows sequential maturation stages through distinct steps

[13]. To derive endothelial progenitor cells from A1 ESCs, we utilize a novel combination of growth factors. ESC maintenance media (2i media) was removed and replaced with -2i differentiation media supplemented with Activin A, BMP4, and EGF to prime the ESCs for 24 hours. The priming period was followed by the reprograming period with -2i differentiation media supplemented with the growth factors VEGF165, FGF2, EGF, and SHH for differentiation into endothelial cells (Fig. 3.9 A).

After the second day of cell differentiation, we evaluated mRNA levels of the endothelial cell marker, Pecam1 (Cd31). On day 5 of differentiation, Pecam1 expression was increased 3- fold in both W4 and A1 ESC lines (Fig. 3.9 B). We additionally evaluated mRNA levels of

Foxf1 to ensure its expression in our differentiated cells at days 2 and 5 (Fig. 3.9 C). There was no difference between the GFP:FOXF1 and parental cell lines, showing that our novel A1 cell line behaves similarly to the parental line during endothelial differentiation.

Highly efficient EPC differentiation. To further evaluate and characterize the endothelial cells derived from A1 and W4 ESC lines, we performed flow cytometry. After removing cell debris and doublets (Suppl. Fig. 3.4), endothelial cells were sorted from among live cells using the classic definition of endothelial cells (CD31+CD45-). Our novel differentiation method yielded

95.2% endothelial cells in the GFP:FOXF1 line (Fig. 3.10 A), similar to the 97.2% yielded by the parental line (Fig. 3.10 B). This highly efficient differentiation method was further evaluated by flow cytometry for the progenitor cell marker, C-KIT [17], with the majority of the cells in each line being C-KIT+ (Fig. 3.10 A-B). These data suggest that we have successfully differentiated our ESCs into endothelial progenitor cells. Furthermore, while the parental W4 line expectedly did not express GFP, our A1 line was > 50% FOXF1+ (Fig. 3.10 A-B).

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Figure 3.9. Generation of EPCs from ESCs through novel differentiation protocol. (A) Schematic representation of the protocol for the generation of EPCs from mouse ESCs. (B) qRT- PCR analysis demonstrated a 3-fold increase in Pecam1 mRNA after 5 days of differentiation in the parental (W4) cell line with a similar 3-fold increase in the GFP:FOXF1 (A1) cell line. P- values for Pecam1 mRNA expression levels: W4 day 2 to day 5: P=0.1031; A1 day 2 to day 5: P=0.0218. (C) qRT-PCR analysis demonstrated a 2-fold increase in Foxf1 mRNA after 5 days of differentiation in the parental (W4) cell line with a 3-fold increase in the GFP:FOXF1 (A1) cell line. P-values for Foxf1 mRNA expression levels: W4 day 2 to day 5: P=0.0816; A1 day 2 to day 5: P=0.1436.

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Figure 3.10. High-yield differentiation of EPCs from ESCs. Flow cytometry dot plots for (A) GFP:FOXF1 (A1) cell line and (B) parental (W4) cell line are shown after 5 days of differentiation for the following markers: live cells, CD31 (PECAM), CD45, C-KIT, and GFP (endogenous). A1 and W4 cell lines both yielded >95% endothelial cells (as defined by CD31+CD45-) with the A1 cells line demonstrating 54.1% of the EPCs as FOXF1+ (as demonstrated through endogenous GFP expression).

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Discussion

In the current study, we developed a novel, murine ESC line to trace FOXF1 expression during cell differentiation. FOXF1 has long been established to be important during embryonic development, lung organogenesis, and for maintenance of lung barrier function [1-3, 12]. We utilized the CRISPR/Cas9 system to knock-in a GFP reporter to the Foxf1 locus in W4/129S cells. This novel cell line, A1, expresses a low GFP fluorescence that is detectable by microscopy and flow cytometry lasers. Our novel GFP:FOXF1 ESC line behaves indistinguishably from the parental line with similar growth patterns, morphology in culture, and stemness marker expression. Altogether, we have successfully generated a novel GFP knock-in reporter system, which will be useful for studying the role of FOXF1 in EPC/EC differentiation and in vivo lung integration.

