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UNVEILING THE BIOLOGY OF COLLECTING DUCT EPITHELIUM REPAIR AND REGENERATION

Pamela Kairath Oliva

Department of Paediatrics Faculty of Medicine, Dentistry, and Health Sciences The University of Melbourne Australia

Submitted in total fulfilment of the requirement of the degree of Doctor of Philosophy

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ABSTRACT

The healthy functioning of the kidney requires the orchestrated action of its two functional units, nephrons and the collecting duct system (CD). Therefore malfunction of either of these two essential compartments can lead to kidney failure. Diseases affecting the collecting duct system (CD), congenital abnormalities of the kidney and urinary tract (CAKUTs), are the most frequent cause of End-Stage Renal Disease (ESRD) in children. At this stage, the only available therapeutic options for treating kidney failure are dialysis or organ transplantation. Given the fact that only one in three patients will receive a transplant, and that dialysis comes with a high risk of mortality, the study of the mechanisms underlying the repair and regeneration of the collecting duct system is vital because it will facilitate new therapeutic strategies for treating kidney disease which are available to more people.

The CD system originates from one of the two progenitor populations which give rise to the kidney, the ureteric bud (UB). The formation of the UB is a substantial part of the nephrogenesis process, and develops into a branched tree-like structure which will ultimately form the ducts of the urinary collecting system. This PhD thesis investigates recent concepts in normal kidney organogenesis, repair, and regeneration, and focuses on the CD system.

In the first part of this research, we investigated the capacity of an endogenous kidney mesenchymal stem cell population (k-MSC), previously reported by our laboratory, which both arose from the collecting duct epithelium and also then integrated into the same compartment. To this end, we generated and examined the functional capacity of Pkd1 defective k-MSCs to trigger autosomal dominant polycystic kidney disease (ADPKD) into wild type mice. Given that micro-injection of double transgenic k-MSCsPkd1del2-4/TMTO+ did not produce significant evidence for cyst formation in the recipient mice, we therefore ruled out the possibility that k- MSCs represent a preferable population for effecting repair. Nevertheless, we showed the capacity to generate k-MSCs with mutant genes, which may prove useful as cellular models of human diseases.

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We then addressed the ambitious aim of recreating the ureteric bud population via the directed differentiation of human pluripotent stem cells (hiPSCs). Here we report a stepwise protocol for the differentiation of hiPSCs towards nephric duct (ND)/ureteric bud (UB) lineage. Optimisation cultures were established using fluorescently-tagged human iPSC lines as a readout of collecting duct differentiation cultures, which also provided a unique opportunity to trace cell lineage within organoids. Cells subjected to this differentiation protocol switched the expression of key genes thought to be of ND/UB lineage, and also showed the ability to epithelialise; this was seen using two- and three-dimensional approaches. The establishment of this methodology provides a valuable platform to now investigate the capacity of this epithelium to respond to a metanephric mesenchyme. We also anticipate that the use of this protocol will ultimately facilitate three-dimensional bio-printing and allow the recreation of larger kidney structures.

This thesis has investigated areas currently applicable to regenerative medicine, including normal tissue development, turnover, repair, and regeneration, with a focus on the collecting duct epithelium. This research has concluded that k-MSCs do not represent a preferable population for effecting structural repair. It has also established the basis of a new methodology to recreate kidney tissue for therapeutic uses.

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DECLARATION

I. The thesis comprises only original work towards the Doctor of Philosophy, II. Due acknowledgement has been made in the text of all other material used, III. The thesis is less than 100,000 words in length, exclusive of tables, figures, bibliographies and appendices.

Pamela Kairath Oliva

Signature: ______December 2017

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PREFACE

Pursuant to the regulations governing to the degree of Doctor of Philosophy at the University of Melbourne, I hereby submit that:

Declaration by author: This thesis is composed of my original work, and contains no material previously published or written by another person except where due reference has been made in the text. I have clearly stated the contribution by others to jointly-authored works that I have included in my thesis.

I have clearly stated the contribution of others to my thesis as a whole, including statistical assistance, survey design, data analysis, significant technical procedures, professional editorial advice, and any other original research work used or reported in my thesis. The content of my thesis is the result of work I have carried out since the commencement of my research higher degree candidature and does not include a substantial part of work that has been submitted to qualify for the award of any other degree or diploma in any university or other tertiary institution. I have clearly stated which parts of my thesis, if any, have been submitted to qualify for another award.

Statement of parts of the thesis submitted to qualify for the award of another degree: None

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Publication during the candidature:

1. Little MH, Kairath P. Regenerative medicine in kidney disease. Kidney Int. 2016 Aug; 90(2):289-99. doi: 10.1016/j.kint.2016.03.030. Epub 2016 May 24. Review

(Attached in appendix 1)

2. Little MH, Kairath P. Does renal repair recapitulate kidney development? J Am Soc Nephrol. 2017 Jan; 28(1):34-46. doi: 10.1681/ASN.2016070748. Epub 2016 Oct 26. Review

(Attached in appendix 2)

Patents:

1. Kairath P, Takasato M, Er X. P, Little MH. Directed differentiation of human pluripotent stem cells to Nephirc duct/Ureteric bud like-cells. Provisional submission in process. (Chapter 3 and 4).

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Contributions by others to the thesis:

Prof. Melissa Little contributed to concept and design of the projects as well as interpretation of experimental data. She also performed GO analysis of single cell RNA-seq and provided critical proofreading for the thesis manuscript. Moreover, she provided constant and valued supervision.

Dr. Joan Li contributed to design, analysis and interpretation of experimental data. She also performed microinjections and assistance in the analysis of k-MSCs (GFP+) (Chapter 2).

Dr. Jessica Vanslambrouck provided advice in techniques and supervision meetings.

Dr. Santhosh Kumar provided advice in techniques and supervision meetings. He also performed cell dissociation for RET/tdTomato+ organoid (Chapter 4).

Dr. Sara Howden contributed to the generation of 1502.2 GATA3:m-Cherry and 2429 GATA3:m-Cherry iPSC lines (Chapter 3 and 4).

Pei Er provided training and technical advice for the maintenance and banking iPSCs (Chapter 3 and 4).

Irene Ghobrial provided technical assistance for iPSCs banking (Chapter 3 and 4).

Luke Zapia performed analysis for single cell RNA-seq (Chapter 4).

Dr. Alexander Combes performed GO analysis for single cell RNA-seq (Chapter 4).

Virginia Nink conducted FACS sorting at University of Queensland (Chapter 2).

Dr. Matthew Burton and Paul Lai conducted FACS sorting at Murdoch Children’s Research Institute (Chapter 3 and 4).

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ACKNOWLEDGEMENTS

Firstly, I would like to thank my supervisor Prof. Melissa Little for her valuable mentoring, guidance, support, and time throughout my PhD journey. I am deeply grateful for her constant debate and criticism of the design of experiments and the analysis of the data, for the opportunities she gave me to assume new challenges, and for always encouraging me to do my best, which has not only sharpened my analytical skills but also helped me to discover my own research interests.

Thank you to the members of the Little Lab, both past and present, for your help and companionship. Special thanks to Dr. Minoru Takasato for giving me the first directions in developmental and stem cell biology. Also, thanks to Pei Er for training me up in stem cell culture, and for her friendship over the years. I would also like to thank the co-supervisors who helped me during different stages of my PhD- Dr. Joan Li, Dr. Jessica Vanslambrouck, and Dr. Santhosh Kumar- for insightful discussions and feedback during PhD meetings.

I would also like to thank the University of Queensland, where I did the first part of my PhD studies. Special mentions to Amanda Carozzi for her constant support, not only in the beginning but also during the transference of my candidature to University of Melbourne. I am also grateful to my committee members, Dr. Shireen Lamande and Dr. John Bateman, for their advice and guidance upon my arrival at the University of Melbourne.

Last but not least, I would like to thank my family for their long-distance support and unconditional love. My most sincere and profound gratitude goes to my husband, Rodrigo, not only for believing in my capabilities and accompanying me to venture overseas to take up this challenge, but also for the endless encouragement, care, and love throughout my PhD candidature which helped me to overcome any difficulties that I encountered along the way.

I was supported by a CONICYT Becas-Chile postgraduate scholarship (#72120118).

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TABLE OF CONTENTS ABSTRACT ...... 2 DECLARATION...... 4 PREFACE ...... 5 ACKNOWLEDGEMENTS ...... 8 FIGURES ...... 13 TABLES ...... 17 ABBREVIATIONS ...... 18 CHAPTER I ...... 20 1.1 INTRODUCTION ...... 21 1.1.1 The kidney: anatomy, cell types and their functional roles in the adult ...... 21 1.1.2 Chronic kidney disease (CKD) ...... 22 1.1.2.1 Malfunction of the collecting duct system ...... 23 1.1.2.2 Limited treatment options ...... 25 1.1.3 Kidney Development ...... 25 1.1.3.1 Fundamental embryonic decisions during kidney organogenesis ...... 26 1.1.3.1.1 Primitive streak determination ...... 26 1.1.3.1.2 Intermediate mesoderm determination ...... 27 1.1.3.2 Formation of three pairs of kidneys ...... 29 1.1.3.3 The metanephric kidney: nephrogenesis and branching morphogenesis ...... 30 1.1.3.3.1 Nephrogenesis ...... 31 1.1.3.3.2 Ureteric bud outgrowth and the branching morphogenesis process ...... 33 1.1.4 Kidney repair in response to injury ...... 35 1.1.4.1 Cellular approaches to improving repair ...... 36 1.1.5 Kidney Regeneration ...... 38 1.1.5.1 Induced Pluripotent Stem Cells (iPSCs): a rapidly evolving technology ...... 38 1.1.6 Significance and approaches of this thesis ...... 40 CHAPTER II ...... 41 2.1 INTRODUCTION ...... 42 2.1.1 The mesenchymal stem cell ...... 42 2.1.1.1 MSCs in the Context of Stem Cells ...... 42 2.1.1.2 General Properties of Bone Marrow MSCs (bm-MSCs) ...... 43 2.1.2 The delivery of bone marrow MSCs into kidney disease models ...... 45 2.1.2.1 Acute Kidney Injury (AKI) ...... 45 2.1.2.2 Chronic Kidney Disease ...... 46 2.1.3 The kidney mesenchymal stromal cell population (k-MSCs) ...... 47 2.1.3.1 Phenotypic Characterization of k-MSCs ...... 47

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2.1.3.2 New Insights and Tracing the Origin of k-MSCs ...... 48 2.1.4 The selected model of renal disease: Dominant polycystic kidney disease ...... 50 2.1.4.1 Autosomal Dominant Polycystic Kidney Disease and the Pkd1 Mouse Model ...... 50 2.1.5 Hypothesis and aims ...... 53 2.2 RESULTS ...... 54 2.2.1 Establishment of a double-transgenic mouse line ...... 57 2.2.2 Generation and phenotypic characterization of a primary k-MSC line deficient for the Pkd1 gen (k-MSCsPkd1del2-4) ...... 59 2.2.3 Generation of genetically-tagged Pkd1 mutant k-MSCs for in vitro and in vivo characterisation (k-MSCsPkd1del2-4/TMTO+) ...... 65 2.2.3 Microinjection experiments ...... 69 2.2.4 In vivo evaluation ...... 71 2.3 DISCUSSION ...... 76 2.3.1 Challenges with the project / reasons why integration may have failed ...... 77 2.3.1.1 Laboratory relocation ...... 78 2.3.1.2 Little evidence of integration after initial injections ...... 78 2.3.2 Future directions ...... 79 CHAPTER III ...... 80 3.1 INTRODUCTION ...... 81 3.1.1 The main regulatory signalling pathway of ureteric bud formation: GDNF-RET ...... 81 3.1.1.1 GDNF/GFRα1/RET complex and relevance of RET expression during UB outgrowth .. 81 3.1.1.2 Regulation of Ret and Gdnf expression ...... 83 3.1.2 The role of retinoids in modulating Ret expression in the developing kidney ...... 84 3.1.3 Intracellular signalling pathways and downstream regulators of RET ...... 87 3.1.4 Other signalling pathways regulating ureteric bud morphogenesis ...... 90 3.1.5 Generation of kidney cell types from induced pluripotent stem cells (iPSCs) ...... 91 3.1.6 Hypothesis and aims ...... 94 3.2 RESULTS ...... 95 3.2.1 GDNF and retinoic acid promote branching morphogenesis in kidney explants ...... 95 3.2.2. Evaluation of GDNF and All-trans retinoic acid (tRA) in kidney organoids ...... 98 3.2.3 Assessment of the iPSC line: 1502.3 GATA3: m-Cherry clone 60 ...... 103 3.2.4 Assessment of the fluorescently-tagged iPSC line: 2429 GATA3: m-Cherry clone 26 ...... 108 3.3 DISCUSSION ...... 116 CHAPTER IV...... 122 4.1 INTRODUCTION ...... 123 4.1.1 Anterior intermediate mesoderm and its derivatives ...... 123 4.1.1.2 Nephric duct lineage specification ...... 123

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4.1.2 Segmentation of the ureteric epithelium ...... 126 4.1.3 Collecting duct epithelial cell differentiation ...... 128 4.1.4 Hypothesis and aims ...... 129 4.2 RESULTS ...... 130 4.2.1 Assessment of the capacity of the protocol to trigger epithelia formation in two- and three- dimensional approaches...... 130 4.2.2 Evaluation of the molecular signature of Cherry+-expressing cells...... 139 4.2.1 Characterization of the response RET/ tdTomato line to the ND/UB protocol ...... 141 4.2.2 Dissecting cell composition in organoids derived from the iPSC RET/tdTomato+ line ...... 145 4.3 DISCUSSION ...... 148 CHAPTER V ...... 154 CONCLUDING REMARKS ...... 155 CHAPTER VI...... 159 6.1 MATERIALS AND METHODS ...... 160 6.1.1 Mice ...... 160 6.1.2 Neonatal Injection Model...... 160 6.1.3 Quantification of cell integration: ...... 160 6.1.4 Isolating and culturing the intact metanephric kidney ...... 161 6.1.5 Cell culture ...... 161 6.1.5.1 Cell Isolation from adult kidney tissue and maintenance of k-MSCs lines ...... 161 6.1.5.2 iPSC and culture on mouse embryonic feeders (MEFS)...... 162 6.1.5.3 Generation of iPSC reporter lines and culture on a feeder layer-free culture ...... 162 6.1.5.4 iPSC culture on a feeder layer-free culture ...... 164 6.1.5.5 Standard differentiation protocol of iPSC lines to form kidney organoids...... 164 6.1.6 Transfection and Antibiotic Selection ...... 165 6.1.7Clonal Selection and Expansion of k-MSCs Mixed Population ...... 165 6.1.8 Cilia induction ...... 165 6.1.9 MRI ...... 165 6.1.10 Cystic index ...... 166 6.1.11 Statistical analysis ...... 166 6.1.12 Imaging ...... 166 6.1.13 Immunofluorescence: ...... 166 6.1.14 Flow cytometry and cell sorting ...... 169 6.1.14.1 Fluorescent activated cell sorting (FACS) ...... 169 6.1.14.2 Flow cytometry ...... 170 6.1.15 RNA extraction, cDNA preparation, qRT-PCR and RT-PCR...... 170 6.1.16 Data generation and analysis for single-cell RNA sequencing ...... 172

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BIBLIOGRAPHY ...... 173 APPENDIX 1 ...... 200 APPENDIX 2 ...... 213

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FIGURES

CHAPTER I Figure 1.1. Anatomical segmentation of the adult mammalian kidney…………………………….....22 Figure 1.2. Fundamental embryonic decisions during kidney organogenesis…………………….….27 Figure 1.3. Formation of three pairs of kidneys during mammalian development…...…..…………..29 Figure 1.4. Schematic representation of kidney organogenesis…………………………………..…..30 Figure 1.5. Overview of cellular compartments and molecular interaction in the nephrogenic niche……………………………………………………………………………………..32 Figure 1.6. Different stages of the renal branching morphogenesis process throughout kidney development……………………………………………...…………………………….……..35 Figure 1.7. Schematic diagram depicting the approaches taken in this thesis………..…………..…..40 CHAPTER II Figure 2.1. Multipotent mesenchymal stromal cells in adult tissues……………………..………..…44 Figure 2.2. Characterization of endogenous kidney MSC (kCFU-F) compared to heart (cCFU-F) and bone marrow populations (bmMSC)…………………………………………..……....48 Figure 2.3. Specific epithelial integration of kidney MSCs after their microinjection into the nephrogenic zone………………………………………………………………………………….49 Figure 2.4. Cysts formation at the level of cell, nephron and kidney…………………………….…..51 Figure 2.5. The function of polycystin-1…………………………………………………....….…….52 Figure 2.6. Specific epithelial integration of k-MSCs (GFP+) is detectable within collecting duct compartments after 4, 6, and 10 weeks post-delivery…………………...……………….……...55 Figure 2.7. Quantification of the long-term integration capacity of k-MSCs (GFP+) into collecting duct structures via the calculation of the ratio GFP+Aqp2+/TotalAqp+2 ……………..…….56 Figure 2.8. Breeding strategy for the generation of the double transgenic mouse model: Pkd1Flox/ Flox;Rosa26TdTomatoFlox/Flox……………………………………………….…..…….58 Figure 2.9. Genotyping of the two key genes: Pkd1 and RosaTdTomato…………………….…...…59 Figure 2.10. Puromycin sensitivity assay for kidney MSC-like cells…………….………….....…….60 Figure 2.11. Experimental in vitro approach used to inactivate Pkd1Flox/Flox gene……………..……..61 Figure 2.12. Selection of stable clones of k-MSCsPkd1del2-4 from a mixed population……………..….62 Figure 2.13. Fibroblast-like morphology of clonally derived k-MSCs…………………………….…64 Figure 2.14. Comparative immunophenotype analysis of wild-type and mutated clones of k-MSCs………………………………………………………………………….…………………64 Figure 2.15. Validation of a double transgenic cell line k-MSCPkd1del2-4/TMTO+…………………..…...66 Figure 2.16. Evaluation of mesenchymal/epithelial protein markers in the double transgenic lines (k-MSCsPkd1 del2-4 and k-MSCsWt)………………………………………….…...…….67 Figure 2.17. k-MSCsPkd1del2-4/TMTO+ resulted in reduced cell number………………………...…...…..68

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Figure 2.18. Evaluation of cell integration of k-MSCsPkd1del2-4/TMTO+ after Microinjection experiments……………………………………………………………….…………..70 Figure 2.19. MRI scanning images of injected mice at 6 months of age…………………..…...…….72 Figure 2.20. Representative MRI scanning images at 12 months of age………...………………..….73 Figure 2.21. Bright-field and fluorescent images of the cystic kidney at 12 month of age…………..73 Figure 2.22. Evaluation of the feasibility of immunodetection of k-MSCsPkd1del2-4/TMTO+………….....74 Figure 2.23. Cyst formation at 12 months post-injection……………………………………..…...….75 CHAPTER III Figure 3.1. Illustration of Ret+cell movements from nephric duct to UB formation…...... …...83 Figure 3.2. Retinoic acid signalling pathway………………………………...…………….....………85 Figure 3.3. Retinoids (cis and trans) are sufficient to maintain Ret in isolated ureteric buds…....…..86 Figure 3.4. Gene networks involved in nephric duct (ND) and ureteric bud (UB) morphogenesis…………………………………………………………………………….…………..89 Figure 3.5. Directed differentiation protocol for the generation of kidney organoids from hPSCs…………………………………………………………………………………………....93 Figure 3.6. Titration of GDNF in kidney explants from E.11.5 HOXB7-GFP mice………………...96 Figure 3.7. AGN193109 ceases arborisation in kidney explants from E.11.5 HOXB7-GFP mice……………………………………………………………………………..……..97 Figure 3.8. Schematic representation of the first differentiation protocol to induce iPSCs to ND/UB tip+ lineage………………………………………………………………….99 Figure 3.9. Bright-field images of differentiation process during the first seven days of the protocol……………………………………………………………………………………99 Figure 3.10. FGF9 at the time of aggregation is critical for survival and structure formation within organoids………………………………………………………………………….100 Figure 3.11. Studying the effect of the presence and absence of GDNF from the aggregation day onward……………………………………………….…………….…………...….101 Figure 3.12. Addition of GDNF to differentiation cultures at the time of aggregation did not increase expression of RET receptor……………………………………………………...…102 Figure 3.13. Sequential strategy for the generation of a fluorescently-tagged cell line (1502.3 GATA3: m-Cherry Clone 60)……………………………………………………………….104 Figure 3.14. Evaluation of different rock inhibitors on differentiation cultures…………...………..105 Figure 3.15. Bright-field and confocal images of the developing organoids derived from the iPSC 1502.3 GATA3: m-Cherry clone 60 line across a time series……………….……....106 Figure 3.16. 24 hours incubation with Y27632 promotes better epithelium organization...... 107 Figure 3.17. Evaluation of the clonality of the iPSC cell line 1502.2 GATA3: m-Cherry clone 60…………………………………………………………………………107 Figure 3.18. Simultaneous strategy for the generation of a fluorescently-tagged

Commercial in Confidence cell line (2429 GATA3: m-Cherry clone 26)………………………………………………………...108 Figure 3.19. Validation of the iPSC 2429 GATA3: m-Cherry clone 26 line………………………..109 Figure 3.20. Experimental technique used to differentiate iPSCs towards an anterior intermediate mesoderm with UB signature………………………………………………………….111 Figure 3.21. Evaluation of different components and growth factors to promote the differentiation of iPSCs toward nephric duct/UB lineage……………………………….…………..112 Figure 3.22. Experimental technique used to differentiate iPSCs toward nephric duct/ureteric bud lineage……………………………………………………………….…………….113 Figure 3.23. Investigation of different growth factors to promote the differentiation of iPSCs towards nephric duct/UB lineage…………………………………………...…..…………114 Figure 3.24. Directed differentiation protocol for commitment of iPSCs to nephric duct/UB lineage………………………………………………………………………………………115 CHAPTER IV Figure 4.1. Stepwise formation of the nephric duct and renal collecting duct……..…………..……125 Figure 4.2. Ureteric bud segmentation and cell types………………………………..…………...…126 Figure 4.3. Bright-field images of differentiation of GATA3/m-Cherry+ reporter line to kidney organoids across the ND/UB protocol………………………………………..………131 Figure 4.4. Kidney organoids generated by ND/UB protocol purely contain collecting duct structures……………………………………………………………………….…………...….132 Figure 4.5. Immunofluorscence of GATA3/m-Cherry+ iPSCs-derived organoids…………..……..133 Figure 4.6. Extended two-dimensional culture of GATA3/m-Cherry+ iPSCs following UB protocol……………………………………………………………..………...………134 Figure 4.7. Live expression of m-Cherry+/expressing cells………………………………..……….135 Figure 4.8. Protein assessment in structures formed in two-dimensional extended cultures with UB protocol………………………………………………………………………..…..135 Figure 4.9. Extended two-dimensional culture of GATA3/mCherry+ iPSCs following standard protocol (ST)……………………………………………...…………………….………….136 Figure 4.10. Evaluation of the capacity of the standard protocol in generating UB/ collecting duct structures…………………………………………………...………………..….137 Figure 4.11. Absence of other kidney components in organoids and 2D-structures generated by the UB protocol……………………………………………………………………….138 Figure 4.12. Enriched expression of key genes of nephric duct/ureteric lineage…………….……..140 Figure 4.13. Schematic representation and differentiation process of the RET/tdTOMATO (GCamp6) iPSCs reporter line…………………………………..……………….141 Figure 4.14. Live expression of the tdTomato+-expressing cells……………………..…………….142 Figure 4.15. Bright-field and confocal images of the developing RET/tdTOMATO+ organoids across a time series………………………………………………………………………..143