Advances in treatment of human lung diseases have led to the development of cell replacement strategies to contribute to lung tissue repair and regeneration. Mesenchymal stromal cells (MSCs) have anti-inflammatory properties and have been used in pre-clinical and phase I and II clinical trials for treatment of chronic inflammatory diseases such as acute respiratory distress syndrome [18], idiopathic pulmonary fibrosis [19], and bronchopulmonary dysplasia

[20]. While MSCs do not engraft into existing lung architecture and are not curative, their benefits as a therapy look promising to treat these and other inflammatory lung diseases. EPCs, however, are known to home to injury sites [8, 9], form de novo blood vessels [8], and engraft into the host lung in human patients [10]. In addition, introduction of EPCs was shown to improve patient health in a phase I pulmonary arterial hypertension clinical trial [11]. Together,

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these preliminary studies highlight the potential importance of EPC-based therapies in clinical treatment of pulmonary diseases.

Since previous studies demonstrated the therapeutic potentials of utilizing EPCs to treat multiple lung disorders, we sought to develop a novel method of differentiation which yields

FOXF1+ EPCs. Published methods have used growth factors and inhibitors to stimulate specific signaling pathway activation and promote ESC differentiation towards the endothelial cell fate with varying levels of success. GSK3 inhibition, which stimulates Wnt signaling, along with

BMP4 treatment was demonstrated to commit induced pluripotent stem cells (iPSCs) to a mesodermal fate [21]. Wnt signaling has been shown to direct differentiation to the mesodermal fate [22, 23] and has additionally been shown to be required for primitive streak formation [24].

Thereafter, exposure to VEGF and forskolin, which is typically used in neuronal differentiation, yielded mature endothelial cells with between 61.8% and 88.8% efficiency by day 5 [21]. EC differentiation was evaluated through VE-cadherin (CD144) positivity. These iPSC-derived ECs were transcriptionally similar to primary vascular endothelial cells as determined by gene expression profiling [21]. In a separate study, GSK3 inhibition for 2 days in the absence of exogenous VEGF or FGF signaling was sufficient to induce hPSC differentiation into 55%

CD34+CD31+ EPCs by day 5 [25]. This EPC population could be further directed to a more mature endothelial cell fate (CD31+CD144+CD34-) through maintenance in endothelial growth media [25]. VEGF has been shown to activate the MAPK signaling pathway [26] and FGF2

(bFGF) is known to activate the PI3K signaling pathway [27]. The current study utilized a GSK3 inhibitor in the ESC maintenance media before initiation of differentiation along with VEGF and

FGF2 and additional growth factors in the differentiation media.

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A recent study was able to convert human iPSCs or ESCs into core-blood endothelial colony forming cells (CB-ECFCs) with a stable endothelial phenotype which can form blood vessels in mice [28]. This study encouraged CB-ECFC formation through exposure to BMP4,

FGF2, and VEGF, with an initial 24 hours of exposure to Activin A in addition to the main growth factors. This protocol yielded NRP-1+ CD31+ cells that could then be expanded to

3x10^4 cells in 12 days. This NRP-1+ CD31+ cell population was determined to have CB-ECFC properties [28]. Activin A is a member of the transforming growth factor beta (TGF-β) super family and has been shown to induce endothelial cell differentiation [29]. BMP4, is known to be a key inducer of mesodermal differentiation from ESCs [30, 31]. Another study utilized VEGF,

BMP4, and FGF2 growth factors after GSK3 inhibition to differentiate hiPSCs and hESCs into between 94.1% and 97.4% endothelial cells (as defined by CD31+ and CD34+ or CD144+), which is the highest percentage of ECs differentiated in vitro to date [32]. The current study utilized both Activin A and BMP4 in the priming period of EC differentiation along with EGF in both the priming and differentiation periods. The growth factor EGF is a typical component of endothelial maintenance media and has previously been used in endothelial cell differentiation protocols [33].

While all these previous studies were successful to varying degrees, only the final study was able to produce an efficient number of ECs without the necessity of further sorting [32].

This is similar to what we found in our current study; however, we additionally looked for the expression of FOXF1, which is known to be important for proper lung function [12], and found that our EPCs were 51.5% positive for FOXF1. This was achieved through our novel use of the

SHH growth factor. SHH is known to be an upstream activator of FOXF1 [34] and along with

FOXF1 and BMP4, SHH has recently been implicated in mouse ureter mesenchyme

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development [35]. Ours is the first study to achieve a 95% endothelial cell population with approximately 50% being FOXF1+.