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Figure 4.16. Gene expression analysis of RET/tdTomato+ expressing cells…………………...…...144 Figure 4.17. Dissecting human RET/tdTomato+ organoids using single-cell RNA sequencing………………………………………………………………………………...………….146 CHAPTER VI Figure 6.1. Reprogramming and gene targeting of human fibroblasts to generate reporter lines…………………………………………………………………………………………………..163

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TABLES

Table 1. Top 2500 differentially expressed (DE) collecting duct genes…………………………….147 Table 2. Top 2500 differentially expressed (DE) ureteric tip genes………………………………...147 Table 3. List of antibodies used for immunostaining………………………………………………..168 Table 4. List of primers used for qRT-PCR in this thesis…...……………………………………....171 Table 5. List of primers used for genotyping by RT-PCR………………………………………..…172

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ABBREVIATIONS

MSCs Mesenchymal stem cells bm-MSCs Bone marrow derived-MSCs k-MSCs Kidney derived-MSCs k-MSCs (GFP+) Kidney derived-MSCs tagged with GFP+ ADPKD Autosomal dominant kidney disease PC1 Polycystin-1 PC2 Polycystin-2 k-MSCsPkd1del 2-4 Primary cell line of k-MSC mutant for both Pkd1 alleles k-MSCsWt Primary cell line of k-MSC wild-type for both Pkd1 alleles k-MSCsPkd1del2-4/TMTO+ Primary cell line of k-MSC mutant for both Pkd1 alleles and fluorescently tagged tdTomato PND1 Post-natal day 1 MRI Magnetic resonance imaging ND Nephric duct WD Wolffian duct UB Ureteric bud NPs Nephron progenitors CD Collecting duct system AKI Acute kidney injury CKD Chronic kidney disease CAKUT Congenital abnormalities of the kidney and urinary tract ESRD End-stage renal disease PS Primitive streak A-P Anteroposterior axis IM Intermediate mesoderm AIM Anterior intermediate mesoderm PIM Posterior intermediate mesoderm ESCs Embryonic stem cells mESCs Mouse embryonic stem cells iPSCs Induced pluripotent stem cells BMP Bone morphogenetic protein E Embryonic day FGF Fibroblast growth factor GDNF Gial cell-derived neurotrophic factor

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RET Receptor tyrosine kinase TGFβ Transforming gowth factor bete tRA All-trans retinoic acid 9-cisRA 9-cis-retinoic acid HA Heparin DE genes Differential gene expression RNA Ribonucleic acid mRNA Messenger ribonucleic acid qPCR Quantitative real time polymerase chain reaction PBTX Triton X-100 diluted in PBS PFA Paraformaldehyde PBS Phosphate-buffered saline MEFs Mouse embryonic fibroblast IF Immunofluorescence FACS Fluorescent activated cell sorting DAPI 4′,6-diamidino-2-phenylindole

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CHAPTER I

Literature review

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1.1 INTRODUCTION

1.1.1 The kidney: anatomy, cell types and their functional roles in the adult

The primary role of the mammalian kidney is to filter the blood through specialised structures called nephrons, thereby regulating fluid homeostasis and removing metabolic wastes from the body. While the mouse kidney contains around 15,000 nephrons (Merlet-Benichou et al., 1999), the human kidney contains and average of million nephrons (Bertram et al., 2011). Anatomically, the adult kidney is divided into two regions, the renal cortex and the medulla. The medulla is subsequently subdivided into the outer and inner medulla, which contain distinct segments of the nephrons and hence perform distinctive renal functions (Figure 1.1). Functionally, the kidney is composed of two fundamental epithelial units, the nephrons and the collecting duct system (Figure 1.1). The nephron is further subdivided into two main components: the glomerulus, where blood filtration occurs, and the renal tubular network, where the filtered blood is modified by solute reabsorption and secretion (Hebert et al., 2001). More precisely, this tubular network can be structurally and functionally divided into three segments: the proximal segment, where reabsorption of two-thirds of the filtered salt and water, as well as all the organic solutes; the intermediate segment, responsible for urine concentration; and the distal segment, where electrolyte interchange and acid-base balancing takes place. In addition, some of the critical renal functions, including the regulation of extracellular volume, osmolarity, and pH, occur in the final portion of the nephron referred to as the collecting duct system (CD) (Figure 1.1). This is formed from two distinctive cellular types, principal cells (PCs) and intercalated cells (ICs), and extends from the connecting segment to the renal papilla. While intercalated cells are more abundant in the cortical collecting ducts and participate in acid-base homeostasis, principal cells are more abundant in the medulla/papilla region where they regulate sodium and water homeostasis (Guo et al., 2014). All nephrons connect with the collecting duct system through which the urine will leave the organ and move to the bladder. Hence, the healthy functioning of the collecting duct system is essential to maintain the fluid/electrolyte balance in the body. Unfortunately, developmental disorders arising due to genetic mutation or structural malformation can affect these essential renal structures leading to impaired function.

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Figure 1.1. Anatomical segmentation of the adult mammalian kidney. Each kidney is formed by an outer layer, and spatially organized into a peripheral layer (the cortex) and an inner layer (the medulla). The medulla is organized into several pyramidal structures. The tip of each pyramidal structure is called a renal papilla, and from here the urine drains into calices that subsequently sub-divide into outer and inner and converge into the renal pelvis. Nephrons are distributed within the cortex and medulla. The glomerulus is followed by the proximal tubule, which is linked to the connecting tubule and subsequently to the collecting duct system (see colour key). (Figure extracted from Davison et al., 2009)

1.1.2 Chronic kidney disease (CKD)

Kidney disease is one of the fastest growing chronic diseases in the world. In fact, one in ten of the adult population worldwide have evidence of kidney disease, according to the first global report on care delivery for kidney disease which was presented this year at the World Congress of Nephrology in Mexico (http://www.theisn.org/gobal-atlas). CKD is characterized by a complex aetiology, and is defined as the gradual loss of the ability of kidneys to perform its essential functions of removing waste and excess water and maintaining acid-base homeostasis. It normally has a gradual and asymptomatic onset, with increasing prevalence with older age, but it leads to high rates of premature mortality. Nevertheless, when CKD is diagnosed and treated in time the deterioration of the renal function can be reduced by up to 50% and may even be reversible (Johnson et al., 2004). The three major risk factors for CKD are diabetes, hypertension, and glomerulonephritis (Jha V., et al., 2013). People with CKD have a twenty times greater risk of dying from cardiovascular events (Keith et al., 2004). While diabetes and hypertension are the most common causes in both developed and developing countries, glomerulonephritis is the most frequent cause of CKD in Asia and sub-Saharan Africa. In

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1.1.2.1 Malfunction of the collecting duct system

Mutations or malfunctions in the collecting duct system are a result of various human diseases. The collecting duct system connects with ureters, the bladder, and the urethra to form the urinary tract, which removes wastes and fluids from the body. Congenital abnormalities of the kidney and urinary tract (CAKUT) comprise a group of structural malformations which occur at the level of the kidney (renal agenesis, hypoplasia, dysplasia, double kidneys), the collecting duct system (hydronephrosis and megaureter), the bladder (ureterocele and vesicoureteal reflux), or the urethra (posterior urethral valves) (Song and Yosypiv, 2011). CAKUT occur in 3-6/1000 of new-borns and frequently lead to renal insufficiency and End- Stage Renal Disease (ESRD) in children (7- Limwongse et al., 1999). Different mechanism can lead to CAKUT including mutations in a single or multiple genes (Song and Yosypiv, 2011), genetic (Sun et al., 2010, Manini et al., 2005, Fain et al., 2005, Hiesberger et al., 2005, Garcia-Gonzales et al., 2007, Kim et al., 2008) or epigenetic modifiers (Patel ., 2007), mode of inheritance and environment (Welham et al., 2005). Although few mutations of single or multiple genes have been described as causative of CAKUT, frequently in an autosomal dominant or more rarely in a recessive pattern of inheritance (Vivante et al., 2014), in many instances the molecular basis remains unresolved. Mutations in either PAX2 or HNF1B have been widely reported as causative of CAKUTs, and are related with ~ 15 % cases of CAKUTs (Weber et al., 2014, Thomas et al., 2011, Madariaga et al., 2013). While genetic aberrations in PAX2 are typically associated with renal dysplasia or hypoplasia, mutations in HNF1 are frequently associated with cystic kidneys (Weber et al., 2014, Thomas et al., 2011, Madariaga et al., 2013). In the same way, genetic aberrations in RET or GDNF, both essential genes in promoting the initial phases of development of the collecting duct system, also lead to this renal condition (Pini et al., 2009). In fact, the ‘budding hypothesis’ arose as one of the first explanations which linked ectopia of the ureteric bud with aberrant insertion of the ureter into the bladder and hypoplasia or dysplasia of the kidneys (Mackie et al., 1975).

Nephrogenic diabetes insipidus (NDI) is another renal condition caused by malfunction of the collecting duct system. Mutations in either Avpr2 (vasopressin type 2 receptor) or Aqp2 have a

Commercial in Confidence genetic prevalence of one per 25,000-30,000 (Robertson et al., 1995; Ananthakrishnan et al., 2009; Heinke et al., 2012) and cause an inability to concentrate urine. The most common symptoms of patients with NDI are polyuria, polydipsia, and an inability to concentrate urine (Robben et al., 2006). Approximately 90% of water in the urinary filtrate is reabsorbed through the water channel aquaporin-1 (AQP1), located in the proximal tubule and the descending limb of the Loop of Henle. The reabsorption of the remaining water is regulated by aquaporin-2 (AQP2), located in the renal collecting duct principal cells. This is controlled by levels of the antidiuretic hormone, arginine vasopressin (AVP), which determines the final urine concentration (Knepper et al., 1997). The interplay between AVP and AQP2 can be disrupted by mutations in both AQP2 (autosomal inheritance) and AVPR2 genes (X-linked inheritance), resulting in NDI. At present, 51 mutations in the AQP2 gene have been reported (Frenton et al., 2013) with these mutations impairing two key processes: 1) the subcellular sorting, with mutations leading to trafficking problems for the protein trying to reach the apical membrane; and 2) the formation of pore-forming structures. Both result in a disruption to water-channel function.

Many of these defects results in hydronephrosis of a swelling of the kidney due to a build-up of urine. Ultrasound, x-rays, computerized tomography, magnetic resonance imagining (MRI), and blood and urine tests are used to confirm the diagnosis. Depending on the severity of the damage, the excess of urine can be drained by catheter. However, if damage progresses, severe urinary blockage can irreversibly damage the kidneys and lead to kidney failure. Primary hydronephrosis can result from abnormalities in the junctions between the kidney, ureter, and bladder, which leads to urinary filtrate backing up into the pelvis. Although some clinical cases are related to physical obstruction of the urinary tract, others are caused by an inherited condition leading to failure of urinary filtrate concentration. For example, a transgenic mice model reported that a simple aminoacidic substitution of a Ser (S256) by a Leu in Aqp2 protein led to its accumulation in the apical membrane of collecting duct cells, resulting in a severe urine concentration defect (Bradley et al., 2005).

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1.1.2.2 Limited treatment options

The prevalence of CKD is increasing at 6.3% per annum (ESRD patients, 2013), representing a significant economic burden (in 2009, USA: $ 42 billion; Australia: $ 1 billion) (Cas., 2006). It is estimated that more than 80% of patients receiving treatment are from high-income countries (White et al., 2008). In poor countries few patients have access to renal replacement therapy (Jha et al., 2009). Hence there is an urgent need to find new therapeutic options which can be accessible to a greater number of patients.

Although the adult kidney has an ability to repair damaged epithelial structures throughout adult-life (Humphreys et al., 2008), this reparative ability reaches a barrier when facing sustained or chronic damage. Indeed, in humans the process of nephrogenesis (formation of new nephrons) is limited to the fetal development. Therefore, the irreversible loss of nephrons provoked by a sustained insult during adult-life initially compromises renal function and ultimately leads to End-Stage Renal Disease (ESRD). At this stage, the only available therapeutic options for treating kidney failure are dialysis or organ transplantation. Only one in three patients will receive a transplant (2016, ANZDATA Registry 39th annual report, available at http://anzdata.org.au), dialysis comes with a high risk of mortality, and both treatments are very expensive. For these reasons there has been a growing interest in the generation of alternative therapies for renal disease. During the last ten years, enormous progress has been made in the generation of pluripotent stem cells (iPSCs), the isolation of tissue-specific stem cells, and the reprogramming of adult cells to stem cells. These developments hold great promise for the recreation of human kidney tissue to treat renal failure. The continuous advance of such technologies relies upon fundamental biological insights into the cellular and molecular basis of normal kidney development, postnatal repair, and regeneration.

1.1.3 Kidney Development

The mammalian kidney is mesodermal in origin and represents one of only a small number of organs that arises from the intermediate mesoderm. The mesoderm in turn is one of the three primary germ layers of the early embryo and, like the endoderm, forms as a result of gastrulation. This section will describe the embryological events that occur from the point of gastrulation, seen as the formation of the primitive streak, to the formation of the permanent mammalian kidney, the metanephros.

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1.1.3.1 Fundamental embryonic decisions during kidney organogenesis

1.1.3.1.1 Primitive streak determination

Organ formation is the result of a series of key decisions during early embryogenesis. In the case of the kidney, this is demonstrated in the primitive streak-derived intermediate mesoderm. Upon epiblast formation, early stages of embryogenesis are dominated by cellular movements which give rise to an elongated group of cells along the axis of the embryo termed the ‘primitive streak’ (PS) at the caudal edge of the embryo (Figure 1.2). This marks the starting point of gastrulation, characterized by formation of the three germ layers- ectoderm, mesoderm, and endoderm- which will generate all the organs of the body. Specification and regionalisation of the primitive streak has been mostly documented, in the mouse embryo, as originating in the endoderm and mesoderm layers (Tam et al., 2007; Takaoka et al., 2012). A differential gradient of morphogens, primarily characterised by BMP4 and Nodal, patterns the PS into anterior and posterior axes (Bachiller et al., 2000). The expression of inhibitory factors including Lefty, Cer- 1, and Dkk-1 of BMPs, Wnt, and nodal signalling determine formation of the posterior-PS which develops into the kidney, muscles, blood, bone and gonads (Little et al., 2016). Loss of BMP4 expression results in mice which fail in mesoderm derivation (Winnier et al., 1995; Beppu et al., 2000). The posterior-PS region is then marked by the expression of T, Meox1, and Kdr (Tam et al., 2007). On the other hand, BMPs, Wnt, and Nodal signalling remain and govern the anterior-PS to originate a different sub-set of organs, such as lung, gut, and liver, derived from anterior PS-endoderm. Even though Nodal is required and expressed throughout the formation of PS, its expression is then gradually restricted to the anterior-PS by E.7.5 (Vincent et al., 2003). Therefore, the right balance between BMPs and Nodal along the PS can specify early formation of the endoderm (lowBMP/highNodal) and mesoderm (highBMP/lowNodal) layers during development.

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Figure 1.2. Fundamental embryonic decisions during kidney organogenesis. A) Upon epiblast formation, cellular movements give rise to primitive streak (PS) at the caudal edge of the embryo, characterized by formation of the three germ layers including ectoderm, mesoderm, and endoderm. B) The PS originates the trunk mesoderm which patterns into: the paraxial mesoderm (giving rise to vertebrate, dermis, ribs, and muscles), lateral plate mesoderm (giving rise to heart, blood vessels, body wall, and blood), and intermediate mesoderm (giving rise to kidneys and gonads). C) Intermediate mesoderm is determined in a temporal and spatial mode along an anteroposterior axis (A-P). PS cells migrating early towards the rostral end form anterior intermediate mesoderm (AIM). This gives rise to the nephric duct or Wolffian duct which elongates caudally to form the ureteric bud (UB). The UB will ultimately form the ducts of the urinary collecting system. D) PS cells migrating later form posterior intermediate mesoderm (PIM). This form the metanephric mesenchyme which reciprocally interacts with the UB to give rise to the permanent kidney. (Figure extracted from Little et al., 2016)

1.1.3.1.2 Intermediate mesoderm determination

Time lapse of chicken embryos has shown that, during elongation of the PS, daughter cells migrate towards the rostral end (James et al., 2003; Sweetman et al., 2008). Once there, and depending on their spatial distribution, they give rise to paraxial (PM), intermediate (IM) and lateral plate mesoderm (LPM) (Figure 1.2b). These events of cell determination and movement are mainly driven by temporo-spatial expression of morphogens including BMPs, Nodal, and FGFs. While lateral plate mesoderm is primordially specified by the action of BMP4 factor (James et al., 2005), inactivation of the same morphogen as well as Nodal signalling (Biben et

Commercial in Confidence al., 1998) is required for determination of the paraxial mesoderm. As expected, low BMP4 levels drive determination of the intermediate mesoderm, positioned between paraxial and lateral plate mesoderm (Obara-Ishihara et al., 1999; James et al., 2005). Gradients of FGF9 are also involved in PM as well as IM determination (Colvin et al., 1999). Odd-skipped related-1 factor (Odd1) is up-regulated early in the intermediate mesoderm (So et al., 1999). While Lhx1 (lim homeobox 1) is initially expressed in the LPM, its expression is then gradually restricted to the intermediate mesoderm (Cirio et al., 2011).

As mentioned earlier, all the renal structures originate from the intermediate mesoderm layer. To this end, the IM is also determined in a temporal and spatial mode along an anteroposterior axis (A-P) (Figure 1.2c and 1.2d). The cells which leave the streak late, and hence become the posterior cells, express Cyp26a1 which inactivates the RA signal, (Duester et al., 2008, Sakai et al., 2001). This enzyme is, therefore, important for the posterior patterning and not the anterior patterning. Conversely, cells that leave early, and hence become anterior, see RA early and are influenced by it because they no longer express Cyp26a1. Hence, of the mesoderm are influenced by high RA/FGF9 and low Wnt morphogens (Xu et al., 2014), and thus retinoic acid signalling is remarkably predominant in patterning the AIM. On the other hand, after the first rostral migration occurs there is a fraction of cells still populating the primitive streak (called the ‘late primitive streak’ or tailbud) characterised by T and Tbx6 expression. This population of cells migrate rostrally later than the earlier AIM cells, giving rise to the posterior-IM (PIM), and thus is less exposed to the RA gradient. Lineage tracing studies have shown that this PIM Eya1+ population forms the metanephric mesenchyme from which the nephron arises (Xu et al., 2014, Sajithlal et al., 2014). Therefore, these two distinct populations of the intermediate mesoderm (AIM and PIM) establish the niches from which the two kidney progenitor populations originate. AIM develops into nephric duct formation at E.8.0 and PIM develops into the metanephric mesenchyme (Taguchi et al., 2014, Takasato et al., 2015) (Figure 1.2c, d).

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1.1.3.2 Formation of three pairs of kidneys

In mammals, the development of the excretory organs entails the formation of three pairs of kidneys, which are referred to as pronephros at E8.0 (E22 in human), mesonephros E9.0 (E24 in human), and metanephros E10.5 (E35 in human), emerging at distinctive times from rostral to caudal, respectively (Figure 1.3) (Saxen et al., 1987, Takasato and Little, 2015). The most rudimentary structure, the pronephros, arises at the rostral end of the intermediate mesoderm (IM) and is characterised by formation of elementary ducts which drain into the nephric duct (ND) (Vetter et al., 1996, Takasato and Little, 2015). While the pronephric tubules form a functional organ in fish and in amphian larvae, they form a transient and vestigial structure with no function in mammals. Although the pronephros and much of the mesonephric kidney in mammal degenerates, some cranial mesonephric tubules contribute to the male reproductive tract (Toivonen 1957; Saxen 1987). Subsequently, although not yet a mature structure, the mesonephric kidney (Wolffian duct or nephric duct) is a more complex structure containing mesonephric tubules some of which open into the ND (Woolf et al., 2009, Vetter et al., 1996). Finally, the metanephric kidney originates caudally to the mesonephros, and is the definitive renal structure that persists after birth. The metanephros initially arises as swelling of the nephric duct and then invades the metanephric mesenchyme (Takasato and Little, 2015).

A) B) C)

Figure 1.3. Formation of three pairs of kidneys during mammalian development. A) The pronephros is the most rudimentary structure and arises at the rostral end of the intermediate mesoderm at E8.0 (E22 in human). B) The mesonephros forms the Wolffian duct or nephric duct E9.0 (E24 in human) which is a more complex structure although not the permanent structure yet. C) The metanephros originates the permanent kidney emerging at E10.5 (E35 in human). (Figure extracted from Takasato and Little, 2015).

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1.1.3.3 The metanephric kidney: nephrogenesis and branching morphogenesis

The metanephros is the permanent kidney structure which persists after birth. It arises as the result of reciprocal interactions between two embryonic populations, the metanephric mesenchyme (MM) and the ureteric bud (UB) (Figure 1.4) (Takasato and Little, 2015). Metanephric morphogenesis starts in humans around embryonic day 35-37 (E10.5 in mice) and is marked by invasion of the ureteric bud within the metanephric mesenchyme (Costantini 2006, Little al., 2016). After this, the ureteric bud commences a series of sequential divisions in a process called branching morphogenesis, resulting in an elaborate structure- the ‘ureteric tree’- from which the collecting duct system arises. The ureteric bud undergoes around fifteen events of dichotomous branching in the case of humans, and thirteen for mice (Short et al., 2014). Thus, the two essential processes underlying the formation of the adult kidney are: nephrogenesis, occurring in the MM; and branching morphogenesis, occurring in the UB.