Although the GFP:FOXF1 EPCs described in the current study express the appropriate markers to be considered endothelial progenitor cells, further in vitro work needs to be done to determine if they function as endothelial cells. Future in vivo studies will determine if the

GFP:FOXF1 EPCs are able to integrate into the host lung endothelial system and successfully rescue mouse models of lung injury. In addition, it is clear that within the endothelial cell field, there is no standard protocol for differentiation nor is there a standard definition that is universally used for EPCs. That being said, in the current study, we successfully generated a novel GFP knock-in reporter system, and have developed a novel protocol to differentiate mESC into 95% endothelial progenitor cells without the necessity of further sorting. These tools and methods will be useful for future studies investigating the role of FOXF1 in EPC/EC differentiation and in vivo lung integration.

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Methods

CRISPR/Cas9 and PCR Screen.

W4/129S6 mouse Embryonic Stem Cells (Taconic, Hudson, NY, USA) were utilized as a parental line to generate a novel GFP:FOXF1 ESC line. W4 cells are a wild-type ESC line derived from 129S6 mice at the blastocyst stage (received from Yueh-Chiang Hu, PhD,

CCHMC). CRISPR/Cas9 was performed as previously described by the Transgenic Animal and

Genome Editing Core Facility at the Cincinnati Children’s Hospital Medical Center (Cincinnati,

OH). Briefly, the 2A-GFP construct along with a CRISPR plasmid were electroporated into W4 embryonic stem cells then sorted for GFP+ cells into three 96-well plates. Approximately 30 clones were recovered from the 96-well plates. The site-specific integration of GFP into the

Foxf1 locus was confirmed by PCR. The A1 clone was identified as a homozygous clone. The primers utilized in the PCR screen are described in Suppl. Fig. 3.2.

Cell Culture and Differentiation Method.

Embryonic stem cells were maintained in 2i media on matrigel-coated plates. The 2i media was made according to protocol (2i Media, Center for Regenerative Medicine, Boston Medical,

Boston University, Boston, MA). Sterile cell culture plates (Falcon) were matrigel-coated according to manufacturer instructions (Corning, Matrigel hESC-qualified Matrix). Cells were split at confluency, which was approximately every 3 days before ESC colonies were allowed to touch and merge.

ESC differentiation was begun one day prior to ESC confluence. Then, 2i media was replaced with -2i media supplemented with growth factors. For 24 hours, priming growth factors were used to supplement the -2i media: Activin A (R&D Systems, 10ng/mL), BMP4 (R&D

Systems, 10ng/mL), and EGF (Gold Biotechnology, 10ng/mL). Thereafter, endothelial

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promoting growth factors were used to supplement the -2i media: VEGF165 (Gold

Biotechnology, 10ng/mL), FGF2 (Gold Biotechnology, 10ng/mL), EGF (Gold Biotechnology,

10ng/mL), and SHH (Gold Biotechnology, 10ng/mL). Differentiating ESCs were harvested on days described in each experiment.

Flow cytometry.

Flow cytometry experiments were performed as previously described [36]. Briefly, cells were removed from matrigel-coated plates with dispase (Stemcell Technologies) and were filtered using a 70μm nylon mesh (Corning). Cells were washed with cell sorting buffer (CSB; Recipe:

50mL phosphate buffered saline (Corning), 500μL fetal bovine serum (BDBiosciences), 500μL

0.5M EDTA (Fisher)) and were re-suspended in CSB with antibodies at a 1:100 dilution. Cells were centrifuged, washed in CSB, and analyzed using FACSAria II. FlowJo software was used to analyze data. The following antibodies were used for FACS: CD117 (BV711, BioLegend),

CD31 (BV605, BioLegend), CD144 (APC, BioLegend), CD45 (AF700, eBioScience), CD34

(PE-Cy5, BioLegend), CD309 (PE-Cy7, BioLegend). Zombie UV viability dye (BioLegend) was used to label dead cells.

Imaging.

Brightfield images were obtained using an Olympus IX70 microscope. Fluorescent images were obtained using a Zeiss AxioPlan 2 microscope. qRT-PCR.

RNA was isolated using GenCatch Total RNA Miniprep Kit (Epoch) according to manufacturer protocol and was reverse transcribed using the High Capacity Reverse Transcription Kit

(Applied Biosystems) according to manufacturer protocol. mRNAs of specific genes were

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measured by qRT-PCR using TaqMan probes (Applied Biosystems) and the StepOnePlus Real-

Time PCR system (Applied Biosystems) as described [37-40]. The following Taqman probes were used for qRT-PCR: Nanog (Mm02019550_s1), Sox2 (Mm03053810_s1), Pecam1

(Mm01242576_m1), Foxf1 (Mm00487497_m1), Kit (Mm00445212_m1), Cd34

(Mm00519283_m1), and Cd309/Kdr (Mm00440088_g1). mRNAs were normalized to Gapdh

(Mm99999915_g1).

Statistical analysis.

Student’s t-test was used to determine statistical significance. P<0.05 was considered to be significant. Values for all measurements were expressed as mean ± standard error of mean.

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References

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Supplementary Information For Chapter 3

Supplemental Figure 3.1. ECFC analysis of tdTomato tracker. Supplemental Figure 3.2. PCR Screen for GFP:FOXF1 knock-in cell lines. Supplemental Figure 3.3. Sequence of Non-GFP allele of heterozygous GFP:FOXF1 clones identified in PCR screen compared to Wild Type (WT) Foxf1. Supplemental Figure 3.4. Flow cytometry gating strategy for EPC analysis.

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Supplemental Figure 3.1. ECFC analysis of tdTomato tracker. Flow cytometry was used to demonstrate tdTomato expression in the ECFCs utilized for transplantation.

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Supplemental Figure 3.2. PCR Screen for GFP:FOXF1 knock-in cell lines. (A) Schematic illustration of the PCR screening used in Fig. 3.6 B and Suppl Fig. 3.2 C. (B) Table showing primer sequences used in PCR screening for Fig. 3.6 B (3887, 3619, 3935, 4102) and Suppl. Fig. 3.2 C (listed on figure). (C) The clones identified in Fig. 3.6 were further analyzed to confirm homozygosity or heterozygosity.

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WT GCATCCCTCGGTATCACTCACAGTCGCCCAGCATGTGTGACAGAAAGGAGTTTGTCTTCT C2 GCATCCCTCGGTATCACTCACAGTCGCCCAGCATGTGTGACAGAAAGGAGTTTGTCTTCT E4 GCATCCCTCGGTATCACTCACAGTCGCCCAGCATGTGTGACAGAAAGGAGTTTGTCTTCT G2 GCATCCCTCGGTATCACTCACAGTCGCCCAGCATGTGTGACAGAAAGGAGTTTGTCTTCT ************************************************************

WT CTTTCAATGCCATGGCCTCTTCTTCTATGCATACAACAGGCGGAGGATCTTACTATCACC C2 CTTTCAATGCCATGGCCTCTTCTTCTATGCATACAACAGGCGGAGGATCTTACTATCACC E4 CTTTCAATGCCATGGCCTCTTCTTCTATGCATACAACAGGCGGAGGATCTTACTATCACC G2 CTTTCAATGCCATGGCCTCTTCTTCTATGCATACAACAGGCGGAGGATCTTACTATCACC ************************************************************

WT AGCAGGTCACCTACCAAGACATCAAGCCGTGTG-TGATGTGAGGTGAGGCCACGGGGCCC C2 AGCAGGTCACCTACCAAGACATCAAGCCGTGTGATGATGTGAGGTGAGGCCACGGGGCCC E4 AGCAGGTCACCTACCAAGACATCAAGCG------TGATGTGAGGTGAGGCCACGGGGCCC G2 AGCAGGTCACCTACCAAGACATCAAGCCGT------***************************

WT TCCAGCCCAGCCTGGCCGGCCCAGGGACCAGGAGCCCACCGCCACAAACTGCTTTACTCT C2 TCCAGCCCAGCCTGGCCGGCCCAGGGACCAGGAGCCCACCGCCACAAACTGCTTTACTCT E4 TCCAGCCCAGCCTGGCCGGCCCAGGGACCAGGAGCCCACCGCCACAAACTGCTTTACTCT G2 ------GAGCCCACCGCCACAAACTGCTTTACTCT *****************************

WT GGAGGTATAACCCGTCAGCAAGTGAAAAGGGATAGCCCCACCCCTAACGGATTATTTGTA C2 GGAGGTATAACCCGTCAGCAAGTGAAAAGGGATAGCCCCACCCCTAACGGATTATTTGTA E4 GGAGGTATAACCCGTCAGCAAGTGAAAAGGGATAGCCCCACCCCTAACGGATTATTTGTA G2 GGAGGTATAACCCGTCAGCAAGTGAAAAGGGATAGCCCCACCCCTAACGGATTATTTGTA ************************************************************