Figure 1.4. Schematic representation of kidney organogenesis. The adult mouse kidney is organized into three main compartments- the cortex (containing glomeruli), the medulla, and the papilla- through which the collecting ducts drain. At the embryonic stage, the metanephric kidney arises from the ureteric bud (UB, brown) and the metanephric mesenchyme (MM, light blue). As the UB invades the MM, the cells closest to the ureteric bud form the cap mesenchyme (CM, dark blue) which undergoes trough nephrogenesis. The first structure is the renal vesicle (RV, purple), which then undergoes extension and segmentation giving rise to S-shaped bodies (SSBs). SSBs will then form a capillary loop and keep elongating to establish the mature nephron. (Figure extracted from Takasato and Little, 2015)

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1.1.3.3.1 Nephrogenesis

Although each progenitor population (MM and UB) is independently specified, normal UB branching and nephron formation depends on their reciprocal communication, which begins from week 8 to 9 of gestation for humans (Osathanondh et al., 1963). In fact, nephrogenesis is initiated by primary signals from the ureteric bud to the metanephric mesenchyme This process induces condensation of a group of cells adjacent to the tip of the ureteric bud, which undergo a mesenchyme-to-epithelial transition to form a renal vesicle (RV) (Figure 1.4) (Osthanondth et al., 1963, Dressler et al., 2006). Each RV then elongates to form the comma-shaped body, then forms a cleft in the distal region resulting in the S-shaped body (Osthanondth et al., 1963, Dressler et al., 2006, Saxen et al., 1987). The resulting S-shaped body segments into three segments: the lower limb (forms the glomerulus, Bowman’s space, and Bowman’s capsule); the central region (Loop of Henle and proximal convoluted tubule); and the upper lower limb which forms the distal tubule and fuses with the ureteric bud (UB). The initial cap-tip interaction is primarily induced by the action Wnt9b, expressed by the ureteric epithelium but then down-regulated to the tips where Wnt11 is highly expressed (Brunskill et al., 2008).

Nephron formation simultaneously occurs with the maintenance of an undifferentiated and densely packed population within MM called cap mesenchyme (CM), which is exclusively located in the periphery of the developing kidney, the nephrogenic zone (Figure 1.5). This self- renewing progenitor population (Six2+ Cited1+) drives branching morphogenesis and gives rise to nephrons (Little and McMahon, 2012, Boyle et al. 2008, Kobayashi et al., 2008). Since all the CM cells are potentially exposed to Wnt9b from the UB, it has been postulated that expression of certain genes within regions of the CM enable survival of the nephron progenitor population. The transcription factor Six2+ has been shown to be one such factor (Self et al., 2006). Deletion of Six2 results in the premature commitment of the nephron progenitor population, severely limiting organ growth and expansion (Self et al., 2006). In contrast, loss of Cited1 has no apparent phenotype suggesting that this is not required for progenitor turnover (Little and McMahon, 2012, Boyle et al., 2007). Expression levels of Six2 show an inverse correlation with rates of cell division (Short et al., 2014), and are high at the periphery of the kidney and gradually diminish moving towards the region of nephron formation (Georgas et al., 2009). However, not all Six2+ cells show equal properties. There is a Six2+ sub-population overlapping with Wnt4, representing the cells that have exited from the undifferentiated pool and are undergoing nephron formation (Little and McMahon, 2012, Kobayashi et al., 2008),

Commercial in Confidence while the population co-expressing Cited1+ forms the progenitor population (Mugford et al., 2008). As the nephrogenic process reaches its end, the cap mesenchyme progenitor population is exhausted, shortly after birth in mice (Hartman et al., 2007, Brunskill et al., 2011, Rumballe et al., 2011) and at around 36 weeks gestation in humans (Hinchliffe et al., 1991). In fact, around 90% of the nephrons are intensively formed during the latest stages of development (after E15.5 in mice). While it is thought that the initial depletion (up to E15.5 in mice) of the cap progenitor population is related to accelerated branching morphogenesis (Reviewed in Short and Smyth, 2016), the question of which factors determine total nephron number or cessation of nephron formation is still unresolved.

Figure 1.5. Overview of cellular compartments and molecular interaction in the nephrogenic niche. Four different compartments can be identified in the developing kidney: CapM: cap mesenchyme, Utip: The ureteric tip, PA-RV: pretubular aggregate-renal vesicle (forming nephron), Utrunk: ureteric trunk. Key molecular modulators are shown in the diagram and arrows indicate molecular interaction between different compartments. (Figure extracted from Little and McMahon, 2012).

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1.1.3.3.2 Ureteric bud outgrowth and the branching morphogenesis process

As mentioned above, the formation of a simple epithelial outgrowth called the ureteric bud distinguishes the formation of the metanephros from that of simpler excretory organs such as the pronephros and the mesonephros. The UB migrates towards the metanephric mesenchyme before commencing branching. This process of UB branching results in the formation of a complex dichotomously branching tree-like structure of tubules which will ultimately form the ducts of the urinary collecting duct system. The function of many other organs apart from the kidney, such as lungs, vasculature, and glandular tissues, also requires the formation of a tubular network. This is achieved by the process known as branching morphogenesis. The correct execution of this process is critical in determining the correct number of nephrons, and therefore of proper kidney function. As mentioned above, disruptions in this process result in a variety of congenital anomalies of the kidney and urinary tract (CAKUTs).

Many studies have investigated how branching morphogenesis is regulated in the developing kidney. Initially, branching morphogenesis was modelled in vitro using the Macdin-Darby canine kidney (MDCK) derived from canine distal tubule/collecting duct (Pollack et al., 1998, Santos et al., 1993, Sakurai et al., 1997). Subsequent cell lineage studies in mouse (25- Shakya et al., 2005) and analysis of differential gene expression by microarray (Schmidt-Ott et al., 2005) have revealed that the ureteric bud epithelium can be divided into two clear sub- populations, the T-shaped bud named ‘tip’, which specifically contains the self-renewing population, and the stalk. More recently, the branching process has also been defined to occur in four different stages (Bush et al., 2004, Nigam et al., 2003, Shah et al., 2004) defined by the presence or absence of different stimulatory and inhibitory factors (Figure 1.6) (Sampogna et al., 2004). These are:

Stage I: Initial UB outgrowth. This is mainly driven by the action of the GDNF/RET signalling pathway.

Stage II: Early rapid UB branching. Tissue recombination studies have provided evidence that the mutual interaction between MM and UB is critical in determining rapid extension of the collecting duct system (Bush et al., 2004, Quiao et al., 1999, Steer et al., 2002). This stage has been also shown to be sustained by the presence of other factors, such as primordially positive

Commercial in Confidence regulators like FGFs, pleiotrophin, heregulin, and, although less significantly, also by negative regulators (TGFβ1, BMP4, activin) in the UB-MM context (Bush et al., 2004, Lu et al., 2006).

Stage III: Late UB branching, MM differentiation. An important shift in the growth factors in the UB-MM context happens. The negative regulators become predominant, down-regulating active branching including transforming growth factor β (TGF-β), endostatin (Bush et al., 2004, Santos et al., 1994, Stuart et al., 2003) and bone morphogenetic proteins (BMPs) are described in this phase in order to terminate active branching. Notably, certain growth factors can play a positive/negative role over the course of this process, depending on which phase they are in and their concentration. For example, BMP7 when expressed at low levels favours branching, while at high levels it acts as an inhibitor (Gupta et al., 1999).

Stage IV: Branching cessation. Although there are still many unresolved question about how the branching morphogenesis reaches its end, it is clear that inhibitory factors from stage III become dominant and exercise a role in this final phase. In addition, it is also thought that nephron formation (mesenchymal structures such as the S-shaped body and comma-shaped body) may also influence branching cessation (Sariola et al., 2002).

This model of branching morphogenesis shows that the appearance of new branches basically entails an active remodelling of the apical cytoskeleton which is driven by the local and specific action of growth factors on the proliferative “tip” cells (Figure 1.6) (Nigam and Shah, 2009). In summary, as the branching morphogenesis proceeds the tip-proliferative cells are specifically restricted to the terminal branches having differential expression of growth factor receptors, extracellular matrix (ECM) components, matrix degrading enzymes compared to the cells populating the stalk (Shakya et al., 2005, Schmidt-Ott et al., 2005), sustaining the sequential rounds of division that establishes the ureteric-tree like structure.

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A B) )

Figure 1.6. Different stages of the renal branching morphogenesis process throughout kidney development. A) The mutual interaction of the metanephric mesenchyme (MM) and the ureteric bud (UB) give rise to the kidney. The ureteric bud epithelium branches in four different stages- Stage I: initial UB outgrowth, which is mainly driven by the action of the GDNF/RET signalling pathway. Stage II: early rapid UB branching, other signalling pathways promoting growth are also present. Stage III: late UB branching MM differentiation, inhibitory factors start to play a role, such as transforming growth factor β (TGF-β) and bone morphogenetic proteins (BMPs) among others. Stage IV: branching cessation, inhibitory factors from stage III remain present and likely nephron formation (mesenchymal structures such as the S-shaped body and comma-shaped body) may also influence branching cessation. In vitro models also support the existence of a budding and branching network. B) Representation of the mechanisms underlying differential formation of the tip and stalk. HSPG: heparan sulfate proteoglycans. GF: gradient formation. ECM: extracellular matrix components. (Figure extracted from Nigam and Shah, 2009).

1.1.4 Kidney repair in response to injury

As indicated, the formation of the entire complement of nephrons occurs prior to birth in humans (Little, 2006). Hence, injury to the organ after birth cannot result in the formation of new nephrons. However, there is extensive evidence that the epithelium of the nephrons themselves has a capacity to undergo proliferation and repair. The endogenous regenerative potential of the kidney is defined as the ability to repopulate and repair damaged structures. Evidence from mice subjected to ureteral ligation has shown that, although the kidney initially exhibits tubular atrophy, interstitial expansion, and ablation of an important part of its parenchyma, once the obstruction is removed the tissue begins a rapid process of remodelling without forming new nephrons (Cochrane et al., 2005). In the last decade many efforts have been made to elucidate the precise cellular origin of epithelial cells involved in kidney repair,

Commercial in Confidence which may contribute significantly to finding new cell-based treatments. Nevertheless, this reparative potential reaches its limit when faced with repetitive episodes of injury as well as chronic damage, when the kidney begins what is known known as ‘maladaptive kidney repair’. This process results in interstitial fibrosis, parenchymal loss, and therefore an irreversible loss of nephrons. In 2012, Grgic and colleagues investigated the response of the kidney to repeated injury in a genetic mice model, with the renal epithelium tagged with six2 to produce a receptor of diphtheria toxin. While these mouse kidneys exhibited an ability to respond and recover when faced with single events of injury, sustained damage resulted in an increase of macrophages infiltrates, myofibroblast proliferation, loss of vasculature, and fibrosis (3- Grgic et al., 2012). This evidence showed once more the primary role of the tubular epithelial cell, not only as a critical regulator of regeneration, but also in triggering the fibrotic reaction. It is thought that events such as increasing age, DNA damage, previous events of AKI, and sustained cells stress may mediate the secretion of growth factors and cytokines that stimulate the inflammatory response. Thus, the sustained release of inflammatory factors might also provoke pericyte dissociation from the endothelium, causing microvascular rare refraction and collagen deposition, leading to progressive fibrotic renal disease.

1.1.4.1 Cellular approaches to improving repair

In the attempt to find new treatments for kidney disease, different cell types have been postulated to hold great promise for contributing to the repair or regeneration of renal tissue. At present four possible origins have been proposed for cells contributing to postnatal renal repair: 1) interstitial cell transdifferentiation (Strutz et al., 1995); 2) recruitment of cells from the bone marrow (Cornacchia et al., 2001, Imasawa et al., 2001, Ito et al., 2001, Ito et al., 2001, Poulsom et., 2001); 3) tubular cell dedifferentiation (Strutz et al., 1995, Witzgall et al., 1994, Bonventre et al., 2003); and 4) the action of adult resident kidney stem cells (Kumar et al., 2015). Variable evidence is available to support the role of options 1 and 2 (Bussolati et al., 2005; Lazzeri et al., 2007; Angelotti et al., 2012). Debate has actually focussed on trying to elucidate whether repair is mediated by tubular epithelial cells or by stem cells resident within the kidney.

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In support of the epithelial model, it has been shown that the proximal tubule expresses mesenchymal markers as vimentin after an injury episode, implying that dediferentiation of the tubular cells may mediate the repair process (Bonventre et al., 2003). On the other hand, a resident tubular stem cell population (CD133+ and CD24+), termed scattered tubular cells (STCs), was initially proposed to be a fixed population resident in tubular compartments which could ameliorate injury in immunodeficient animals with AKI (Cornacchia et al., 2001, Imasawa et al., 2001, Ito et al., 2001, Ito T et al., 2001, Poulsom et., 2001). Later, Berger et al., (2014) reported that the first tracking study of STCs provided strong evidence that this stem cell population was not a fixed intratubular progenitor population, and that these cells might rather arise from any tubular cell as a result of spontaneous de-differentiation triggered by local injury (Berger et al., 2014).

Aside from the possibility of intraepithelial progenitors that contribute to repair, it has also been proposed that the kidney, like many other organs, contains a resident mesenchymal stem cell (MSC) population that can play a role in tissue repair. Mesenchymal stem cells (MSCs) were first isolated from the bone marrow and defined based upon their capacity to form fibroblastic colonies (Friedenstein et al., 1982). The capacity for MSCs, either isolated from the bone marrow or a wide variety of other tissues, including fat and umbilical cord, to act as regenerative cells for use in cellular therapy is of major interest for many disease states. Indeed, MSC clinical trials around the globe have administered to more than thousand patients, either by autologous or allogenic transplantation (Duffield et al., 2005, Da Silva Meirelles et al., 2006, Nombela-Arrieta et al., 2011, Dominici et al., 2006, Pittenger et al., 1999). There is a broad consensus that MSCs are able to ameliorate tissue damage in response to injury and allograft rejection. Although these cells were originally found in the bone marrow (bm-MSCs), they have also been found to reside in many adult tissues including the kidney, possibly representing a perivascular cell population involved in normal tissue repair (Rinkevich et al., 2014, Lasagni et al., 2015, Kusaba et al., 2014). The current knowledge on MSCs and how they contribute to tissue repair will be broadly covered in Chapter Two, since one of the major aims of this thesis is the examination of the functional contribution of an endogenous kidney mesenchymal stem cell population to kidney repair. Some authors have also claimed that macrophages have a role in contributing to kidney repair (Ricardo et al., 2008). In fact, their reparative action has been related to the release of interleukin (IL-4, IL-10) and inducible nitric oxide synthase (Kluth et al., 2001; Riquelme et al., 2013; Cao et al., 2010; Lu et al., 2013; Ferenbach et al., 2010). Taken together, renal progenitor cells (RPCs), mesenchymal stem cells (MSCs), and ultimately

Commercial in Confidence macrophages, have exhibited the capacity to contribute to postnatal repair, although their contribution is limited, and it is still unclear if the administration of these cells to patients will have long-term efficacy or side effects.

1.1.5 Kidney Regeneration

While improvements in tissue repair are of great importance, and contribute to the field of regenerative medicine, they are limited to the amelioration of renal damage. Consequently, there has been enormous interest in attempting to regenerate kidney tissue using stem cells from a variety of sources, including human foetal stem cells, cells from the amniotic fluid, or cells derived from the directed differentiation of human PSCs (ESCs, embryonic stem cells or iPSCs, induced pluripotent stem cells) into kidney cell types. While human foetal stem cells derived from human foetal kidney or from the amniotic fluid represent optional sources of cells, they are not easily accessible. Also, they have not shown enough evidence of being able to generate the variety of cell types required when attempting to regenerate an organ. These limitations were later resolved when the emergence of iPSCs revolutionised regenerative medicine.

1.1.5.1 Induced Pluripotent Stem Cells (iPSCs): a rapidly evolving technology

The existence of induced pluripotent stem cells was firstly demonstrated in mouse (Takahasashi et al., 2006), and a year later in humans (Takahasashi et al., 2007). These cells have the unique ability to differentiate into any somatic cell type from any available cell type. The pluripotency state is restricted to the epiblast phase of the developing embryo, thus representing cell lines equivalent to an embryonic derived cell line. In a process referred to as ‘reprograming’ adult somatic cells were converted to an embryo-like state by the overexpression of four transcription factors, including Oct3/4, Sox2, and Klf14, which also repressed lineage-specific genes. The resulting iPSCs are genetically and morphologically like human ECSs, and exhibit the ability to differentiate into different cell types from the three germ layers- endoderm, mesoderm and ectoderm (Takahasashi et al., 2007). Later, other researcher also successfully reprogrammed iPSCs using a slightly different cocktail of transcription factors. This demonstrated the reproducibility and robustness of this approach and began a new era in the field of stem cell biology (Yu et al., 2007, Wernig et al., 2007; Park et al., 2008).

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Since the generation of iPSCs from adult human skin fibroblast began of routine use for research, this technology has been rapidly progressing. iPSCs can now be rapidly generated by transfection of episomal and retroviral cassettes. This process introduces the reprogramming genes while avoiding any genetic footprint from either the donor or the reprograming process. Recent advances in genome editing technologies, including the transcription activator-like effector nuclease (TALEN) and clustered regularly interspaced short palindromic repeats (CRISPR), have enabled the correction of inherited mutations within patient-derived iPSCs (Hotta et al., 2015). Also, the development of fluorescently-tagged iPSC lines has provided a valuable tool for the study of, developmental processes or the dissection of important lineage relationships, for example.

The first study reporting the generation of kidney cell types dates back to only 2012, with the derivation of podocyte-like cells from ESCs (Song et al., 2012). Since then there has been an extraordinarily rapid development of the nephrology field, with reports of different protocols for deriving kidney cell types from iPSCs. This will be reviewed in detail in Chapter Three. In the last three years the recreation of many kidney cell types has become possible. With the generation of kidney organoids by our group being one of the major breakthroughs to the kidney field in 2015 (Takasato et al., 2015 and 2016). We have seen evidence of structures containing ureteric, nephron, vascular and stromal derived cells. Despite of this remarkable discovery, this approach is self-limiting due to the lack of sustained renal progenitor populations across time, and the relatively short time period for cultures in this format. To make this technology of transferable use, the two key progenitor cell types, the nephron and the ureteric bud progenitors, needs to be recreated to ensure ongoing morphogenesis.

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1.1.6 Significance and approaches of this thesis

As we have discussed throughout this literature review, the increasing worldwide prevalence of chronic kidney damage has increased the need to find renal replacement therapies. Coupled with the reducing availability of organs for transplantation, there is a necessity for development of novel approaches to repair or regeneration. The work described in this thesis examined and challenged recent concepts in normal kidney organogenesis, repair, and regeneration with a focus on one renal compartment, the collecting duct system. In Chapter Two, we examined the capacity of an endogenous kidney MSCs population (k-MSCs) to functionally integrate into the mature collecting duct system (Figure 1.7). While in this thesis, the approach was to prove that this integration would effect the recapitulation of a disease state, the objective long term was to potentially use this approach as a potential means for effecting repair. In Chapters Three and Four, we addressed the ambitious attempt to recreate one of the two progenitor populations which gives rise to the kidney, the ‘ureteric bud population’ (UB), via the directed differentiation of human pluripotent stem cells. It is hoped that this might improve our capacity to recreate kidney tissue for therapeutic uses (Figure 1.7). In addition, the establishment of fluorescently-tagged human iPSCs cultures provided a unique opportunity to trace lineage within organoids. This allowed the evaluation of progress in real time and with a much higher- throughput than with animal models. We also anticipate that the generation of an improved protocol for the generation of UB progenitor cells will ultimately facilitate bioprinting in 3D and allow the recreation of larger kidney structures.

Figure 1.7. Schematic diagram depicting the approaches taken in this thesis.

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CHAPTER II

Evaluating the potential contribution of k-MSCs to collecting duct epithelium.

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2.1 INTRODUCTION

New treatments for acute kidney injury (AKI) and chronic kidney disease (CKD) are of high priority in nephrology. Although the literature has described a cellular reparative capacity in the adult kidney (a capacity to repair after acute injury), it has not proven structural regeneration (a capacity for regenerate structures). A deeper revision of the current knowledge about the molecular mechanisms involved renal repair, as well as recent advances in regenerative medicine, is presented in the two published reviews authored during this PhD candidature listed as appendix # 1(Does Renal Repair Recapitulate Kidney Development?) and # 2 (Regenerative medicine in kidney disease).

In an attempt to create innovative interventions, recent studies have tested stem cell-based technology, with mesenchymal stem cells (MSCs) the most widely used cell. MSCs provide the exciting prospect of a powerful treatment to repair acutely damaged organs by virtue of their unique stem cell tropism and pro-regenerative capacity. Here, we present a review of the literature focused on current knowledge of the biology and therapeutic advantages of bone marrow versus kidney derived MSC-like cells. This chapter proposes to elucidate if kidney- derived MSCs represent a preferable population for further use in kidney disease.

2.1.1 The mesenchymal stem cell

2.1.1.1 MSCs in the Context of Stem Cells

During the last decade, much progress has been made on stem cell research due to their unique properties. Stem cells are undifferentiated cells which are mainly described by their self- renewal and long term proliferation capacity, and their ability to give rise to more specialized cells (Wagers and Weissman, 2004). Regarding differentiation potential, stem cells are classically classified in five categories: totipotent (able to give rise to all embryonic and extraembryonic cell types), pluripotent (able to give rise to any cell derived from the three germs layers and the germ cells); multipotent (able to give rise to multiple cell lineages but usually tissue restricted); oligopotent (able to give rise to a restricted number of cell lineages); and, finally, unipotent (able to give rise to an exclusive cell type) (Wagers and Weissman, 2004). Stem cells can also be classified according to the developmental stage from where they

Commercial in Confidence are isolated, and are therefore termed embryonic or adult stem cells. It is well known that stem cells from embryonic origin are more potent than adult stem cells, due to their greater differentiation potential. Nevertheless, their use is still a controversial topic. In this context, adult stem cells appear to have several advantages as they can be easily isolated from a patient to be utilized for transplant, thereby avoiding immunological rejection compared to non- autologous pluripotent foetal cells. As the focus of this chapter is to evaluate the potential of mesenchymal stem cells, such cells will be reviewed in detail.

2.1.1.2 General Properties of Bone Marrow MSCs (bm-MSCs)

Mesenchymal stem cells or multipotent mesenchymal stromal cells (both referred to as MSCs) (Horwitz et al., 2005) are stem-like cells, traditionally of bone marrow origin (bm-MSC). They were first described based on their ability to adhere to plastic and their fibroblastic phenotype (Friedenstein et al., 1982). However, in recent years, MSCs have also been identified, based on cell surface immnunophenotype and mesodermal differentiation capacity, to reside in many postnatal organs (Figure 1), possibly representing a perivascular cell population involved in normal tissue homeostasis (Crisan et al., 2008, da Silva Meirelles et al., 2006, Nombela-Arrieta et al., 2011).

Given that this cell type has a high potential use for cellular therapies in humans, the number of studies related to MSC has enormously increased during the last years. To unify the criteria to identify human MSCs, Dominici et al., reported the basic characteristics of human MSCs as: multipotentiality (notably to mesenchymal cell types), adherence to plastic in culture, the expression of marker of proteins such as CD73, CD90, and CD105, and the lack of expression of hematopoietic lineage markers as CD45, CD34, CD14, CD11b, CD79α, CD19 and HLA- DR surface molecules (Dominici et al., 2006).

MSCs deserve attention for their proven plasticity, being capable of differentiating into cell types other than their tissue of origin. Under specific conditions these cells can differentiate to osteocytes, adipocytes and chondrocytes (Figure 2.1) (Pittenger et al., 1999, Dominici et al., 2006). Mesenchymal stromal cells have been also reported to elicit a paracrine role, releasing growth factors such as vascular endothelial growth factor (VEGF), hepatic growth factor (HGF), insulin-like growth factor-1 (IGF-1), and anti-apoptotic cytokines that might be

Commercial in Confidence released to the place of engraftment (Bi et al., 2007, Togel et al., 2005, Imberti et al., 2007). However, an accurate profile of these pro-reparative factors is still unclear.

Clinical trials have also taken advantage of some properties of MSCs. For example, their immunomodulatory capacity, including suppression of T cell proliferation, and the alteration to the cytokine secretion profile of dendritic cells, natural killer and T cells promote a more tolerant phenotype by diminishing their inflammatory environment (Aggarwal and Pittenger, 2005, Krampera et al., 2003, Di Nicola et al., 2002, Stagg, 2007). Consequently, MSCs are clearly a promising candidate for cell-based therapies based mainly on immunomodulatory and/or tissue repair properties. This cell population has been also described to amilorate some diseases, such as chronic kidney disease (CKD), and to reduce allograft rejection (Reinders et al., 2010). In the recent years, more than 1000 patients have received cellular therapy based on the use of MSCs (Rabelink TJ 2012; Salem HK 2010; Reinders ME 2014; Fibbe WE 2007; Nauta AJ 2007; Tan MD 2013). However, it is still unclear if those cells have long term efficiency or side effects (Le Blanc and Ringden, 2007, Reinders et al., 2010).

Figure 2.1. Multipotent mesenchymal stromal cells in adult tissues. The plastic-adherent cellular fraction of many organs contains stromal progenitor cells that can give rise to colonies of fibroblastic morphology, the hallmark that defines mesenchymal stromal cells is their ability to differentiate into osteoblasts, adipocytes and chondrocytes when placed under inductive stimuli. Differentiation into multiple non-mesenchymal mature cell types (such as muscle cells, endothelial cells and neural cells) has been reported, but remains a matter of debate (Image extracted from Nombela-Arrieta C et al., 2011)

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2.1.2 The delivery of bone marrow MSCs into kidney disease models

2.1.2.1 Acute Kidney Injury (AKI)

In the kidney as well as other organs, numerous studies have reported beneficial effects of bone marrow MSCs in tissue repair and regeneration. These cells are delivered into defective animal models induced by chemical-induced (glycerol, mercury, chloride and cisplatin) or ischemia- induced injury (Tögel et al., 2005, Togel et al., 2009, Imberti et al., 2007, Bi et al., 2007). It was initially thought that these cells exerted their reparative effect through the replacement of renal tubular epithelial cells (Herrera et al., 2004). However, bone marrow MSCs have not been able to undergo direct differentiation into epithelial cells in order to contribute to tubular regeneration. In support of this, one report, using a model of ischemia reperfusion injury, showed that kidney structure was repaired by intrinsic tubular cells, rather than by exogenous cells (Humphreys et al., 2008), suggesting that bone marrow-MSCs elicit their effects by the secretion of paracrine factors. Ample literature exists supporting this claim. Bi B et al. (Bi et al., 2007) studied the treatment of a kidney injury model induced by cisplatin (an antineoplastic drug known for its nephrotoxicity effects) with bm-MSCs. The method was carried out through independent injections of bone marrow-stromal cells or conditioned medium obtained from the same type of cells to different animal groups. The authors found that both groups of animals showed increased survival and renal injury was limited, suggesting that repair was due to secreted reparative factors and did not require the cells themselves. A better understanding of what are the exact factors and their proportions in the conditioned medium is still required. To date, several reports support the fact that these cells release growth factors as HGF, IGF1, VEGF and EGF which are widely indicated to improve renal function (Bi et al., 2007, Togel et al., 2005, Imberti et al., 2007, Togel et al., 2009).

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2.1.2.2 Chronic Kidney Disease

As mentioned above, our knowledge of the paracrine reparative effect of MSC on AKI has grown tremendously in the past few decades. However there has not been much progress on whether MScs exert a positive effect on chronic kidney disease.

To identify the role of MSCs play in a progressive model of renal failure, a glomerulonephritis model (GN) was used, where renal parameters were followed until 10, 30 and 60 days after MSCs injections. Here, the parameters of renal function were restored in the first stage. Surprisingly, the study showed after long-term assessment that MSCs underwent maldifferentiation into cells with adipocyte features within the glomerular structure (Kunter et al., 2007). This obviously raised the concern about the safety of MSCs-based cell therapy. Controversially, another report that performed weekly injections of MSCs between 6 to 10 weeks in an Alport syndrome model (collagen type Iα2-deficient mouse), indicated that there were no response to the administered cellular therapy in terms of blood urea nitrogen, creatinine and proteinuria. In this model, MSC- based therapy failed to delay the renal phenotype, but there was a reduction in interstitial fibrosis (Ninichuk et al., 2007).

Similarly, another genetic model described for the same pathology (collagen type I alpha 2- deficient mouse) was treated with human first-trimester blood MSCs. An improvement in this glomerulopathy was shown to occur by the preferential recruitment of donor cells by damaged col1α2-deficient glomeruli. A decrease in collagen type I deposition in the glomeruli of transplanted mice compared to non-transplanted was also reported (Guillot et al., 2008). These findings reinforce the high potential of human foetal MSCs for the treatment of genetic diseases. However, this study did not resolve the question of whether renal function was re- established.

Due to the available information, it is clear that several critical questions remain unresolved and cause ongoing debate especially in treatment of chronic kidney diseases. One important question is whether these cells will be able to undergo a long-term response and if there are any side effects. As a summary of the current knowledge about treatment based on bone marrow-MSCS, two points can be highlighted: 1) They have a short-term reparative effect for the treatment of kidney diseases; and 2), bone-marrow MSCs cannot turn into epithelial cells, and thus, they have mainly been assessed in acute kidney injury models.

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2.1.3 The kidney mesenchymal stromal cell population (k-MSCs)

2.1.3.1 Phenotypic Characterization of k-MSCs

As mentioned earlier, it is currently recognized that mesenchymal stromal cells are present in many post-natal organs (Crisan et al., 2008), possibly representing a reparative niche. These postnatal-derived MSC populations were initially proposed to arise from the perivascular niche, based on the observation of the following expression markers: CD146+NG2+PDGF- Rβ+ (Crisan et al., 2008, da Silva Meirelles et al., 2006, Meirelles Lda and Nardi, 2003). Our knowledge of adult-derived MSCs has grown tremendously in the past few years, and our laboratory has made important contributions, particularly related to the mesenchymal stromal population resident in postnatal kidney.

In 2012, our research group reported and characterized an endogenous kidney population of mesenchymal stromal cells (k-MSCs). This population was isolated from adult murine kidneys, and compared to populations from adult murine bone marrow (bm-MSCs) and heart (h-MSCs), observing several common features of MSCs including an ability to form fibroblastic colonies and differentiate to adipocytes, osteocytes, chondrocytes (Figure 2.2A) (Pelekanos et al., 2012). Regarding the kidney population, these showed a long-term clonogenic potential of 1%, and could be expanded as fibroblastic colonies for more than 30 passages under tissue culture conditions. Additionally, the k-MSCs inmunophenotypic profile was positive for CD29+, CD44+, CD90+, all markers previously described as being positive on BM-MSCs (Figure 2.2B). Immunosuppressive capacity was also assessed by measuring the ability to suppress T- cell proliferation of each population, using a low (3x103) and high (3x104) number of MSCs. In particular, it was shown that bone marrow MSCs had effective suppressive capacity in both cases, whereas the kidney MSC population only showed this potential in response to a higher number of cells (Figure 2.2C) (Pelekanos et al., 2012). Whereas this study showed clear phenotypical similarities among the three populations, the expression of some genes was only selectively over-expressed in specific populations. For example, in K-MSCs, expression of collecting duct markers such as natriuretic peptide precursor type B, Uroplakin 1b and Hoxb7 showed an enrichment for kidney-specific gene expression (Pelekanos et al., 2012). These data reinforced the concept of the retention of organ specific roles, and raised the question of

Commercial in Confidence whether the kidney mesenchymal stem cell-like population within the adult mouse kidney has important therapeutic implications, leaving room for further research.

A) B)

Figure 2.2. Characterization of endogenous kidney MSC (kCFU-F) compared to heart (cCFU-F) and bone marrow populations (bm-MSC). A) Analysis of the mesodermal potential of the three populations a-c) Light microscopy of adherent cultures. d-f) Light microscopy of Giemsa-stained cystospin preparations. g-i) Oil red-o staining of lipid droplets after 21 day culture in adipogenic media. j-l) Alizarin red staining of osteoid matrix after 21 day culture in osteogenic media. m-o) Alcian blue staining of proteoglycans after 21 day pellet culture in chondrogenic media. B) Comparative immunophenotyping of bone marrow, cardiac and kidney MSCs showing strong similarity across 81 epitopes. C) Immunosuppresive activity of bone marrow and kidney derived MSCs in a mix lymphocyte reaction. (Figure adapted from Pelakanos et al., 2012).

2.1.3.2 New Insights and Tracing the Origin of k-MSCs

In order to assess whether k-MSCs can play a role in tissue homeostasis and/or renal repair, our group decided to further investigate the origin and fate of these cells when they are put back into the kidney (Li et al., 2015). Firstly, to study the fate of the cells, GFP+ kidney MSC- like cells were microinjected into the renal parenchyma of neonates at postnatal day 1 (PD1), using GFP+ bone marrow MSCs as controls. The injections were performed at the time when nephron formation and papillary maturation were still occurring (Vanslambrouck et al., 2011, Rumballe et al., 2011). Kidneys were then analyzed up to 14 days post-delivery. Although GFP+ BM-MSCs were not detected in the kidney, surprisingly GFP+ k-MSCs were detected in

Commercial in Confidence collecting duct structures within the medullary/papillary region, co-localizing with Aquaporin 2 positive cells. In this study, all the cell incorporations were detected in the same region of the kidney, suggesting an in vivo capacity for the injected MSCs to undergo a mesenchyme to epithelial transition (Figure 2.3) (Li et al., 2015). The second approach was the identification of the cellular origin. To achieve this, cells were sorted using three different methodologies, 1) the expression of CD surface markers 2) manual dissection of the papilla or medulla region and 3) the expression of the specific collecting duct promoter, Hoxb7. After 14 days in tissue culture conditions, the papilla region displayed the strongest potential to form colonies and the Hoxb7 fraction higher clonogenicity. Therefore, the population from the medulla/papilla showed a stronger stem cell-like phenotype (Li et al., 2015). Even more promising was the fact that HoxB7-sorted k-MSCs retained their epithelial potential, forming E-cadherin+ branching epithelial structures under in vitro and in vivo conditions. Additionally, a repair activity was also identified from k-MSC-conditioned media. Based on the previous facts, the k-MSC population holds greats promise for the treatment of chronic kidney disease. A) B)

Figure 2.3. Specific epithelial integration of kidney MSCs after their microinjection into the nephrogenic zone. Bone marrow and kidney mesenchymal stromal cells isolated from ubiquitously GFP-expressing mouse. A, a) Ultrasound image of neonatal mouse kidney outlined by dashed line. b) Section of neonatal kidney after injections of k-MSCs with green fluorescent beads to assist in the localization. B) Low resolution confocal images show the GFP+ tubular structures exclusively in the medulla region of those mice injected with k-MSCs. C) High magnification of confocal images for k-MSCs against GFP+ and collecting duct epithelium (Aqp2+) showing co-localization in the medulla region. (Figure adapted from Li et al., 2015.)

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2.1.4 The selected model of renal disease: Dominant polycystic kidney disease

As it has been discussed through this literature review, the main purpose of this research project is to identify whether kidney MSC-like cells display a functional integration into the medullary collecting duct as was proposed in Li J et al., 2015. If the latter is true, it will further open the possibility of human clinical trials and the development of this cell type for use in new cell- based therapies. However, this relies upon the long term functional integration of these cells into collecting duct epithelium. To assess this possibility, it was proposed to test the epithelial potential of these cells to trigger a defective kidney phenotype. The chosen gene for this purpose was Pkd1 due to its described role in the generation of Autosomal Polycystic Kidney Disease (ADPKD) in which cyst formation is the dominant feature.

2.1.4.1 Autosomal Dominant Polycystic Kidney Disease and the Pkd1 Mouse Model

Polycystic kidney disease is a genetic disorder that can be transmitted as autosomal recessive or as dominant inheritance. Autosomal dominant polycystic kidney disease (ADPKD) is the most common mode of inheritance, with ADPKD occurring in 1 in 1000 live births but the disease commonly presenting in adult life (Fick and Gabow, 1994, Wilson, 2004, Harris and Torres, 2009). It is characterized by the slow formation of numerous cysts throughout the cortex and medulla with cysts arising from every segment of the nephron because of an expansion of epithelial cells. Although the initiating factor of cyst formation is unknown, evidence exists that cysts formation is accompanied by loss of the remaining copy of the mutated gene such that the disease is recessive at the level of the epithelium. It is thought that these affected cells then undergo selective proliferation which proceeds indefinitely (Sullivan et al., 1998). Finally, as such epithelial structures grow, they disconnect from the tubular structure, but still accumulate liquid within the lumen. Indeed, the enlargement of the cysts leads to a decline in the number of functional nephrons causing the progression of the disease (Figure 2.4). Although ADPKD has considerable variation in terms of its phenotype, clinical manifestations can be defined as the inability to concentrate urine, acute and chronic pain. In some cases, there is a progression to end-stage renal disease (Fick and Gabow, 1994, Sullivan et al., 1998).

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Figure 2.4. Cysts formation at the level of cell, nephron and kidney. Defects in the genes encoding PC1 or PC2 lead to aberrant gene transcription, cell proliferation and ion secretion which in turn result in the formation of fluid-filled cysts. As cysts balloon out from individual nephrons, their collective effect lead to the displacement of the normal renal parenchyma and the formation of cysts-filled kidney, with reduced functional capacity (Chapin et al., 2010).

There are two types of ADPKD: Type I is caused by mutations in the PKD1 gene and accounts for 85 to 90 percent of cases (1994). Type II is caused by mutations in the PKD2 gene, and accounts for 10 to 15 percent of cases (Mochizuki et al., 1996). For both genes, mutation can be either truncation or missense, however, both appear to inactivate the genes involved (Calvet, 1998). Moreover, it has been proven that lowering of Pkd1 expression is sufficient to initiate cystogenesis in mice (Lantinga-van Leeuwen et al., 2004). Mice with homozygous targeted disruptions of the pkd1 or pkd2 gene die in utero or perinatally (Lu et al., 1997), showing that those genes are essential for normal development and renal tubular morphology.

The proteins encoded by PKD1 and PKD2 genes have different functions. Polycystin-1 (PC1) is an integral membrane protein with 11 transmembrane domains and an extracellular region consisting of a variety of domains. It interacts with many molecules eliciting intracellular responses through phosphorylation pathways. On the other hand, polycystin-2 (PC2) has been identified as a non-selective calcium channel (Gonzalez-Perrett et al., 2001, Koulen et al., 2002). It is thought that both proteins function as a complex, regulating levels of intracellular calcium. Polycystin-1 and 2 have been reported to be located in the lateral domain of the plasma membrane, playing a role in cell-cell, cell-matrix interactions and also in the primary cilia (Figure 2.5) (Pazour et al., 2002, Yoder et al., 2002, Ibraghimov-Beskrovnaya et al., 1997). It is still not fully understood why mutations in these proteins result in cyst formation. Yet it is

Commercial in Confidence well-known that they are associated with several vital biological processes, such as signal transduction, cell-cell interactions, cell-cycle control and transcriptional regulation (Chapin and Caplan, 2010, Ibraghimov-Beskrovnaya and Bukanov, 2008). Recently, several clues have pointed towards their role in the primary cilium, which functions in mechanosensation of urine- flow (Berbari et al., 2009, Jonassen et al., 2008). Cilia defects also play a role in the establishment of improper planar cell polarity in epithelial cells, which might have an effect on tubular dilation and cyst formation (Bagherie-Lachidan and McNeill, 2010).

Figure 2.5. The function of polycystin-1. Polycystin-1 complexes are found at the cell– matrix interface, cell–cell contacts, and luminal cilium, where they are thought to function as sensors of the extracellular environment and interact with proteins of the cell membrane and actin and tubulin cytoskeleton and transduce signals by means of intracellular phosphorylation cascades to regulate gene transcription in the nucleus. The polycystin-1 C- terminal contains sites for phosphorylation on serines (by protein kinases A and X) and on tyrosines (by c-src and focal adhesion kinase), as well as proline-rich src homology 3 (SH3) and putative WW sites. Signal-transduction cascades induced by the polycystin complex include those of the Wnt pathway (by means of b-catenin and T-cell [TCF] and lymphoid- enhancing [LEF] transcription factors), the focal adhesion pathway (by means of MAPkinase [MAPK], JUN kinase [JNK], and activating protein 1 [AP-1] transcription), and the JAK2–STAT1 pathways, suggesting transcriptional regulation of proliferation, apoptosis, epithelial differentiation, polarity, adhesion, migration, cell shape, and tubular diameter, which are all components of renal morphogenesis. (Image adapted from Wilson et al., 2004)

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A most remarkable finding was published in 2015 by Leonhard et al., describing a cystic snowball effect in a tamoxifen inducible Pkd1 mice. Here, where the number of Pkd1-deficient cells (total loss of Pkd1) was reduced to 8%, cysts did occur and increase rapidly across time. This was the first study in animals in which the deletion in the Pkd1 gene was not active in a large number of cells. Interestingly, this report showed that scattered pkd1 deletion leads to a long dormant period of six months which is then followed by a rapid onset of severe polycystic kidney disease where most of the mice developed massive cysts structures through all the kidney around 9-11 months of age. Therefore, this study represented closely a model in animals that mimicked human ADPKD showing that only a small number of mutant epithelial cells was enough to trigger the phenotype. Hence, this study provided strong evidence that the functional integration of even a small number of Pkd1 mutant k-MSCs should lead to cystic disease in the normal recipient mouse.

2.1.5 Hypothesis and aims

Hypothesis:

Kidney-derived MSCs can functionally integrate into collecting duct epithelium and contribute to collecting duct function through adult life.

AIM 1. Evaluate the longevity of integrated kidney MSC-like cells after neonatal injections

AIM 2. Generation and Characterization of the futures of Pkd1 mutant cell lines

AIM 3. In vivo recapitulation of polycystic kidney as a measure of functional integration into collecting duct

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2.2 RESULTS

AIM 1: Evaluate the longevity of integrated k-MSCs (GFP+) after neonatal injections.

As mentioned earlier, a recently published study from our laboratory has shown that kidney derived MSC-like cells are enriched in the papilla region and that a robust cell population can also be obtained when those cells are sorted from collecting duct epithelium (Li et al., 2015). Even more promising was the fact that when those cells were put back into the neonatal kidney at postnatal day 1 (PND1), k-MSCs were uniquely found in collecting duct epithelium. Such integrations have been shown to persist up to 15 days post-delivery. Hence, these data supported the hypothesis that k-MSCs were able to undergo a successful homing and integration process. However, whether these cells can be detected after a long period of time, and whether they play a functional role in the kidney, are questions that remain unresolved. To investigate this, we first evaluated whether k-MSCs persist within collecting duct epithelium over a long period of time. To achieve this, a round of microinjections of k-MSCs was performed into neonates following the same protocol previously established by Li et al., 2015.

Based on the fact that the capacity of EGFP+ cells to integrate into collecting duct structures was previously found and reported by our laboratory (Li et al., 2015), we investigated the capability of k-MSCs (GFP+) to remain integrated within kidney structures over a long period of time. In this study, we used the same constitutionally expressed GFP+ cells as in our previous publication. Long-term evaluation was carried out up until 10 weeks after the post- delivery of k-MSCs into CD1 mice. k-MSCs (GFP+) were detectable in the kidney and were exclusively co-localised with Aqp2+ cells (Figure 2.6 and 2.7). In addition, we were able to detect k-MSCs (GFP+) over all three time-points evaluated and within the same kidney compartments as was previously reported. Notably, except for the beads themselves, no GFP+ signals were detected in the control samples performed.