WT AAGAAAATCCCAACACAGACTGGGAGCAGCGTCTCTACCCTCACTCCCTCA C2 AAGAAAATCCCAACACAGACTGGGAGCAGCGTCTCTACCCTCACTCCCTCA E4 AAGAAAATCCCAACACAGACTGGGAGCAGCGTCTCTACCCTCACTCCCTCA G2 AAGAAAATCCCAACACAGACTGGGAGCAGCGTCTCTACCCTCACTCCCTCA ***************************************************

Supplemental Figure 3.3. Sequence of Non-GFP allele of heterozygous GFP:FOXF1 clones identified in PCR screen compared to Wild Type (WT) Foxf1. Nucleotides identical to the WT sequence are indicated with *. Sequence homology was generated by the Clustal Omega online resource provided by European Bioinformatics Institute, . Clone C2 has a 1 base pair (BP) insertion. Clone E4 has a 5 bp deletion. Clone G2 has a 60 bp deletion.

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Supplemental Figure 3.4. Flow cytometry gating strategy for EPC analysis. Flow cytometry diagram showing gating strategy for parental (W4) line. Single cells were isolated from debris and doublets then sorted for live cells. The final panel is the same live/dead stain as shown in Fig. 3.10 B.

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Chapter 4: Discussion and Future Directions

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Liver

Hepatic injury can result from a variety of infectious, toxic, and metabolic agents, and can lead to hepatic fibrosis [1]. Hepatic fibrosis is characterized by an excessive deposition of extracellular matrix (ECM) and collagen which disrupts the liver architecture and impairs liver function [2]. The fibrous lesions of hepatic fibrosis are produced by myofibroblasts, which differentiate from hepatic stellate cells (HSC), and are initially a protective response to shield the liver from further damage [2, 3] (Fig. 4.1 A). The Forkhead Box F1 (FOXF1) transcription factor is expressed in HSCs and is essential for liver repair after acute liver injury [4] (Fig. 4.1 B).

FOXF1 is additionally known to stimulate HSC activation in response to liver injury [4, 5] (Fig.

4.1 C).

To determine the role of FOXF1 in hepatic fibrosis, we generated αSMACreER;Foxf1fl/fl mice to conditionally inactivate Foxf1 in myofibroblasts during carbon tetrachloride-mediated chronic liver injury. Deletion of Foxf1 in MFs exacerbated hepatic fibrosis, increased collagen deposition, disrupted liver architecture, and stimulated expression of profibrotic genes Col1α2,

Col5α2, and Mmp2 in the liver tissue (Fig. 4.1 D). Our studies indicate that Foxf1 expression in

MFs is critical to prevent aberrant MF accumulation and ECM deposition during hepatic fibrosis by repressing pro-fibrotic gene transcription in myofibroblasts and HSCs. Further work can determine if FOXF1 directly binds to the profibrotic genes found and can further tease out the transcriptional networks of the myofibroblasts during and after hepatic fibrosis. In addition, the role of FOXF1 in the reversion of hepatic fibrosis remains unknown.

Since FOXF1 has important roles in HSC activation, liver repair after injury, and regulation of collagen deposition by MFs during the progression of fibrosis, it is possible and

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Figure 4.1. FOXF1 activity during the progression of liver injury. Schematic illustrations demonstrates (A) the normal wound healing response to hepatic injury. (B) FOXF1 is expressed in both quiescent hepatic stellate cells and activated myofibroblasts (MFs) during liver injury. (C) FOXF1 expression is necessary for proper activation of MFs to protect the liver from damage. (D) Loss of FOXF1 after HSC activation to MFs results in MF accumulation and excessive collagen deposition. (E) The role of FOXF1 in the resolution of hepatic fibrosis remains unknown but could aid in MF apoptosis, senescence, or MF reversion to an HSC-like cell.

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likely that FOXF1 plays a role in fibrosis reversion (Fig. 4.1 E). Upon removal of the underlying etiological agent of fibrosis, MFs disappear and fibrosis subsides. Recovery from fibrosis is associated with remodeling of the excess matrix and restoration of normal hepatic architecture.

There are currently three main theories in regards to MFs during fibrosis resolution: Reversion

[6, 7], senescence [8], and apoptosis [9].