To determine the relative contribution of k-MSCs (GFP+) to the papillary tubular collecting duct epithelium, a quantification of the ratio GFP+/Aqp2+ was performed. Previous analyses indicated that 12.0±0.8% of medullary collecting duct cells were derived from injected k-MSCs at 4 days after neonatal injection (Li et al., 2015). We therefore calculated the integration percentage of the injected cells after an exhaustive examination of the GFP+/Aqp2+ immunofluorescence of kidney medulla/papilla sections obtained at 4, 6, and 10 weeks post-

Commercial in Confidence delivery (detailed in the Materials and Methods Section). For each time-point, three different independent injection experiments were analysed and, from them, between two to three different frozen kidneys sections were studied, previously stained against GFP+ and Aqp2+ antibodies. This analysis showed the following relative incorporation: P4 weeks, 9.7 ±0.04% Aqp2+ collecting duct cells were GFP+ (3,917 Aqp2+ nuclei counted); P6 weeks, 6.3±0.01% GFP+ (2,365 Aqp2+ nuclei counted); and P10 weeks, 7.0±0.03% GFP+ (2603 Aqp2+ nuclei counted) (Figure 2.7). Taken together, the immunofluorescence studies clearly showed that microinjected k-MSCs remain detectable in collecting duct structures over a long-term period, showing an integration percentage average of 7.6% in the three groups studied. After comparison with the previous data from our laboratory, we can see that there is not a significant change to this percentage of integration over time. These data do not support the possibility that the integrated cells are being phagocytosed by the collecting duct (Kim et al., 1996) or replaced after a longer period of time; rather, they support the hypothesis of functional integration. Hence delivering k-MSCs into animal models, either to apply a treatment or to induce a defective phenotype, appears feasible.

Figure 2.6. Specific epithelial integration of k-MSCs (GFP+) is detectable within collecting duct compartments after 4, 6, and 10 weeks post-delivery. A representative panel of immunofluorescence assays performed against GFP and Aqp2+ antibodies is shown from a random sample. A) Low- magnification of a confocal merged image (20X) showing GFP+Aqp2+ tubular structures in the medullary region of injected mice. B) and C) High resolution of confocal images (40X) showing independent detection of GFP+ and Aqp2+ within the close-up region shown in A (n=3, for each age group).

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Figure 2.7. Quantification of the long-term integration capacity of k-MSCs (GFP+) into collecting duct structures via the calculation of the ratio GFP+Aqp2+/TotalAqp+2. Immunofluorescence analysis of kidney sections collected from mice injected with k-MSCs (GFP+) was performed against GFP+ and Aqp2+. Three injected mice were analysed for each group and a total of three sections were analysed for each animal. All images (300 dpi at X20) were captured and analysed with Imares software. Data are presented in the bars as the mean of the group. Statistical analysis was performed by t-test. * means statistical significance using the group of mice injected at 4 weeks as control group with a p value =0.05.

AIM 2: Generation and phenotypic characterisation of Pkd1 mutant cell lines

To demonstrate the functional ability of k-MSCs to integrate into collecting duct structures, this aim proposed to generate and characterise mutant kidney MSC lines for further microinjection experiments into wild-type neonates at PND1. This approach was supported by previous findings from our laboratory: 1) k-MSCs were able to undergo epithelial tubular formation in vitro; and 2) microinjection of those cells into the renal parenchyma showed that they ‘home’ to and integrate back into the collecting duct (Li et al., 2015). To assess this, defective k-MSCs were generated through the conditional deletion of both copies of Pkd1 gene (k-MSCsPkd1 del2-4) and then micro-injected back into wild-type neonates at PND1. The Pkd1 gene was selected because several mutations in this gene have been shown to cause autosomal dominant polycystic kidney disease (ADPKD) (Calvet, 1998, Lantinga-van Leeuwen et al., 2004, Lu et al., 1997), and the clinical diagnosis of ADPKD relies mainly on cyst formation throughout all kidney tubular compartments. Therefore, if kidney-derived MSCsPkd1del2-4

Commercial in Confidence functionally integrate into kidney structures, the introduction of these cells into normal mice is proposed to result in the formation of cysts within collecting ducts. This model was also chosen because the genesis of cyst formation in ADPKD suggests that a second hit is required and hence that all cells of a cyst are clonal (Chapin and Caplan, 2010). For this reason, we proposed that even at a 10% level of incorporation we are likely to trigger the phenotype.

2.2.1 Establishment of a double-transgenic mouse line

To establish k-MSCs mutant for Pkd1 gene which were able to be lineage-traced after microinjection, we generated a double-transgenic mouse line using the breeding strategy outlined in Figure 2.8. The main purpose of this breeding is to generate a mouse line carrying the following genes: PkdFlox/Flox and RosaTdTomatoFlox/Flox. The selection of these two genes is based on the following properties:

1) Pkd1: many disruptions on this gene have been widely described to cause Autosomal Dominant Polycystic Kidney Disease (ADPKD). The Pkd1Flox/Flox allele allows us to isolate the cells from an unaffected mouse and generate a Pkd1 deletion in vitro using transfected Cre recombinase.

2) tdTomato+: As the tracking of administered k-MSC in the recipient kidney is critical to the interpretation of our experimental results, the cells were genetically label with tdTomato+.

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Figure 2.8. Breeding strategy for the generation of the double transgenic mouse model: Pkd1Flox/Flox; Rosa26 tdTomatoFlox/Flox. The strategy was organised in two breeding steps. Breeding 1: Homozygous Pkd1 mice were crossed with Rosa26TdTomato mice. Based on Mendelian inheritance, a 50% probability was estimated for getting the heterozygous expected genotype in the filial 1 generation (F1) (Pkd1Flox/ - ;Rosa26tdTomatoFlox/-). To get the homozygous genotype, the Filial 1 generation was then backcrossed with a genetic probability of 12,5% of getting Pkd1Flox/Flox; Rosa26tdTomatoFlox/Flox and Pkd1Flox/Flox.

The Pkd1Flox/Flox mice were bred with Rosa26tdTomato mice. Based on Mendelian inheritance, we estimated that 50% of the filial generation 1 (F1) would acquire the desired genotype. As a result of this crossing, all offspring were heterozygous for the Pkd1 gene (Pkd1+/Flox), so mice from F1 were then backcrossed with an estimated Mendelian inheritance of 12.5%. All the offspring acquired from the described crosses were genotyped by PCR in order to follow the co-inheritance of the transgenes (Figure 2.9).

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A) B)

Figure 2.9: Genotyping of the two key genes: Pkd1 and RosatdTomato. Representative ethidium bromide-stain agarose gel with PCR products amplified from tail DNA isolated from offspring on the second filial generation of Pkd1x Rosa26dTomato mating. PCR reactions were independently carried out for each gene. M denotes the molecular weight marker. A) For Pkd1: the final products were 132 bp for the wild-type allele, and 250 bp for the mutant allele. Samples in line 1- heterozygous genotype; lines 2, 3, 5- wild-type genotype; lines 4 and 6- homozygous genotype. B) For RosatdTomato: the final products were 297 bp for the wild-type allele, and 196 bp for the mutant allele. For this gene, two separate PCR reactions were set up for each animal to detect wild-type and mutant genotype. Samples in lines 1, 4, 7, 8- homozygous mutant genotype; lines 3, 5- heterozygous genotype; and lines 2, 6- wild-type genotypes.

2.2.2 Generation and phenotypic characterization of a primary k-MSC line deficient for the Pkd1 gen (k-MSCsPkd1del2-4)

Whilst generating the double crossing of mice, we proceeded with the derivation of untagged k-MSCs from Pkd1Flox/Flox mice to perform in vitro characterisation. This was important to establish whether k-MSCs mutant for Pkd1 were likely to behave any differently to previously characterised wild-type k-MSCs. In order to produce a primary cell line of k-MSCs mutant for both Pkd1 alleles (k-MSCsPkd1del2-4), kidney MSC-like cells were isolated from a previously reported Pkd1Flox/Flox mouse (Piontek et al., 2004). Kidney-derived MSCs were obtained and expanded under culture conditions, as described in Material and Methods. Once these cells were at passage 5, the conditional inactivation of the Pkd1 gene was carried out by CRE in vitro excision of the floxed sites. To achieve this, k-MSCs were transfected with a plasmid

Commercial in Confidence containing a CRE-IRES-PUROMICIN gene cassette. The expression of the puromycin reporter gene allowed the selection of the successfully transfected cells under antibiotic exposure. A kill curve for the antibiotic was performed to find the optimal concentration of puromycin to kill the un-transfected survival cells (Figure 2.10). Six different concentrations of antibiotic were tested. Concentrations from 4 to 8 ug/ml of puromycin caused an undesirable massive kill of k-MSCs from day 2 onwards, which could have also led to the loss of some successfully recombinant cells. With this in mind, the most suitable concentration was selected as 2 ug/ml. At this dose (approximately 95% of k-MSCs killed at 4 days post-treatment) we were able to cause the gradual death of un-transfected cells without an overkill of the plated cells. Once this was established, k-MSCs were transfected using the described plasmid above (Figure 2.11A). Three different conditions of DNA and lipofectamine were tested, as described in Materials and Methods, followed by puromycin treatment at 2 ug/ml to evaluate the effectiveness of the CRE excision and the antibiotic selection, DNA derived from untreated k-MSCs and CRE- excised clones was analysed by PCR using a pair of primers which aligned outside of both floxed sites (Figure 2.11B), revealing that the three treatments achieved a similarly high, but not complete, inactivation of the Pkd1 gene (Figure 2.11C).

Figure 2.10. Puromycin sensitivity assay for kidney MSC-like cells. Cells were cultured in medium containing 0.5, 1.0, 2.0, 4.0, 6.0, 8.0 ug/ml of puromycin. The survival of the cells at different concentrations was daily examined up to day 4 of culture and estimated in percentages by comparing each plate to the control plate under light microscopy. As control plate we used k-MSCs plated and cultured under normal tissue culture conditions at confluence.

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Figure 2.11. Experimental in vitro approach used to inactivate Pkd1Flox/Flox gene. k- MSCs were plated in a 35 mm petri dish and then transfected using three different treatments of varying Lipofectamine/DNA ratios, detailed in Materials and Methods section. A) Schematic representation of the plasmid used for transfection assays selected in order to optimise CRE and puromycin expression. Recombination of the floxed sites was induced by expression of CRE protein, and then three different treatments were cultured in medium containing 2 ug/ml of puromycin antibiotic for 4 days. B) Diagram of the primers used for gene knock-out evaluation which aligned outside of both floxed sites; a smaller PCR product of ~ 220bp is expected when both alleles are deleted, but a 1.2Kb product is yielded, showing the presence of floxed sites. C) DNA analysis of the effectiveness of CRE excision treatment in the three transfected conditions. DNA from untransfected cells was analysed as control, yielding the expected fragment of 1.2 Kb, whereas DNA extracted from conditions 1, 2, and 3 showed the shorter (220bp) and larger (1.2 Kb) fragments.

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Figure 2.12. Selection of stable clones of k-MSCsPkd1del2-4 from a mixed population . DNA of 44 clones was extracted and then analysed by PCR using a pair of primers to amplify the floxed genomic region. A) Schematic representation of the primers used for clonal screening. A product of 1.2 Kb is yielded before CRE exision and a smaller product of ~ 220 bp shows the deletion of both alleles. B) The region between exon 2 and exon 4 of the Pkd1 gene was amplified for the genomic DNA of 44 clones. PCR products were resolved on a 1.5% agarose gel stained with ethidium bromide, obtaining one null clone of the Pkd1 gene (220 bp) (k-MSCsPkd1del 2-4).

Because none of the three treatments led to a complete Pkd1 inactivation, and to exclude the possibility that wild-type k-MSCs might affect further in vitro characterisation, clonal selection was carried out. Cells were plated at low-density in 100 mm petri dish and clones were then isolated via ring cloning. In total, 44 clones derived by manual isolation were picked and the genomic DNA was analysed by PCR analysis, using the same primer strategy used for mixed cultures (Figure 2.12A). Out of all the clones analysed, just one clone proved to be mutant Pkd1 for both alleles (Pkd1del2-4), identified by detection of the lower PCR product of 220 bp. The 43 remaining clones were wild-type genotypes, identified by a PCR product of 1.2 Kb (Figure 2.12B). A wild-type clone k-MSCWt was then randomly selected to compare with the mutated

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Pkd1 clone (k-MSC Pkd1del2-4) for the following in vitro studies. The establishment of a single- cell clone is important for the subsequent generation of labelled k-MSCs from the double mouse Pkd1Flox/Flox; Rosa26tdTomatoFlox/Flox, which is the ultimate goal.

As mentioned earlier, the Mesenchymal and Tissue Stem Cell Committee of the International Society for Cellular Therapy in 2006 defined the minimum criteria that cells must meet to classify them as MSCs (Dominici et al., 2006). Based on these, the selected wild-type and mutant clones were characterised to validate them as MSCs and to investigate whether the conditional inactivation of the Pkd1 gene had any effects on mesenchymal stromal cell properties. Firstly, all cells isolated from bulk cultures after clonal selection adhered to plastic when maintained in standard culture conditions (Figure 2.13). Secondly, we performed FACS analysis to evaluate selected clones (wild-type and mutated) for the presence of the key surface markers, including CD24, CD44, CD49e, CD51, CD81, CD140a, CD140b, and Sca-1 (Figure 2.14) (Li et al., 2015, Pelekanos et al., 2012). As expected, both the wild-type and the mutant clones were positive for CD44, CD49e, and Sca-1, which have been previously reported as positive for murine immunophenotypic profile (Meirelles Lda and Nardi, 2003). Interestingly, k-MSCsWt (52%) and (93%) k-MSCsPkd1del2-4 clonal populations were positive for a higher percentage of CD24 than the approximately 30% previously reported for bulk cell populations by our laboratory (Li et al., 2015). CD24 has been described as marking tubular progenitor/stem cells, including renal progenitors (Challen et al., 2004, Swetha et al., 2011, Li et al., 2015). Other markers associated with pericyte origin were also evaluated, including CD140a, which was completely negative for both clones, and CD140b, which epitope was positive for only 15 % of mutant clones and 20% of wild-type clones. This data confirms that Pkd1 inactivation does not affect the expression of the characteristic murine MSCs immunophenotypic profile. However, it also indicates some differences in immunophenotype between these clones and the previously isolated k-MSC clones.

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Figure 2.13. Fibroblast-like morphology of clonally derived k-MSCs. The wild-type (k- MSCsWt) and mutated (k-MSCsPkd1del2-4) clones were similarly adherent to plastic and had a fibroblast-shaped morphology under normal tissue culture conditions. Bright-field images were made at low magnification (4X) using an inverted bright-field/fluorescence microscope (Nikon Eclipse Ti-U).

Figure 2.14. Comparative immunophenotype analysis of wild-type and mutated clones of k-MSCs. Cells were originally isolated from bulk cultures and then subjected to CRE in vitro treatment and a clonal selection process. Both populations k-MSCsWt and k- MSCsPkd1del2-4 were analysed at passage 11 for MSC and pericytic cell markers.

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Based on the above in vitro analysis, we can conclude that Pkd1 gene inactivation did not alter the properties of mesenchymal stem cells. Although further in vitro studies need to be performed to obtain a full MSC characterisation, the preliminary data presented here showed that k-MSCPkd1del2-4 met with the defined minimal criteria of an MSC, including adherence to plastic, a fibroblastic morphology, and CD expression profile. In addition, there is down- regulation of epithelial markers after isolation. These data support the feasibility of microinjections of genetically labelled Pkd1 mutant MSCs into kidney compartments. If our hypothesis is correct, the integration of these cells should lead to dysfunctional epithelial growth.

2.2.3 Generation of genetically-tagged Pkd1 mutant k-MSCs for in vitro and in vivo characterisation (k-MSCsPkd1del2-4/TMTO+)

In order to produce a primary cell line of k-MSCs tagged by tdTomato+ and mutant for both Pkd1flox alleles (k-MSCPkd1del2-4/TMTO+), kidney MSC-like cells were isolated from the double transgenic mice Pkd1Flox/ Flox; Rosa26tdTomatoFlox/Flox. Once these cells were at passage 5, the conditional inactivation of the Pkd1 gene was carried out by CRE in vitro excision of the floxed sites, which produced simultaneous activation of the reporter tdTomato+. These cells were then FACS-sorted for tdTomato+, expanded, and the genic inactivation of Pkd1 investigated by PCR (Figure 2.15).

Due to the described ability of k-MSCs to undergo an epithelial mesenchymal transition (EMT) (Li et al., 2015), we next addressed the epithelial capacity of k-MSCsPkd1del2-4/TMTO+. As a control, we used the bulk cell line (k-MSCWt/Wt). The wild-type and the mutant cell lines were then plated for immunofluorescence at passage 12. Both cell lines were positive for the mesenchymal marker vimentin and NG2, while negative for epithelial markers such as Ecad and Aqp2+ (Figure 2.16), which suggests the dedifferentiation of both cells towards a mesenchymal state. Expression of tdTomato+ was evaluated by detection of its native fluorescence.

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Figure 2.15. Validation of a double transgenic cell line k-MSCPkd1del2-4/TMTO+. Kidney derived-MSCs isolated from the double transgenic mice were subjected to simultaneous CRE in vitro excision of floxed sites for Pkd1 and tdTomato+ genes. A) FACS-sorting for tdTomato+ of the double transgenic cell line. B) Live expression of the reporter during cell expansion. C) Flow cytometry of k-MSCsWt/Wt and k-MSCsPkd1del2-4/TMTO+. D) DNA analysis of the Pkd1 gene for the sorted tdTomato+ population. DNA from untransfected cells isolated from Pkd1 mice was used as positive control, which yielded the expected fragment of 1.2Kb, whereas DNA extracted from the sorted tdTomato+ population showed the shorter fragment of 220 Kb.

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Figure 2.16. Evaluation of mesenchymal/epithelial protein markers in the double transgenic lines (k-MSCsPkd1 del2-4 and k-MSCsWt). Both cell lines were at passage 12 at the time of seeding for immunofluorescence. The wild-type and mutant cell lines were positive for mesenchymal vimentin/NG2 markers and negative for the epithelial proteins Ecad/Aqp2. tdTomato+ expression was detected by the native fluorescence of the reporter. Scale bar = 20 µm.

Given that the primary cilia have been widely described as a common denominator in kidney cystic pathogenesis, we evaluated the effect on the primary cilia of the gene deletion between exon 2-4 in the Pkd1 gene. Although there is still little consensus as to whether there is an intrinsic correlation between mutations in polycystin 1 and defects in the primary cilia structure, some studies have also related cyst formation to, not only mutations in either

Commercial in Confidence polycystin protein, but also to cilia defects in mice models (Ma et al., 2013, Jonassen et al., 2008). To assess this, the mutant and control cell lines were plated with equal numbers of cells and, after 24 hours of seeding, ciliation was induced by reducing the amount of serum in the basal medium (serum starvation) for 24 hours (Seeley and Nachury, 2010). After this, immunofluorescence assay was performed against Arl13b. This is a protein localised in the cilia which has been widely described to be essential for maintenance of the ciliary structure. Several mutations in Arl13b have also been defined in ciliopathies (Cantagrel et al., 2008). Hence, the presence of cilia was evaluated in the control and double transgenic cell lines by counting a range of nuclei between 93-106 in ten different z-stacks for each cell line. Cilia were detected in a high percentage of the wild-type cell line (80%), whereas it was significantly reduced (to 37%) in the k-MSCsPkd1del2-4/TMTO+ (Figure 2.17). This result indicates that deletion between exon 2-4 of the Pkd1 gene structurally affected the primary cilia of the MSCsPkd1del2- 4/TMTO+ line. This showed that these cells have great promise for further use in the proposed in vivo experiments.

Figure 2.17. k-MSCsPkd1del2-4/TMTO+ resulted in reduced cell number. A) Representative confocal images of cilia stained with Arl13b in the wild-type (k-MSCWt/Wt), and mutant (k- MSCPkd1del2-4/TMTO+) cell lines (lower left panel) and at high magnification (right panel). B) Quantification of number of cells with cilia. Ten different z-stacks were taken from different regions to count a range of between 93-106 nuclei per cell line. *** P<0.001 significance level from a two-tailed t-test. Scale bar = 20 µm.

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AIM 3. In vivo recapitulation of polycystic kidney as a measure of functional integration into collecting duct.

By the time of performing microinjection experiments for this aim, Leonhard and colleagues had reported the first mouse model which mimicked human ADPKD as a result of a small number (8%) of Pkd1 mutant cells, where evidence of cysts appeared after a long dormancy period of six months (Leonhard, 2015). Based on this study, and on evidence from Aim 1 where microinjections of k-MSCs (GFP+) resulted in an average integration of 7.6%, we anticipated that cell injection of mutant cells into wild-type neonates at PND1 would trigger cyst formation with a long dormancy period of about 6 months.

2.2.3 Microinjection experiments

Having generated and characterized k-MSCsPkd1del2-4/TMTO+, these cells were then used for microinjection purposes, because we hypothesised that if k-MSCs integrate effectively into collecting-duct, the autosomal dominant polycystic kidney disease (ADPKD) phenotype should arise in the recipient wild-type mice. Microinjection experiments were carried out in the same way as described in Aim 1 and in Materials and Methods. Notably, two different mouse strains, Swiss and C57BL/6, were used as recipients of mutant cells for two reasons. Firstly, by the time of performing experiments for Aim 2, our laboratory had relocated from the University of Queensland to the Murdoch Children’s Research Institute, involving the transfer of mice and the adaptation of experiments to a new animal facility. At MCRI, the strain most similar to CD1 (used for Aim1) was outbred Swiss. Secondly, we anticipated that injecting k-MSCsPkd1del2-4/TMTO+ isolated from the C57BL/6 strain might invoke a graft rejection response when injected into a different strain. As this may negatively influence the experimental outcome, C57BL/6L mice were used also as recipients of k-MSCs Pkd1del2-4/TMTO+ to maintain the genetic identicality of donor and acceptor mice.

A total of 24 mice of both genetic backgrounds were microinjected for both genetic backgrounds. This number considered our previous success rate for microinjection experiments as ~ 60-70%. We estimated that at least ~16 mice may proceed to further cyst evaluation. Successful injection was evaluated based on the presence of the fluorescent beads combined with the cells at the time of injection (Figure 2.18A). However, this could only be evaluated for the test controls, not for all animals. Initially, a small group of mice, with three injected

Commercial in Confidence mice from each genetic background, were sacrificed shortly after injection (1 month) to look for Tomato+ cells in the collecting duct epithelium. Only scattered tdTomato+ cells were observed in all the kidneys collected. These were located within the medullary interstitium and cortical tubular structures (Figure 2.18; B, C, D), not the collecting duct, suggesting that these cells had only low levels of integration and had not integrated as previous k-MSC (GFP+) lines had been reported to do. We did not observe any difference related to genetic background. This preliminary evaluation was a major concern for the success of the project, and based on the possibility that we may need a longer time for cyst formation to occur, we further evaluated microinjected mice at six months of age.

Figure 2.18. Evaluation of cell integration of k-MSCsPkd1del2-4/TMTO+ after microinjection experiments. Three injected animals of each genetic background were harvested for assessment of cell integration at 1 month old. A) Schematic representation of the overview of microinjection experiments and steps of assessment. B) Representative panel of a successful microinjected kidney. C), D), and E) Immunofluorescence against Aqp2 and F) laminin and tdTomato+ detection. Scale bar = 20 µm.

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2.2.4 In vivo evaluation

The series of 24 injected mice from four litters were evaluated at six months of age by performing Magnetic resonance imaging (MRI) in eight mice to evaluate cyst formation. Four mice from each genetic background were randomly selected for evaluation of cysts. Only two mice showed some incipient evidence of possible cyst formation (Figure 2.19), although this was not definitive due to the size of the bright spots. Thus, adult injected kidneys were still apparently resistant to the development of overt cystic disease. Only these two animals were then followed for a further 12 months. Surprisingly, at 12 months of age one injected mouse showed robust evidence of cyst formation under MRI scanning, as shown in Figure 2.20. Cyst formation was also confirmed at the time of harvest by macroscopic detection of a cyst formation. The kidney contained fluorescent beads, proving the success of the micro-injection (Figure 2.21).

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Figure 2.19. MRI scanning images of injected mice at 6 months of age. MRI analysis was performed using T2-weighted scans in the injected mice with k-MSCs Pkd1del2- 4/TMTO+. Some evidence of small cysts was found in two animals, highlighted in the yellow dashed circle.

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Figure 2.20. Representative MRI scanning images at 12 months of age. One big cyst structure was detected in one of the two animals evaluated (shown as yellow dashed square). Images represent coronal sections.

Figure 2.21. Bright-field and fluorescent images of the cystic kidney at 12 month of age. A) Cystic kidney of a 12 month injected mice. B) Successfully micro-injected kidney. Fluorescent beads persist a year after the microinjections.

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Despite the fact that one mouse was not sufficient to complete this study to any degree of statistical validity, we hoped to examine and quantify the involvement of injected cells in the cystic tissue. However, detection of the tdTomato+ was not possible using either native fluorescence or antibody detection. The latter proved unfeasible because of background issues in the tissue collected from this species (Figure 2.22). Collectively, these data suggest the mutant generated cells differed in their derivation properties from the original k-MSCs (GFP+) used for the long-term analysis of integration. Nevertheless, we proceeded to examine kidney sections of the harvested mouse. The affected mouse showed a prominent cystic structure in the cortical region which was detected by MRI, but there were some other smaller structures randomly located in the tissue. The cystic index was quantified by calculating the ratio between cystic/total kidney areas in haematoxylin and eosin renal sections, resulting in a percentage ~ 14 % (Figure 2.23). The contralateral kidney was used as control sample and two different sections were used for quantification.

Figure 2.22. Evaluation of the feasibility of immunodetection of k-MSCsPkd1del2-4/TMTO+. An antibody against tdTomato+ was used to detect microinjected cells by immunofluorescence in frozen tissue sections of cystic and contralateral kidneys. A), B) Representative images of immunofluorescence assay against Laminin, tdTomato+, and Dapi. Arrowheads and dashed boxes show clusters of cells on cystic tissue. C) Representative image of the negative control for tdTomato+ using a contralateral kidney section. Scale bar = 100µM.

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A) B)

Figure 2.23. Cyst formation at 12 months post-injection. A) Mice subjected to MRI at 12 months of age were sacrificed and their kidneys stained with haematoxylin and eosin. B) The ratio between total cystic areas and total kidney area was calculated in two different sections. The contralateral kidney was used as control.

Overall, these findings indicate that while derivation of k-MSC lines mutant for the Pkd1 gene was achievable, there was not significant detection of cell integration with the tagged mutant cell line. So, although cyst formation was detected in a single injected mouse maintained until 12 months after injection, we cannot rule out the possibility that this cyst formation was a rare random event.

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2.3 DISCUSSION

The objective of this chapter was to further evaluate the potential use of k-MSCs for the treatment of collecting duct disease and injury, based upon prior evidence that these cells arose from the collecting duct and were able to return there and integrate (Li et al., 2015). To do this, we chose the challenging approach of recreating ADPKD via cell injection of Pkd1 defective cells, to achieve a closer approximation to human ADPKD than has been reported to date.

Here we were able to derive Pkd1 mutant k-MSC lines that showed no apparent defects in the MSC state. Although these lines lack the functional expression of the Pkd1 gene, they fulfil the criteria issued by the International Society for Cellular Therapy for mesenchymal stem cells, including their expression of CD markers and their adherence to plastic. In addition, k-MSCs Pkd1del2-4/TMTO+ showed selective expression of mesenchymal proteins and non-expression of epithelial markers, corroborating their mesenchymal state. On the other hand, cilia count was lower in k-MSCsPkd1del2-4/TMTO+ than in the control line, suggesting that the deletion between exon 2-4 in the Pkd1 gene resulted in a detectable primary cilia abnormality which may have contributed to defects in normal cell polarisation. While it is true that both Polycystin 1 and 2 have been defined as playing a sensory/signalling role rather than an architectural role in the primary cilia, the genetic association between Pkd1 and the cilia structure has not been thoroughly assessed to date.

We then addressed the feasibility of cell integration by performing long-term in vivo experiments. Although we were able to detect evidence in an initial k-MSC (GFP+) line of long-term integration into kidney structures, k-MSCsPkd1del2-4/TMTO+ failed to show evidence of efficient integration. It is possible that inefficiency of the delivery of cells via micro-injection is such that insufficient numbers of mice were examined. However, this also highlights the possible inefficiency of the technique were it to be used for repair. It is also technically problematic given that it is difficult to predict and/or assess the success of an injection without sacrificing the mouse. As a result, it is possible that many of the animals later evaluated by MRI actually had no injected cells active in them. By the time of undertaking this project, there were quite a few mice studies reporting cystic kidney disease related to mutations in either Pkd1 or Pkd2, but all of these studies were based on animal models using much larger numbers of mutant cells than seen in human patients with ADPKD. Surprisingly, in 2015, Leonhard et al., reported an animal model which closely mimicked human ADPKD by reducing the number

Commercial in Confidence of affected homozygous cells to 8%. Despite the similarity of this model with the injection model reported here, it is important to point out some differences. The reported model of induced human ADPKD in mouse was a genetic approach which did not require delivery of exogenous cells. Although this model showed the feasibility of triggering overt cystic formation with scattered Pkd1 mutant cells, it still used a genetic tamoxifen inducible Pkd1 mouse. In contrast, in this study we injected cells which then had to migrate and integrate.

Taking the above study as a reference, injected mice were firstly evaluated at 6 months of age by MRI scanning, showing incipient evidence of cyst formation in only two of eight animals evaluated, both of which were of C57BL/6 genetic background. However, we expected that this cellular approach might produce a milder phenotype and consequently a much longer time interval for triggering cyst formation than in the study. Finally, the two mice which exhibited preliminary signs of cyst formation were analysed at 12 months of age and one showed strong and clear cyst formation under MRI scanning. This was later corroborated by kidney histology sections identifying one big and prominent cyst in the cortical zone and smaller structures in the medulla/papilla region. Despite this positive outcome, we cannot exclude that this was a rare random event. Therefore, we acknowledge that further experiments are required to validate the significance of the evidence of cyst formation outcome.

Overall, based on a systematic evaluation of cell integration as well as of cyst formation we concluded that k-MSCs do not represent a preferable population to functionally integrate to collecting duct epithelium. Although the induction of nephric injury in the recipient mice might assist cell integration, there appears to be no justification for persisting with this approach. However, we did show the capacity to generate k-MSCs with mutant genes which may prove useful as cellular models of human diseases. This ADPKD cell line may be a valuable platform for drug-screening assays.

2.3.1 Challenges with the project / reasons why integration may have failed

As discussed throughout this thesis, the microinjection of k-MSCsPkd1del2-4/TMTO+ did not show capacity of significant integration in the recipient mice being a major concern for this project. Therefore, it is worth to discuss additional technical and logistical challenges which may have also contributed to this negative outcome.

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2.3.1.1 Laboratory relocation

As mentioned above, after the first year of the candidature, our laboratory was relocated from The University of Queensland (Queensland) to the Murdoch Children’s Research Institute (Victoria), which had a big impact on my PhD. This involved allocating time to stop work and reproduce mice colonies for transference, apply for animal ethics approval, and re-standardise ongoing procedures for approval, e.g. microinjection experiments in a new environment. The microinjection approach was originally established using outbred CD1 mice as the recipient strain. To date, the only data generated showing significant cell integration, either in this study or prior studies in the laboratory, was generated using a specific kMSC line introduced into CD1 strain recipients (Li et al., 2015). With relocation, we had to adapt this experiment to use in outbred Swiss mice. Another variation required was the use of C57BL/6 strain as recipient mice because the mutant Pkd1 cell lines also came from C57BL/6. Taken together, it is possible that these variations affected the experimental outcome.

2.3.1.2 Little evidence of integration after initial injections

In this study, integration of tdTomato+ cells was a rare event. Even when we were able to detect tdTomato+ cells in tubular compartments, the efficiency of cell integration was still lower than in previous findings by our laboratory (Li et al., 2015). Given the importance of this outcome to the desired long-term phenotype, this was a major concern. It is possible that the inefficiency of delivery of cells using micro-injection is such that insufficient numbers of mice were examined. However, this is in itself a deficiency of the technique because the only way to evaluate injection efficiency is through sacrificing of mice. The problem is compounded by the laborious nature of the analysis method. Once the kidneys were harvested, their analysis involved cutting thin sections of 10 µm of almost the entire adult kidney to find the region containing fluorescent beads considered to be the region where the injected cells should integrate.

It is also possible that the k-MSCsPkd1del2-4/TMTO+ generated differed in their properties from the original k-MSCs (GFP+). Due to the lack of a proper control line derived with the same reporter (tdTomato+), this was not possible to assess. Therefore, if it was a specific rare event which gave rise to the prior k-MSCs (GFP+), then subsequent k-MSC lines may not have this

Commercial in Confidence capacity. It is not yet clear where variation arises during the derivation process, and hence this cannot be controlled for.

Taking the experimental and technical limitations discussed above together, along with the fact that MRI scanning of eight mice at the age of six months only showed preliminary evidence of cyst formation in two mice, cyst evaluation was only completed in these two animals. This was an insufficient number to reach any clear conclusions.

2.3.2 Future directions

A number of additional aims had been proposed for the project, all of which were contingent upon the outcome of this chapter. These included ‘proof of concept’ for treatment of collecting duct disease using a mice model of diabetes insipidus. Indeed, substantial work was performed towards this rescue aim that has not been included in this thesis. In short, while there was clear evidence of a disease phenotype in the analysed Aqp2 mutant mice, no amelioration of the disease occurred as a result of the introduction of wildtype k-MSCs (GFP+). The lack of positive data from our work towards these aims was a major concern, and the project shifted from examining the utility of k-MSCs to recreating collecting duct epithelium from human pluripotent stem cells. This work is discussed in Chapters Three and Four.

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CHAPTER III

Evaluation of culture methods, cell lines, and optimal growth factors required for improving patterning to ureteric epithelium

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3.1 INTRODUCTION

In this thesis, the focus has been on the recreation of one of the two progenitor populations which give rise to the kidney referred as ureteric bud (UB). These cells form the branching collecting ducts of the mature kidney through which the urine leaves the organ. This work is covered in Chapters Three and Four, with this Chapter focusing on developing tools and a protocol for generating ureteric bud, and Chapter Four covers the refinement of the process and the evaluation of resulting tissue. Critically, the development of this protocol relied upon our comprehensive understanding of the ureteric bud lineage derivation, cell-cell interactions, and markers of specific components of cell types involved in mouse kidney development. In Chapter One, we reviewed the basis of kidney organogenesis, focusing on the metanephric kidney. Here, we will comprehensively examine the molecular mechanisms and signalling pathways involved in the derivation of ureteric bud lineage and the branching morphogenesis process.

3.1.1 The main regulatory signalling pathway of ureteric bud formation: GDNF-RET

3.1.1.1 GDNF/GFRα1/RET complex and relevance of RET expression during UB outgrowth

Recent advances in differential gene expression studies, lineage tracing, and in vitro models of the branching morphogenesis process have revealed the role of several molecules acting as positive and/or negative regulators of the arborisation process. Extensive literature refers to GDNF/RET signalling as the key master pathway to induce the initial budding of the nephric duct (ND), the primitive structure (mesonephric kidney) which precedes UB formation, as well as promoting continued branching morphogenesis throughout development (Blake et al 2014, Costantini and Kopan, 2010, Little and MacMahon 2012). However, the precise molecular mechanism which occurs remains to be elicited. Indeed, mice lacking either Gdnf or Ret exhibit failure of ureteric bud formation (Moore et al., 1996, Pichel et al., 1996, Sanchez et al., 1996, Schuchardt et al., 1994). In humans, 5-30% of the patients suffering from CAKUT have a RET mutation (Davis et al., 2014), and 5-10% have a GDNF mutation (Costantini, 2006). In the early stages of development, RET (receptor tyrosine kinase) is expressed throughout the nephric duct, however its expression becomes gradually restricted to the tip population over the course of metanephric development. It was initially proposed that UB outgrowth was

Commercial in Confidence caused by an increase in cell proliferation (Michael et al., 2004), but years later a study showed that this event was rather related to cell movements (Chi et al., 2009). In particular it showed that, upon localized GDNF stimulation, nephric duct cells enter into competition for their inclusion in the nascent UB outgrowth. Here, cells with high RET activity were able to move towards the swelling region of the ND, giving rise to the ureteric bud (UB) and leaving behind cells with null or low RET activity which populate the nephric duct (Figure 3.1). Finally, cells with high RET activity forming the UB remained restricted to the tip region (Chi et al., 2009). This study conclusively proved that UB formation is driven by cell competition based on RET activity.

Currently, four ligands have been described to activate RET (Airaksinen et al., 1999). Glial cell line derived neurotropic factor (GDNF) is the only active ligand in kidney organogenesis. GDNF is a distant relative of the transforming growth factor (TGF)-β superfamily (Lin et al., 1993). During early stages of development, this growth factor is expressed by cells in the nephrogenic cord (Costantini et al., 2006) is then seen in the MM (Dressler., 2009) and has been reportedly express in the nephron progenitors arising from the MM (McMahon, 2016). Recent single cell profiling has identified GDNF expression in cortical stromal cells as well as the nephron progenitors (Magella et al., 2017). Fate mapping studies using inducible mice (Gdnf-CreERT) confirmed GDNF expression as restricted to the nephron progenitor population throughout kidney development (Sanchez et al., 1996, Cebrian et al., 2014, Hellmich et al., 1996). It binds to its co-receptor GFRα1, promoting auto-phosphorylation of RET receptors forming the molecular signalling complex GDNF- GFRα1-RET (known also as “RET signalling”) (Durbec et al., 1996, Treanor et al., 1996, Trupp et al., 1996). However, RET signalling is also dependent on the availability of heparin sulfate glycosaminoglycans (Barnett et al., 2003, Sariola et al., 2003, Shah et al., 2011). Calcium binding is also required for RET ligand activation (Anders et al., 2001, Ibanez 2013). Upon RET signalling activation, a series of intracellular signalling pathways, including ERK MAPK, PI3Kinase/Akt, PLCγ, and others, is triggered downstream (Costantini, 2006, Costantini and Kopan, 2010).

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Figure 3.1. Illustration of Ret+cell movements from nephric duct to UB formation. Cellular representations: Ret positive cells (blue) and Ret negative (green). A) Ret positive cells are dispersed along the nephric duct. Cell movements start to form the first swelling of the UB (yellow arrows). B) Competition between cells with high, low, or null Ret expression to reach the UB outgrowth. C) Cells with high Ret activity (blue) lead primary ureteric bud formation. (Figure extracted from Chi et al., 2009).

3.1.1.2 Regulation of Ret and Gdnf expression

Several genes have been identified in promoting and maintaining Ret and Gdnf expression during metanephric development. Some of them are upstream genes of the Ret signalling pathway which also contribute during nephric duct (ND) elongation and UB swelling. In this context, one of the key regulators during kidney organogenesis is the zinc-finger transcription factor GATA3, which is not only described as playing a crucial role in controlling ND morphogenesis (Grote et al., 2006), but also is an essential regulator of Ret expression over the course of organogenesis (Grote et al., 2008, Boualia 2011). However, Ret expression is also regulated by the presence of other genes, including Pax2, Pax8, Lhx1, and Ctnnb1 (encoding β-catenin) (Grote et al., 2006, Grote et al., 2008, Boualia et al., 2011, Marcotte et al., 2014, 70- Chatterjee et al., 2012, Jeanpierre et al., 2012, Bouchard et al., 2002.), which form an essential regulatory core (Pax2/Pax8/Gata3/Lhx1) during nephric duct development (Marcotte et al., 2014). Gata3 null mice exhibit premature extinction of Ret expression, abnormal ureteric bud budding, and subsequently aberrant kidney formation (Grote et al., 2008). Another important regulator of Ret expression is retinoic acid, secreted by stromal cells (Lokmane et al., 2010, Moreau et al., 1998), which will be reviewed in this chapter in more detail. In addition to upstream regulators, there are also downstream regulators, such as Wnt11, which are widely described as sustaining a positive feedback loop governing Ret expression (Majumdar et al., 2003). On the other hand, less is known about the transcriptional regulation of Gdnf. Mice

Commercial in Confidence models have helped to reveal some important regulatory genes, including Pax2, Six1, Eya1, and Hox11 paralogs, which are all genes expressed in the MM (Marcotte et al., 2014, Lokmane 2010, Boyle et al., 2006; Batourina et al., 2001, Schedl et al 2007). Moreover, over the course of metanephric development, Wnt11 not only maintains a positive loop modulating Ret expression but also signals to the MM to sustain Gdnf expression (Majumdar et al., 2003, Basson et al., 2006).

3.1.2 The role of retinoids in modulating Ret expression in the developing kidney

As previously discussed, proper development of the kidney requires Ret expression, which is not only dependent on GDNF+ but also on retinoic acid (RA). An extensive literature exists on the essential role of vitamin A and its metabolites (RAs) in the formation of several tissues and organs, including the kidney. Several mice studies have reported that vitamin A deficiency is associated with impaired branching morphogenesis and a subsequent reduction in nephron numbers (Lelievre-Pegorier et al., 1998, Moreau et al., 1998). The isomer All-trans retinoic acid (tRA) is a naturally occurring derivative of vitamin A and is also widely documented as a component in kidney embryogenesis (Ross et al., 1993, De Luc A, 1993, Means and Gud.AS, 1995). tRA signals via nuclear transcription factors belonging to the retinoic acid receptor family (RAR), comprised of eight major isoforms (RARα1 and α2, RARβ1, β2, β3 and β4, and Rγ1 and γ2). These isoforms are differentially expressed during embryonic development and display evolutionary conservation (Dolle et al., 1990, Giguere et al., 1990, Krust et al., 1989, Leroy et al., 1991, Zelent et al., 1991). 9-cis retinoic acid (9-cisRA) is another isomer of RA, which not only binds with high affinity to RAR but also to the retinoid X receptor (RXR) family (Levin et al., 1992, Wolf, 1991). Endogenous activity of this isomer was first reported in kidney and liver (Heyman et al., 1992), and later in other tissues. Since retinoic acid signalling plays an important role in patterning towards ureteric bud lineage, 9-cis retinoic acid is used in combination with tRA in most of the studies attempting in vitro propagation of the isolated ureteric epithelium or kidney explants when investigating Ret expression (Rosselot et al., 2010, Batourina1 et al., 2001). Overall, once retinoids reach the nucleus they promote heterodimerization of their receptors, RAR/RXR, binding to enhancer elements (RARE) and activating transcription of responsive genes (Figure 3.2) (Chambon, et al., 1996, Mangelsdorf et al., 1995).

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Figure 3.2. Retinoic acid signalling pathway. Retinoic acid (RA) can be intracellularly synthesized or diffused from an adjacent cell (red arrow). The former is assisted by cellular retinol-binding proteins (CRBP) which present retinol to retinol dehydrogenases (RDHs). Where RA is intracellularly available cellular retinoic acid-binding proteins (CRABPs) assist the translocation of RA to the nucleus. In the nucleus, RA promotes heterodimerization of the receptors RAR/RXR which are able to bind to enhancer elements (RARE), activating transcription of responsive genes by assembling the complex composed of RNA polymerase II (Pol II), TATA-binding protein (TBO), and TBP-associated factors (TAFs). (Figure extracted from Rhinn et al., 2012).

In 1999, Mendelsohn and colleagues examined the compound mutant mice lacking RARαβ2, which phenotype resulted in small malformed kidneys and reduced numbers of nephrons with abnormal branching morphogenesis, demonstrating the essential and coordinated action of RARα and RARβ2 in response to retinoic acid signalling. Notably, the expression of these two RAR receptors was restricted to stromal cells. Gene analysis expression revealed down- regulation of the proto-oncogene c-ret whose expression is spatially separated from RAR- containing cells. Based on this, the authors postulated the existence of a sort of signal release, by RAR-containing cells, which stimulates the expression of c-ret in the ureteric bud population. Interestingly, the same authors later observed the rescue of all the phenotypic abnormalities in mice lacking RARαβ2 when the Ret gene was constitutively expressed in ureteric bud cells (Batourina et al., 2001). Both of the above studies suggested the major role of retinoic acid is in regulating Ret expression and they identified the stromal cell as the primary site of retinoid action.

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For almost ten years, the consensus was that RARE activity was confined to stromal cells, but later reports described retinoic acid activity in ureteric bud and collecting duct cells, in embryonic and adult mice respectively. In 2010, Rosselot et al. described tRA-RAR-activity in ureteric bud cells of embryonic kidneys. This finding was later reinforced by another group using RARE-hsp68-lacZ transgenic mice, which also identified endogenous retinoic acid action in principal and intercalated cells, which are the two mature cell types of the collecting duct system. This seemed to change the primordial role of stromal cells (Wong et al., 2011). However, these studies did not resolve the question of whether synthesis and metabolism of RA in ureteric bud cells was involved in supporting renal development.