MF reversion has long remained a theory, however, recent publications have independently revealed evidence for the reversion of MFs into an HSC-like state, primed to return to the MF phenotype [6, 7]. These reverted HSCs have an increased responsiveness to recurring fibrogenic stimulation [6, 7] with a stronger contribution to subsequent fibrosis [6].

Since FOXF1 is expressed in MFs during the normal progression of hepatic fibrosis and is responsible for regulating collagen deposition, it would stand to reason that FOXF1 may additionally be responsible for reversion to a HSC-like cell.

Cellular senescence is a stable form of cell cycle arrest that can be triggered in a myriad of cell types in response to cell damage or stress [10]. In hepatic fibrosis, (methods) were used to determine that MFs senesce and limited fibrotic response [8]. Previous studies have shown that p53 contributes to cellular senescence [11, 12]. When a recent study described FOXF1 as a target of the p53 family [13], it became reasonable that the role FOXF1 plays in fibrotic reversion is to trigger cellular senescence.

Apoptosis is the favored theory for MF clearance during hepatic fibrosis resolution [7].

Apoptosis is the programmed death of a cell and occurs in MFs after cessation of liver injury to contribute to recovery of normal hepatic architecture. Previous studies have shown conflicting results regarding FOXF1 and apoptosis. Endothelial FOXF1 deletion in mouse embryos revealed that apoptosis is increased which lead to defects in vascular development [14], whereas lung

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endothelial deletion of FOXF1 in adult mice did not induce apoptosis in an acute lung injury model [15]. More work will need to be done to determine if FOXF1 expression could induce apoptosis. Since loss of FOXF1 either induced fibrosis or had no effect on the mechanism [14,

15], it is unlikely that FOXF1 plays a role in fibrotic reversion by triggering apoptosis.

Each of these theories of MF clearance could be tested with lineage-tracing experiments.

A conditional reporter gene could be included to trace desmin+ (Des) cells. Carbon tetrachloride could be used to induce hepatic fibrosis, which would in turn activate HSCs to their MF-like state. After chronic liver injury, fibrosis would be allowed to subside for 2 weeks before mice were harvested and liver tissue collected. 14 days were chosen based on a wound-healing study which demonstrated a high number of fibroblasts between 6 and 15 days after injury with that number subsiding by day 30 [16]. Liver sections could be stained to test each theory: For reversion: cells would express Des but not αSMA. For senescence: cells would either express p21 or p27 [17] but not Ki-67 or PH3. For apoptosis: cells would express cleaved caspase 3 but not Ki-67 or PH3. The results of this experiment would round out our knowledge of FOXF1 in hepatic fibrosis from HSC activation, MF regulation, and finally to the role of FOXF1 in hepatic injury resolution.

ESCs

Pulmonary endothelial cells regulates the passage of nutrients and fluids and are essential for endothelial barrier maintenance and lung homeostasis. Our lab recently demonstrated that the transcription factor FOXF1 promotes normal lung homeostasis by regulating endothelial barrier function [15]. Acute lung injury (ALI) and the more severe acute respiratory distress syndrome

(ARDS) both increase endothelial permeability, which causes pulmonary edema and

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inflammation. Currently, novel treatment approaches to increase patient survival for ALI and

ARDS remain elusive. Other severe lungs disorders such as chronic obstructive pulmonary disease, an inflammatory lung disorder which results in obstructed air flow from the lungs [18], and pulmonary arterial hypertension, a disease characterized by microvascular loss resulting in an increase in blood pressure [19], additionally have tenuous treatment options with lung transplant being the most promising [20, 21], even with its low 5-year survival rate [22]. Recent advances in treatment of human lung diseases have introduced cell replacement strategies to contribute to lung regeneration (outlined in Chapter 3).

To improve endothelial barrier function of mice with ALI, we introduced endothelial colony forming cells (ECFCs) into the bloodstream of injured mice and increased survival of

53.8% of injured mice, which was associated with a marked decrease in lung injury (Chapter 3

Part A). Since ECFC integration was not observed due to tools used, we generated a novel embryonic stem cell line where we have knocked GFP in to the Foxf1 locus (GFP:FOXF1, A1).

Since the original ECFC were not positive for FOXF1, we developed a novel differentiation protocol which yielded 95.2% endothelial cells with a sub-population of endothelial progenitor cells that were 51.5% FOXF1-positive (Chapter 3 Part B). These newly-developed tools can be utilized in future studies regarding endothelial barrier function and lung regeneration and to determine the precise mechanism by which the introduction of ECFCs functioned to increase mouse survival and prevent exacerbated lung injury.