To address this question, deficient mouse models of retinol dehydrogenase enzymes Raldh2 (specific to cortical stroma) and Raldh3 (specific to UB cells) were examined (Rosselot et al., 2010). Strikingly, Raldh3 had a minor effect in organogenesis, whereas mutation of Raldh2 resulted in similar phenotypes to those described for RARαβ2 double mutant mice. As expected, the double compound mutant Raldh2-3 showed a more aggressive phenotype. These findings clearly proved that Raldh2 has a major role in kidney development, and re-established the essential role occupied by stromal cells in maintaining stromal-UB communication during ureteric bud remodelling. Moreover, the observation that GDNF+ and a combination of both isomers of RA are sufficient to trigger Ret in isolated cultivated UBs ruled out the participation of other signals secreted by the stroma (FIGURE 3.3).

Figure 3.3. Retinoids (cis and trans) are sufficient to maintain Ret in isolated ureteric buds. A) Isolated UB at E.11 prior to culture. B) UBs were maintained in culture in serum-free conditions with GDNF + RAs (cis and trans) for 2 days. C) Ret expression was detected by in situ hybridization in samples cultivated with GDNF + RAs (cis and trans). D) Isolated UBs cultivated with GDNF+ and without RAs (cis and trans). E) Ret expression was undetectable in wholemount E.11 ureteric bud maintained for two days with GDNF and without RAs. (Extracted from Rosselot et al., 2010)

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While there is not much information about retinoic acid responsive genes, and even less is known about it at the mechanical level, some studies have recently appeared. Retinoic acid responsive genes are related to many developmental process. In the case of the kidney, they are mainly related to biological functions of the collecting duct, such as regulation of oxygen levels, and steroid hormones (Wong et al., 2012). In 2014, Nishida and colleagues identified 33 candidate genes, using kidney explants exposed to RAs and antagonists. The expression of some genes, such as Elf5 and Scnn1b (sodium channel) in the ureteric trunk, were dependent on retinoic acid addition. Collectively, these findings showed that retinoids are not only crucial for turning on the expression of the master gene Ret, but they also trigger the unique expression of other genes restricted to the ureteric bud linage.

3.1.3 Intracellular signalling pathways and downstream regulators of RET

RET signalling is implicated in the activation of several intracellular pathways, including Ras- Erk MAP kinase, PLCγ/Ca+, PI3K-Akt pathways, and others (Takahashi, 2001)). Transgenic mouse models varying either RET tyrosine residues or other key genes, and organ culture studies using activators/inhibitors, have shed some light on the molecular pathways and gene networks involved in the branching morphogenesis process throughout development (Figure 3.4). For instance, blocking Erk Map-kinase-kinase (MEK) did not have a major apparent effect on elongation, but caused significantly slower branching morphogenesis in kidney cultures (Watanabe and Costantini, 2004, 102- Fisher et al., 2001). Moreover, the later loss of MAPK activity in a genetic model led to not only fewer branches but also to several defects in cell-cell and cell-matrix adhesion (Ihermann-Hella et al., 2014). Mutation in the residue of tyrosine 1015 in the Ret gene also provoked inhibition of PLCγ activity, leading to multiple ureters and smaller kidneys (Wong et al., 2005, Jain et al., 2006). On the other hand, PI3K inhibition impaired UB outgrowth and subsequently halted extension of the epithelium (Tang et al., 2002). However, deletion of the enzyme PI3K phosphatase (PTEN), a PI3K antagonist, caused abnormal cell migration and irregular branching, suggesting that PI3K has a role in cell motility during UB development (Kim and Dressler, 2007). Despite our knowledge of the signalling pathways having grown tremendously in the past few decades- revealing that the main biological processes involved in UB morphogenesis are cell proliferation, survival, motility, and migration- there is not yet a complete understanding of how the above molecular events trigger or affect the arborisation process in the developing kidney.

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The identification of downstream regulators of Ret signalling became of interest to many researchers (Figure 3.4). In 2009, Lu and colleagues performed a screening for genes upregulated by Ret signalling on isolated ureteric buds, cultured with and without Gdnf, even though Wnt11, Spry1 and Ret had already been described as up-regulated by Gdnf. Their study identified novel regulators such as Cxcr4 (chemokine receptor), Crlf1 (cytokine), Myb, Etv4 and Etv5 (transcription factors), and Dusp6 and Spread2 (both inhibitors) (Lu et al., 2009). Although a clear role was not defined for Myb, Dusp6 and Spread2, the study undoubtedly made a great contribution by identifying Etv4 and Etv5 as key downstream molecules of Ret signalling in kidney development. During the same year, another report described the role of Cxcr4 in UB branching and nephrogenesis after an in vitro assay (Ueland et al., 2009). This study identified its role in kidney cell migration, as has been previously reported in other developing cell systems and the immune system (Murdoch, 2000, Schier, 2003).

Since then, numerous efforts have been made to elucidate the roles of Etv4 (Pea3) and Etv5 (Erm), which belong to the Pea3 family of ETS transcription factors. They were both initially documented to be involved in spermatogonial, neuronal, and limb cell growth and then later in kidney organogenesis (Livet et al., 2002, Chen et al., 2005, Brent and Tabin, 2004). Etv4 and Etv5 expression requires Ret signalling via PI3K (Figure 3.4) (Lu et al., 2009), and they are co- expressed in the ND and UB tip-zone, although weaker expression has also been detected in MM and nascent nephrons. Mice lacking Etv4 or Etv5 have a mostly normal renal phenotype. Conversely, the double compound mutant (Etv4-/-;Etv5+/-) led to a damaged renal phenotype, exhibiting either renal agenesis (lack of kidneys) or abnormally small and malformed (dysplastic) kidneys, and absence of a nephrogenic zone (Lu et al., 2009). This data conclusively identified these two ETS transcription factors as key downstream regulators of receptor tyrosine kinases which collectively support branching morphogenesis.

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Figure 3.4. Gene networks involved in nephric duct (ND) and ureteric bud (UB) morphogenesis. A) Genetic networks of nephric duct development. B) Gene networks downstream of tyrosine kinase receptors (RET, FGF2 and MET) involved in UB development. C) Representation of interaction between metanephric mesenchyme (MM) and ureteric bud (UB) (Adapted from Costantini and Kopan, 2010).

Sox8 and Sox9 are also important transcription factors which act together as downstream regulators of Ret. Similarly, the examination of the double knock-out mice Sox8/Sox9 revealed renal agenesis and abnormal branching morphogenesis (Reginesi et al., 2011). Furthermore, it was shown that Ret expression was unaltered, and Sox9 expression was not dependent on RET activity. However, downstream targets, including Etv4, Etv5, Wnt11, Cxcr4, Dusp6, Met, and Spry1, resulted in reduced or null expression (Reginesi et al., 2011). Taken together, the activation of the GDNF/Ret signalling triggers several intracellular pathways and gene networks, some of which are critically regulated by the co-action of downstream transcription factors such as Etv4/Etv5 and/or Sox8/Sox9. Hence, orchestrated action of upstream and downstream regulators sustains a molecular network which is not only essential to initiate epithelial branching morphogenesis, but also to modulate the biological process underlying kidney organogenesis.

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3.1.4 Other signalling pathways regulating ureteric bud morphogenesis

1. WNT/β-catenin signalling: WNT/β-catenin is a well-established signalling pathway in nephron formation, but its role in UB formation has only been documented recently. The tissue- specific loss of β-catenin in mice leads to failure in UB development, with diminishing expression of hierarchic ureteric bud genes including Pax2, Lim, Ret, and Wnt11 (Bridgewater et al., 2011). Indeed, this pathway was also reported role in the maintenance of the progenitor population in undifferentiated state (Marose et al., 2008). Nevertheless, apart from acting as a positive regulator, the WNT/β-catenin pathway has also been reported as an inhibitor of the branching process. Examination of embryos deficient in ICAT, a blocker of WNT/β-catenin signalling, revealed a retarded and abnormal UB branching process (Hasegawa et al., 2007). In support of this, Bridgewater et al. (2011) reported failure in the branching process in a mice model constitutively expressing β-catenin associated with upregulation of the negative regulator Tgfb2. Although the WNT/β-catenin pathway acts as a dual regulator favouring/inhibiting extension of the ureteric bud epithelium and there are still many unanswered questions at the mechanical level, there is currently consensus that it is necessary for the gene network to be modulated by canonical WNT/β-catenin during UB morphogenesis.

2. FGF signalling: The signalling system formed by fibroblast growth factors (FGF) and their receptors (FGR) is widely reported to play an essential role supporting cell survival in kidney organogenesis (Qiao et al., 1999). Among them, FGF7 (Qiao et al., 1999), FGF10 (Michos et al., 2010, Qiao et al., 2001), and FGFR2 (Bates, 2011, Zhao et al., 2004) have been reported as essential components in ureteric bud formation. In fact, FGFs also exert an effect in modulating the expression of important genes related to branching morphogenesis, such as Ret and Wnt11 (Bates, 2011). Moreover, FGF10 knock-out mice models exhibited mild effects across the renal phenotype. Abnormalities in branching morphogenesis were detected when the Ret signalling pathway was reduced or blocked (Michos et al., 2010), suggesting that this is another important contributor to UB epithelial extension.

3. WNT11 signalling: As mentioned earlier, Wnt11 is a tip+ specific protein whose expression is dependent on Ret activity (Majumdar et al., 2003). It is a downstream factor of Ret signalling, forming a positive expression loop with the GDNF-RET complex (Lu et al., 2009, Majumdar et al., 2003). Wnt11 mutant mice exhibit severe renal abnormalities related to branching dysfunction (Majumdar et al., 2003). Wnt11 has also been shown to be capable of signalling to

Commercial in Confidence the metanephric mesenchyme where it positively modulates GDNF expression (Majumdamar et al., 2003), possibly via activation of Frizzled 4/8 (FZD4/8) receptors in UB (Ye et al., 2011).

4. TGF-β signalling: Another factor which plays a role in the formation of different ductal organs is transforming growth factor-β (TGF-β). In the case of the kidney, TGF-β signalling has however been described as an inhibitor of branching morphogenesis. In 2007, Michos and colleagues reported a reduction in BMP4 (a member of the TGF-β family) levels through gremlin1 (GREM1) impact positively on UB branching morphogenesis (Michos et al., 2007). In addition, kidney explants treated with TGF-β1 blocked ductal formation and reduced nephron numbers (Clark et al 2001). The same outcome was observed with Tgfb2 (Sims-Lucas et al., 2008) and alk3 (BMP receptors) (Hartwig et al., 2008) in studies characterising mutant mice models. While little is known about the mechanism underlying TGF-β, it is undoubtedly a relevant inhibitor factor in the morphogenesis of branched epithelia.

5. Other activation mechanisms apart from GDNF-RET: Although there is still not much knowledge at the mechanical level, some evidence has also been collected which identifies other signalling pathways than GDNF-RET as modulating formation of the UB epithelium. As earlier mentioned, FGF10 signals via the tyrosine kinase receptor FGFR2 and regulates ductal epithelial formation (Michos et al., 2010), just as the hepatocyte growth factor (HGF) does via its receptor MET proto-oncogene (Ishibe et al., 2009, Woolf et al., 1995). The latter has also been reported to jointly act with the epidermal growth factor (EGF)-EGFR, mediating not only branching morphogenesis but also maintenance of the collecting duct system (Ishibe et al., 2009). More recently a tissue-specific mutant mouse of Kif3, a primary cilium protein, exhibited signalling via GLI3 which caused a surprising diminution of Ret and Wnt11 expression and an abnormal renal phenotype (Chi et al., 2013), suggesting Kif3- GLI3 as an interesting new mechanism to be further investigated.

3.1.5 Generation of kidney cell types from induced pluripotent stem cells (iPSCs)

The development of protocols for generating kidney structures has relied significantly upon our comprehensive understanding of kidney development. To date, several stem-cell-based approaches have demonstrated the generation of different kidney cell types (Taguchi et al., 2014, Mae et al., 2013, Xia et al., 2013, Morizane et al, 2015, Freedman et al., 2015). In 2013, Mae et al., using a fluorescently-tagged OSR1 iPSC line differentiated with activin A, bone

Commercial in Confidence morphogenetic protein (BMP7), and CHIR99021 co-cultured with embryonic mouse tissue, reported evidence that these cells could differentiate into podocytes and renal tubule epithelia. A year later, Lam and colleagues reported the differentiation of human ESCs to intermediate mesoderm (PAX2+LHX1+), using CHIR99021 during the first days, followed by FGF2 and retinoic acid. These cells could furthermore form LTL+ CDH16+ tubular epithelium, which showed capacity to integrate into kidney explants. Similarly, in 2013, Xia et al. first reported the technique for driving hPSCs toward UB progenitor–like cells lineages with chemically defined media. Here, monolayer cultures of human iPSCs and ESCs were induced with two days in BMP4 and FGF2, followed by two days in retinoic acid, activin A, and BMP2. The sole expression of HOXB7+, RET+, and GFRα1+ was shown as proof of ureteric bud fate. This study therefore did not perform an exhaustive examination of the differentiation cultures, examining the expression of a very limited set of genes. It also lacked evidence for UB protein markers, and formation of epithelia structures both regarded as ureteric epithelium. While differentiated cells were subjected to re-aggregation cultures, and these displayed the ability to integrate into embryonic UB structures, this approach also failed to provide evidence of any functional signs of ureteric epithelium.

Much effort has also been devoted to the generation of the nephron progenitor population (NPs). As discussed in Chapter One, NPs derive from the metanephric mesenchyme located in the periphery of the embryonic kidney, where they interact with the UB to give rise to the nephron. Recently, several researchers have reported the generation of segmenting nephrons. In 2014, Taguchi et al. developed the first approach and demonstrated the derivation of iPSCs to SIX2+WT1+SALL1+PAX2+ metanephric mesenchyme. Differentiated cells also showed evidence of three-dimensional models of nephrons containing both glomeruli and renal tubules. A more recent study, using co-culture of iPSCs in Matrigel (Corning, NY) and activation of Wnt canonical signalling, induced the formation of epiblast-like spheres which also resulted in segmenting nephrons (Freedman et al., 2015). Although these were all important advances, none of them described the formation of the entire organ.

In 2015, research by our group developed a method to turn human iPSCs into small models of the developing human kidney referred to as ‘organoids’. Again, this involved the stepwise addition of a series of growth factors and chemicals to direct iPSCs to kidney fate, resulting in kidney organoids which simultaneously form structures containing ureteric, nephron, vascular, and stromal progenitors (Figure 3.5) (Takasato et al., 2016). Briefly, this protocol initiates the

Commercial in Confidence differentiation of iPSCs toward intermediate mesoderm by activating Wnt-signalling during four days of treatment with CHIR99021, followed by three days with FGF9 and heparin (HA). At Day 7, the two progenitor populations are simultaneously induced, the cells are used to form aggregates which are pulsed with one hour of CHIR99021 to induce nephrogenesis, and then they are cultured with FGF9 and HA for the next five days (Figure 3.5). After this period, organoids are only cultured in the basal medium APEL. After aggregation culture up to day 25, each organoid contains up to ten nephrons, each displaying evidence of proximal distal segmentation. Notably, these nephrons even showed evidence of maturation, by podocytes foot-process formation, and capacity of albumin uptake in proximal tubular cells.

Figure 3.5. Directed differentiation protocol for the generation of kidney organoids from hPSCs. Cells are exposed to CHIR99021 (8µM) for four days to induce the formation of the posterior primitive streak, before switching to FGF9 from days 4–7 to induce the formation of intermediate mesoderm. At day 7 of differentiation, cells are aggregated to form organoids cultured onto a transwell filter. After aggregation, organoids are briefly exposed to CHIR9902 pulse before culturing in FGF9 + HA until day 7+5. After this period, growth factors are removed, which enables the kidney organoid to self-organise into each of the various renal subcompartments (Adapted from Takasato et al., 2016).

In summary, our capacity to generate kidney cell types has grown enormously in the last five years, and the generation of kidney organoids by our laboratory was one of the major breakthroughs in the kidney field in 2015. Indeed, the generation of both nephrons and kidney organoids has great promise for therapeutic uses, for example in nephrotoxicity screening, disease modelling, and drug screening. While many of these studies have already had successful results (Narayanan et al., 2013, Li et al., 2014, Freedman et al., 2015), we have become aware that our recently reported technique for producing kidney organoids is not transferable to therapeutic use because of the lack of a sustained renal progenitor population

Commercial in Confidence and the relatively short time period for culture. To overcome this limitation, we need to be able to produce large quantities of high-quality kidney cell types. To this end, one of the purposes of this thesis is the recreation of one of the two progenitor populations that give rise to the kidney, the ureteric bud epithelium population. The recreation of these two progenitor populations will also allow the investigation of human kidney embryogenesis, answering important questions about nephron formation in humans. Furthermore, it will be the basis of the generation of large scale cultures of progenitors, making this technology transferable to tissue engineering and a vast number of therapeutic uses.

3.1.6 Hypothesis and aims

Hypothesis: Directed differentiation of iPSCs towards nephric duct/ureteric bud lineage by patterning cells towards anterior intermediate mesoderm followed by addition of GDNF and retinoids.

AIM 1: Investigating the effect of GDNF and RA signalling on the ureteric branching morphogenesis process.

AIM 2: Validating the use of new collecting duct reporter cell lines on a feeder layer-free system.

AIM 3: Investigating the action of chemicals, media, and growth factors upon nephric duct/ureteric bud patterning.

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3.2 RESULTS

AIM 1: Investigating the effect of GDNF and RA signalling on ureteric branching morphogenesis process.

3.2.1 GDNF and retinoic acid promote branching morphogenesis in kidney explants

A long literature exists proposing that retinoic acid (RA) and GDNF are the two most important components needed to trigger the expression of RET receptors (Qiao et al., 1999, Rosselot et al., 2010, Riccio and Michos, 2012). The presence of the tyrosine kinase receptor is an important marker of the ureteric bud and it is also required for subsequent branching morphogenesis. In order to evaluate the action of GDNF and RA in UB branching we initially utilized an organ culture system. While this system does not provide a completely accurate model of kidney development, the metanephric kidney grows remarkably well at the air- medium interface and organ culture is the easiest and most reliable way to test the action of grow factors and small molecules during in vitro organogenesis. We thus then began using organoids generated from induced human pluripotent stem cells (iPSCs) to establish a new protocol which favours the expression of critical genes which are expressed in the ureteric bud epithelium.

Kidney rudiments were dissected from 11.5dpc HOXB7-EGFP embryos. The expression pattern of this transgene has been widely described as present throughout the renal collecting duct (Srinivas et al., 1999). HOXB7-EGFP kidneys were selected to facilitate the visualisation and monitoring of the ureteric bud epithelium in vitro. Embryonic kidneys were placed on Transwell filters and cultured for 96 hours in serum-free medium (DMEM/F12) to test the action of GDNF and AGN. Three different culture conditions were evaluated to test the activity of GDNF -basal medium, 50, and 150 μg/ul of GDNF. All three successfully promoted arborisation from a single branched structure resulting in extensive proliferation with several branches of ureteric bud due to the existence of the MM and UB within kidney explants (Figure 3.6). However, swollen tips were exclusively detected in the presence of GDNF, indicating the activity of this component with either both concentrations for its further use in stablishing culture conditions with iPSCs. As anticipated, treatment with AGN193109, an efficient antagonist of retinoic acid action (Chapla et al., 1996), disrupted branching morphogenesis, even in the presence of GDNF (Figure 3.7). The results of these organ culture studies were in

Commercial in Confidence line with previous reports which showed the relevance of both signalling pathways in triggering successful arborisation of the ureteric bud epithelium (Qiao et al., 1999, Rosselot et al., 2010, Odysee et al., 2012), and they also validated the effectiveness of these reagents for further studies.

Figure 3.6. Evaluation of the activity of GDNF in kidney explants from E.11.5 HOXB7-GFP mice. Kidney explants were placed in three different culture conditions: only basal medium (BM), basal medium + 50 ng/ml GDNF, and basal medium + 150 ng/ml GDNF. Branching morphogenesis was examined by following GFP+ expression every 24 hours for four days.

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Figure 3.7. AGN193109 ceases arborisation in kidney explants from E.11.5 HOXB7- GFP mice. The presence of GDNF could not rescue the effect of the retinoid antagonist in culture. Kidney explants were placed in four different culture conditions: only basal medium (BM), basal medium+2.5 µM AGN, basal medium+2.5 µM AGN +50 ng/ml GDNF, and basal medium+2.5 µM AGN +150 ng/ml GDNF. Branching morphogenesis was examined in these cultures by following GFP+ expression every 24 hours for four days.

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3.2.2. Evaluation of GDNF and All-trans retinoic acid (tRA) in kidney organoids

Based on our understanding of embryogenesis, we believe that the anterior intermediate mesoderm gives rise to the nephric duct/UB/collecting duct lineage (Takasato and Little, 2015). This epithelium is positive by immunofluorescence for PAX2, GATA3, DBA and CDH1. Using this knowledge as a base, we then addressed the question of whether addition of GDNF and All-trans retinoic acid (tRA) to the differentiation cultures would induce expression of key ureteric bud genes. We used as reference the protocol reported by our laboratory for generating kidney organoids, reviewed in detail in the introduction section of this chapter (Takasato et al., 2016). While this approach showed evidence of tubular segments positive for PAX2+ECAD+GATA3+, assumed to be collecting duct, there was neither evidence of a complex epithelium nor expression of UB markers such as RET+ and WNT9B+ (Takasato et al., 2016). Therefore, the major objective of this chapter is to appropriately specify nephric duct/ureteric epithelium expressing RET (UB/tip+). While the ureteric epithelium as a whole gives rise to the collecting ducts, it is cells of the branching tips which drive both self-renewal of NPs and the generation of new ureteric branches, without which the organ will not reach optimal size and structure (Costantini and Kopan, 2010)

As a first approach, the wild-type human iPSC cell line CRL1502 (clone C32) was differentiated via a shorter period and lower concentration of CHIR99021 (6µM), based on previous evidence that shorter incubation with the glycogen synthase kinase GSK-3 inhibitor, CHIR99021, promotes a more prominent induction to the anterior intermediate mesoderm from which ureteric epithelium arises (Takasato et al., 2016). After two days of culture, cells were then treated with All-trans retinoic acid (0.1µM) (tRA), FGF9 (200 ng/ml), and heparin (HA) (1 µg/ml) for the next five days. At day 7, cells were used to make aggregates termed ‘organoids’, as a way to promote cellular organization and better epithelisation. Because this approach did not intend to trigger nephrogenesis, organoids were not treated with 1 hour of CHIR pulse. Based on our previous data from organ culture, aggregates were then cultured under the two following conditions from day 7 onwards- condition one: GDNF (50 ng/µl) + tRA (0.1µM) and condition two: only tRA (0.1µM) (Figure 3.8). During the first seven days of differentiation, cells initially organized as a monolayer which then became multilayer with some zones of high cellular density (Figure 3.9).