Despite the extensive research being conducted to differentiate into and understand EPCs, each published article reveals a novel genetic signature (reviewed in Chapter 3 Part B). Since no standard genetic signature exists for early and late EPC identification, studies cannot be properly compared to establish consistent protocols for differentiation. While each study reviewed in

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Chapter 3 Part B, including the study conducted in Chapter 3 Part B, listed markers associated with ECs, no two studies evaluated the same markers while claiming to have produced early [23]

(Chapter 3 Part B) or late [23-26] EPCs. Elucidating these early and late EPC gene signatures will be important when future studies differentiate EPCs with the purpose of utilizing the EPCs as potential therapeutic treatments for patients with lung diseases. All of this being said, EPC differentiation is a relatively new field and gene signatures will be revealed which stand the tests of time and repeated experiments.

Since the EPCs differentiated in Chapter 3 Part B were CD31+CD45-KIT+, we have classified them as early EPCs; however, we need to further characterize these EPCs to ensure they have EC properties and not just the genetic signature. Since ECs are known for their ability to home to injury sites in vivo, we can perform a scratch assay which will evaluate the ability of our EPCs to migrate [27] (Fig. 4.2 A). In addition, ECs are able to perform angiogenesis. To test this ability, we can perform in vitro tube-forming assays with our EPCs [28] (Fig. 4.2 B).

Finally, we can perform a morphological analysis of our EPCs to ensure they resemble the cobblestone-like morphology of cultured vascular EC [29] (Fig. 4.2 C).

Overall, more work needs to be done to drive research towards translating pre-clinical

ESC and EPC transplantation studies to clinical trials. In order for EPC populations to constitute effective clinical treatments, they will need to exhibit pro-survival, self-renewal, engraftment properties, and increased angiogenesis. For now, proof-of-concept studies can utilize the mouse

EPCs derived in Chapter 3 Part B to rescue murine models of human diseases. Since we know

ECFCs can increase mouse survival (Chapter 3 Part A). We can inject our GFP:FOXF1+ EPCs into the same ALI model to see if we can determine how the ECFC were originally able to improve survival. We can test if the GFP:FOXF1+ EPCs transiently engraft for a few days then

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Figure 4.2. Proposed studies to test GFP:FOXF1+ EPCs. (A) Scratch assay will evaluate cell migratory ability. (B) Tube-forming assay will test angiogenic capacity of EPCs. (C) Cultured EPCs should exhibit cobblestone-like morphology. (D) GFP:FOXF1+ EPCs could be transplanted into disease models of acute lung injury (ALI), bronchopulmonary dysplasia (BPD), and alveolar capillary dysplasia with a misalignment of pulmonary veins (ACD/MPV) to see if these cells can improve mouse survival, rescue disease phenotype, and to determine if the GFP:FOXF1+ EPCs can integrate into the lung vasculature.

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shed, engraft full and integrate into the endogenous lung barrier system, or neither. Since human studies have shown engraftment of stem cells that underwent in vivo differentiation into ECs

[30], it is likely that EPC engraftment will occur in these mice with ALI (Fig. 4.2 D).

Since ALI is not the only lung disease in need of better therapeutic treatments, we could use our GFP:FOXF1+ EPCs to rescue mice with either a bronchopulmonary dysplasia (BPD) phenotype or the lethal perinatal lung disorder, alveolar capillary dysplasia with a misalignment of pulmonary veins (ACD/MPV) (Fig. 4.2 D). For the former, hyperoxic treatment is known to result in lung simplification, mimicking a BPD phenotype [31]. For the latter, we could utilize our recently published murine model of ACD/MPV with a S52-mutation in Foxf1 [32]. In both of these studies, mice could be injected with our GFP:FOXF1+ EPCs and evaluated for survival, disease recovery, and EPC integration.

In summary, stem cells are a useful tool for studying both differentiation and disease mechanisms in culture. Although FOXF1 has been extensively described in ECs in vivo

(reviewed in Chapter 1), our novel GFP:FOXF1 ESC line can help elucidate the role of FOXF1 in EC differentiation in EPCs. Doing so will additionally aid in identification of progenitors that could repopulate and restore normal airways of a multitude of lung diseases.

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