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Figure 3.8. Schematic representation of the first differentiation protocol to induce iPSCs to ND/UB tip+ lineage. iPSCs (CRL1502, clone C32) were cultured during the first two days with CHIR99021, followed by five days with FGF9, HA, and tRA. At day 7, cells were used to form aggregates which were then cultured under two different conditions- condition one: GDNF (50 ng/µl) + tRA (0.1µM) and condition two: only tRA for the next eighteen days.

Figure 3.9. Bright-field images of differentiation process during the first seven days of the protocol. Cells were cultured in APEL medium supplemented with 6 µM CHIR99021 for two days, followed by five days with FGF9, heparin (HA) and All-trans retinoic acid (tRA).

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In the second step of the protocol, at the time of aggregation (day7+0 onwards) we observed that absence of FGF9 in our cultures severely affected cell survival, and thereby structure formation. Although there were some visible small structures resembling cyst formations at early stages (day7+3), these subsequently disappeared, and the organoids clearly lacked evidence of epithelial formation (Figure 3.10). This finding strongly established FGF9 as a critical component which remains necessary to promote cell survival at the time of aggregation

In view of the role of FGF9 at the time of aggregation, we again established differentiation cultures to evaluate the addition of FGF9 and tRA, with and without GDNF. Using this strategy, organoids survived and successfully formed structures under both conditions studied. These structures resembled small cysts under bright-field microscopy (Figure 3.11). Subsequently, organoids were subjected to immunofluorescence assay against collecting duct protein markers. The structures formed within organoids were positive for the three CD protein markers tested, GATA3/PAX2 and KRT8 (Figure 3.12A). Therefore we then investigated whether organoids cultured with and without GDNF might have a differential gene expression in Wolffian (nephric) duct and metanephric mesenchyme markers. To this end, organoids were collected at day 7+18 for RNA extraction. Although the expression of Wolffian (nephric) duct genes, including GATA3 and RET, was higher than that of the metanephric mesenchyme genes SIX1 and HOXD11, addition of 50 ng/ml of GDNF at the time of aggregation did not significantly increase expression of RET receptors (Figure 3.12B).

Figure 3.10. FGF9 at the time of aggregation is critical for survival and structure formation within organoids. iPSCs (CRL1502, clone C32) were differentiated during the first two days with CHIR99021, followed by five days with FGF9, HA, and tRA. Bright- field images of organoids cultured for eighteen days with tRA (0.1µM) and GDNF (50 ng/µl) + tRA (0.1µM). Organoids cultured under both experimental conditions were unable to survive and form structures in the absence of FGF9. . 100

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Figure 3.11. Studying the effect of the presence and absence of GDNF from the aggregation day onward. A) Schematic representation of the differentiation protocol used. Organoids were culture for five days under condition one: GDNF (50 ng/µl) + tRA (0.1µM) + FGF9 (200 ng/ml), and heparin (1 µg/ml), and condition two: tRA (0.1µM) + FGF9 (200 ng/ml), and heparin (1 µg/ml). FGF9 and HA were removed from the medium from day 7+5 onwards. B) Bright-field images of organoids cultured with and without GDNF, with evidence of cyst-like structures.

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Figure 3.12. Addition of GDNF to differentiation cultures at the time of aggregation did not increase expression of RET receptor. A) Representative immunofluorescence assay of kidney organoids showing evidence for GATA3+PAX2+KRT8+ structures. B) Organoids were collected at day 7+18 to assess m-RNA relative expression of ureteric bud (RET and GATA3) and metanephric mesenchyme markers (SIX1 and HOXD11). Gene expression of each gene was normalised to glyceraldehyde-3-phosphate dehydrogenase. Data represent mean ± S.E.M of three biological replicates.

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AIM2: Validating the use of new collecting duct reporter cell lines on a feeder layer-free system.

The generation of kidney organoids by our laboratory, as well as the first experiments performed for this thesis, were based on iPSCs lines derived and maintained on mouse embryonic feeders (MEFs). Although this was the classical method of culturing iPSCs, it has clear disadvantages, most notably significant batch-to-batch variations. To overcome this, the culture of iPSC lines has progressed to using defined culture conditions to improve transferability and reproducibility. Recent advances in systems of culture have made possible the culture of pluripotent stem cells in a feeder-free system by using the utilization of extracellular matrices instead of feeders. As this switch in approach was happening with the laboratory, particularly with newly derived reporter lines, it was decided to adapt any new differentiation protocol to feeder-free conditions. Given that we wanted to identify a specific end cell type, the research was thought to likely benefit from the use of fluorescently-tagged iPSC reporter lines. Hence this project moved to standardise the use of reporter pluripotent stem cell lines generated by others using CRISPR/CAS9 gene editing and grown in a feeder- free system.

3.2.3 Assessment of the iPSC line: 1502.3 GATA3: m-Cherry clone 60

The generation of the two progenitor populations which give rise to the kidney will provide a wonderful model to illuminate many unknown aspects of the human nephrogenesis process at the molecular level. To pursue this ambitious aim, we had access to unique reporter lines generated by others using CRISPR/Cas9 gene editing of hPSCs as readout of differentiation cultures, isolation and characterisation of specific cell types. We chose the GATA3 gene because intermediate mesoderm (PAX2+LHX1+) forms a GATA3+ nephric duct which extends along the embryo from the rostral end. The ureteric bud arises as a side-branch of this nephric duct and becomes the ureter and collecting ducts of the kidney. In this context, a human GATA3 iPSCs cell line was generated by a sequential approach, as described in more detail in Materials and Methods. Briefly, episomes encoding reprogramming factors are transfected into fibroblasts to create iPSCs, which are then transfected with the homologous reporter cassette and CRISPR/Cas9 RNA. The fluorescence cassette was site-specifically integrated upstream GATA3 start codon, linked via a T2A peptide (Figure 3.13) (Howden et al., 2015).

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Figure 3.13. Sequential strategy for the generation of a fluorescently-tagged cell line (1502.3 GATA3: m-Cherry Clone 60): A) Cherry reporter inserted prior to GATA3 start codon to mark collecting duct following differentiation into kidney organoids. B) Schematic representation of reprogramming and gene targeting by sequential approach. (Adapted from Howden et al., 2015).

The transferability of organoid culture to iPSCs being grown in a feeder-free culture system also required the standardisation of a new step in our differentiation protocol applied at the time of seeding cells for differentiation. In this step, cells were dissociated using TrypLE Select (Thermo Fisher Scientific) as to provide the opportunity to passage the hPSCs as single cells, and then plated in E8 medium (Chen et al., 2011) supplemented with ROCK inhibitor to increase the survival and attachment of cells. Because the success of differentiation experiments relies on plating a precise and reproducible number of cells before starting to induce the cells towards the desire fate, we first investigated whether varying incubation times and commercially available types of ROCK inhibitors, including Y27632 and RevitaCell, made a difference in the outcome of the experiment. This was investigated because several recent reports have argued that Rho kinase has many effects upon cell shape and behaviour, depending upon the timing and the cell type. In 2016, Maldonado et al. concluded that ROCK inhibitor (Y-27632) has an effect on mesendodermal (primitive streak) patterning via E-cadherin inhibition. To assess this, the iPSC line (1502.3 GATA3: m-Cherry clone 60) was seeded using E8 medium supplemented with an equal concentration of ROCK inhibitor-Y27632 5µM,

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Figure 3.14. Evaluation of different rock inhibitors on differentiation cultures. A) Bright-field images during the first seven days of differentiation using the iPSC1502.3 GATA3: m-Cherry clone 60 line. Cells were plated using E8 medium supplemented with r inhibitor- Y27632 (for 5 and 24 hours), and RevitaCell (for 24 hours). B) Flow cytometry of monolayer cultures at day 7 of differentiation.

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Next, we evaluated the capacity of the iPSC line (1502.3 GATA3: m-Cherry Clone 60) by plating with ROCK inhibitor-Y27632 (5 and 24 hours) and RevitaCell (24 hours) to form organoids. While the cell line showed the ability to grow organoids under the three evaluated conditions, each experimental condition resulted in organoids with evident differences in structure formation (Figure 3.15). To evaluate this, organoids were subjected to immunofluorescence assay. Differentiated cells were subjected to treatment with 24 hours of RevitaCell and originated organoids with abundant small structures within the organoid (Figure 3.16). Conversely, organoids derived from cells plated with ROCK inhibitor-Y27632 for 5 hours showed fewer, but larger, epithelial structures preferentially located around the edge of the organoid (Figure 3.16). Surprisingly, incubation with ROCK inhibitor-Y27632 for 24 hours produced better epithelial organization characterized by larger epithelial elements, but these were randomly located throughout the organoid and resembled tubule formation (Figures 3.15 and 3.16). This enhanced the formation of epithelial structures, so we established the standard of 24 hours incubation with ROCK inhibitor-Y27632, applied at the time of seeding cells, for differentiation experiments.

Figure 3.15. Bright-field and confocal images of the developing organoids derived from the iPSC 1502.3 GATA3: m-Cherry clone 60 line across a time series. Differentiation experiments were carried out plating the cells with E8 medium supplemented with equal concentration of ROCK inhibitor-Y27632 5µM (incubated for 5 and 24 hours), and Revitacell (24 hours). The cell line showed ability to differentiate into kidney organoids, forming visible structures under all the three evaluated conditions. The organoids also exhibited live expression of m-Cherry+/expressing cells.

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Figure 3.16. 24 hours incubation with Y27632 promotes better epithelium organization. Immunofluorescence against PAX2 revealed that the use of different ROCK inhibitors resulted in organoids with different epithelial organization. Incubation with: A) RevitaCell for 24 hours, resulted in organoids with several abundant small structures around the whole organoid. B) Y27632 for 5 hours produced a principal and big epithelial structure, C) Y27632 for 24 hours resulted in a big epithelium accompanied by randomly located structures resembling tubule formation. Having validated the use of the iPSC line (1502.3 GATA3: m-Cherry clone 60) to grow organoids, and the use of ROCK inhibitor in our differentiation protocol, we then evaluated the use of this fluorescently-tagged iPSC line as a readout of differentiation. It was therefore critical to ensure that the presence of fluorescence coincided with the presence of the GATA3 protein on all occasions. To this end, immunofluorescence studies were performed. Co- localisation of the native GATA3 protein, identified by antibody staining with the reporter m- Cherry, showed strong evidence of lack of clonality, as shown in Figure 3.17. Several tubular structures were positive for m-Cherry but not for the native protein detected by antibody immunostaining, which invalidated further use of 1502.3 GATA3: m-Cherry Clone 60 for this project.

Figure 3.17. Evaluation of the clonality of the iPSC cell line 1502.2 GATA3: m-Cherry clone 60. Differentiation cultures were established, and organoids collected at day 7+11. Immunofluorescence assay was carried out against, A) GATA3, and B) m-Cherry proteins. C) While some structures showed co-localisation between the native GATA3 protein and the reporter m-Cherry, others lacked co-localisation (arrow heads).

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3.2.4 Assessment of the fluorescently-tagged iPSC line: 2429 GATA3: m-Cherry clone 26

Given the importance of having a clonal collecting duct reporting cell line, a new cell line for the same gene was generated. This time, the cell line was derived by a simultaneous reprogramming and gene editing approach, due to its higher chance of generating correctly targeted clones (Howden et al., 2015). Briefly, episomal vectors encoding reprogramming factors, plasmids encoding guide RNA for the target gene, the homologous reporter cassette, and RNA encoding CRISPR were simultaneously transfected into fibroblasts via electroporation (Figure 3.18).

Figure 3.18. Simultaneous strategy for the generation of a fluorescently-tagged cell line (2429 GATA3: m-Cherry clone 26): A) Cherry reporter inserted prior to GATA3 start codon to mark collecting duct cells following differentiation into kidney organoids. B) Schematic representation of simultaneous reprograming and gene targeting by a sequential approach. (Adapted from Howden et al., 2015).

We therefore examined the clonality of the new fluorescently-tagged iPSC line referred to as 2429 GATA3: m-Cherry clone 26. To do this, cells were used for differentiation experiments and organoids were subsequently made. Organoids were then FACS-sorted at day 7+11, revealing the existence of three populations, Negative-mCherry+, Low-mCherry+, and High- mCherry+ (Figure 3.19A). We therefore proceeded to examine the expression level of GATA3 by qPCR analysis of these three populations. As expected, mRNA expression levels of GATA3 ranged from low (Negative-mCherry+), through intermediate (Low-mCherry+), to high (High- mCherry+) according to the population assessed (Figure 3.19B). Similarly, organoids were collected at the same time-point and evaluated by immunofluorescence. At the protein level,

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Figure 3.19. Validation of the iPSC 2429 GATA3: m-Cherry clone 26 line. Differentiation cultures were established, and organoids collected at day 7+11. A) Flow cytometric analysis of GATA3/m-Cherry+ reporter line at day 7+11. Three different populations were collected: Negative-mCherry+ (14.6%), Low-mCherry+ (41.6%), and High-mCherry+ (23.3%). B) The expression level of GATA 3 was evaluated by qPCR. Gene expression was normalised to glyceraldehyde-3-phosphate dehydrogenase. Data represent mean ± S.E.M of three replicates. C) Immunoflourescence assay against GATA3 and m-Cherry proteins was performed in organoids, showing co-localisation between the reporter m-CHERRY+ and the native GATA3 protein. Scale bar 50 µm.

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AIM3: Investigating the action of chemicals, media, and growth factors upon ureteric bud patterning.

Having validated a GATA3 reporter cell line for identifying collecting duct cells, we then assessed the effects of different chemicals and growth factors known to be critical during ureteric bud lineage specification. With the goal of increasing the expression of RET, we evaluated the addition of 9-cis-retinoic acid (9-cisRA) to our differentiation cultures, based on some studies of the propagation of isolated ureteric bud cells which reported a requirement for both isomers of retinoic acid (-cis and –trans), as well as GDNF, for ureteric branching and expression of the RET receptor in vitro, in the absence of a surrounding mesenchyme (Rosellot et al., 2010). Applying this new understanding, along with previous evidence reported here, a new combination of components and growth factors was tested in our cultures, including: 1) titration of different CHIR99021 concentrations; 2) addition of 9-cis-retinoic acid; 3) increase in concentration of both isomers of retinoic acid; and 4) optimisation of the basal medium. By the time of performing these studies, a critical component of the medium used for our differentiation experiments (STEMdiff™ APEL™ Medium) had been discontinued (late 2016). To test whether the replacement product could give us a better outcome (STEMdiff™ APEL™ 2 Medium lacks protein-free hybridoma medium), we examined all the above described conditions using both media.

In this context, the iPSC 2429 GATA3: m-Cherry clone 26 line was subjected to new differentiation experiments using the technique outlined in Figure 3.20. After seven days of differentiation, monolayer cultures were harvested for RNA extraction to perform gene expression evaluation under all the experimental conditions. Firstly, we found that the condition using STEMdiff™APEL™ medium supplemented with 6µM CHIR99021 led to the highest upregulation of ureteric bud (UB) markers including GATA3, PAX8, RET, and GFRα1, alongside down-regulation of metanephric mesenchymal (MM) markers including SIX2, EYA1, and HOXD11 (Figure 3.21). Secondly, the new basal medium STEMdiff™APEL™ 2 caused a significant reduction in the expression of all the UB and MM genes evaluated, which occurred under all the conditions evaluated (Figure 3.21). Surprisingly, the trend of expression of most of the genes swung in the opposite direction when APEL 2 medium was used as basal medium. Therefore, we concluded that STEMdiff™APEL™ medium supplemented with 6µM CHIR99021 was the optimal condition to commit cells to an anterior intermediate fate with ureteric bud identity.

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We then investigated whether the addition of different growth factors, known to be essential for ureteric bud morphogenesis at earlier stages of the protocol, might trigger increased expression of RET and other genes regarded as being of nephric duct/UB lineage. To do this, we fixed the use of 6µM CHIR99021 in our cultures during the first two days of differentiation, followed by different combinations of growth factors including: 1) earlier addition and higher concentration of GNDF; 2) addition of BMP7; and 3) different combinations of GDNF and BMP7 (Figure 3.22).

Figure 3.20. Experimental technique used to differentiate iPSCs towards an anterior intermediate mesoderm with UB signature. Four different approaches were evaluated during the first seven days of differentiation cultures including: 1) titration of different CHIR99021 concentrations; 2) addition of 9-cis-retinoic acid; 3) increase in the concentration of both isomers of retinoic acid; and 4) optimisation of the basal medium.

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Figure 3.21. Evaluation of different components and growth factors to promote the differentiation of iPSCs toward nephric duct/UB lineage. CHIR99021 titration, both isomers of retinoic acid (-cis and -trans), and different basal mediums were evaluated during the first seven days of differentiation in monolayer cultures using the iPSC 2429 GATA3: m- Cherry clone 26 line. RNA was extracted from all the evaluated conditions at day 7. A) Gene evaluation of nephric duct/ureteric bud markers GATA3, PAX8, GFRα1, and RET; and, B) Metanephric mesenchyme SIX2, HOXD11, and EYA1 markers, revealed increased and preferential expression of ureteric bud lineage when cells are differentiated with STEMdiff™APEL™ medium supplemented with two days of 6µM CHIR99021, followed by 5 days with FGF9, HA, and RA-cis and –trans. Gene expression of each gene was normalised to glyceraldehyde-3-phosphate dehydrogenase. Data represent mean ± S.E.M of three biological replicates. *P <0:05; **P <0:001; ***P <0:0001. A two-tail t-test was performed for APEL vs APEL2.

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Figure 3.22. Experimental technique used to differentiate iPSCs toward nephric duct/ureteric bud lineage. The use of 6µM CHIR99021 was established in our cultures during the first two days, followed by evaluation of the action of different growth factors (GDNF+/BMP7/ GDNF+ BMP7+) in our differentiation cultures. All these conditions were also evaluated with the basal media STEMdiff™APEL™ and STEMdiff™APEL™ 2.

Surprisingly, when adding GDNF at higher concentrations and at earlier stages of the protocol this growth factor was able to trigger significant expression of GATA3 and the RET receptor (Figure 3.23), although there was no significant change in the relative expression levels of GFRα1 and PAX8. As expected and in line with our previous data, the expression levels of SIX2, EYA1, and HOXD11, the metanephric mesenchyme genes, were very low in all the culture conditions evaluated. Moreover, the use of STEMdiff™APEL™ 2 medium again ablated the expression of almost all the genes. Thus we totally ruled out the use of this newly marketed medium in further differentiation experiments and established the use of GDNF+ from day 2 of differentiation onwards to significantly up-regulate the expression of RET and GATA3. Collectively, the results presented in this Chapter demonstrate that the protocol outlined in Figure 3.24 promotes the differentiation of iPSCs toward nephric duct/UB +tip lineage.

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Figure 3.23. Investigation of different growth factors to promote the differentiation of iPSCs towards nephric duct/UB lineage. The iPSC 2429 GATA3: m-Cherry clone 26 line was differentiated with two days of 6µM CHIR99021, followed by 5 days with FGF9, HA, and RA-cis and –trans. Here, we also evaluated the addition of different growth factors GDNF+, BMP7, and GDNF+ BMP7+ from day two onwards. RNA was extracted from all the evaluated conditions at day seven. A) Gene evaluation of nephric duct/ureteric bud markers GATA3, PAX8, GFRα1, and RET; and, B) Metanephric mesenchyme SIX2, HOXD11, and EYA1 markers, revealed upregulation of GATA3 and RET expression when STEMdiff™APEL™ medium was also supplemented with GDNF was added from day two onwards. Gene expression of each gene was normalized to glyceraldehyde-3-phosphate dehydrogenase. Data represent mean ± S.E.M of three biological replicates. *P <0:05; **P <0:001; ***P <0:0001. Two tail t-test was performed for APEL vs APEL2.

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Figure 3.24. Directed differentiation protocol for commitment of iPSCs to nephric duct/UB lineage. Cells are plated and incubated for 24 hours with ROCK-inhibitor Y26372 prior to differentiation experiments. Then, cells are exposed to 6µM CHIR99021 for two days, followed by five days with FGF9, HA, GDNF, 9-cisRA and tRA. At day 7, cells are aggregated to form organoids which are cultured for the next five days in the same cocktail of growth factors. After this period, FGF9 and HA are removed to allow epithelial maturation.

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3.3 DISCUSSION

This chapter describes different optimisation methods, including the use of growth factors, chemicals, and particular cell lines, for improving the patterning of induced human pluripotent stem cells towards nephric duct/ureteric epithelium. Generation of kidney organoids containing different cell types, including collecting duct cells, was one of the major breakthroughs in the kidney field in 2015 (Takasato et al., 2015 and 2016). However, this is self-limiting because of the lack of a sustained progenitor population across time and the relatively short time period for culture. This is turn results from the absence of an apparent branching ureteric epithelium surrounded by a self-renewing progenitor zone. To overcome this limitation, this thesis has focused on the generation of one of the two progenitor populations that give rise to the kidney, the ‘ureteric bud’ population. Here we aimed to generate a new protocol that promotes the directed differentiation of iPSCs to an epithelial population expressing RET, the receptor responding to GDNF produced by the nephron progenitor population, or WNT9B, the ligand which drives both survival and commitment of the nephron progenitor population.

The initial organ culture analysis from E.11.5 kidneys not only validated the activity of key components such as GNDF+, but also supported the essential role played by retinoic acid and GDNF signalling pathways in driving the branching morphogenesis process. Because our major aim was the generation of an epithelial population, we started to perform titration of different components at the time of forming three-dimensional aggregates (organoids) which facilitate self-organising events. At this stage, the sole addition of GDNF and tRA, the two main ligands involved in activating signalling pathways associated with ureteric bud development, was not only insufficient to derive structure formation within the aggregates, but unable to sustain survival of the organoid throughout the culture. This was explained by the absence of FGF signalling in our organoid cultures, beginning on the day of aggregation. FGFs are required in the developing kidney both for their role in cell survival (Qiao et al., 1999), and for modulating the expression of important UB genes such as RET and Wnt11 (Bates, 2011). Therefore, FGF9 and HA were added to the cultures in combination with tRA, plus and minus GDNF, from the day of aggregation. While this new culture condition promoted organoid survival and the formation of small structures resembling cysts in the aggregate, addition of 50 ng/ml of GDNF to our cultures was not sufficient to promote either the expression of RET or GATA3. In fact, the m-RNA levels of GATA3, a critical transcription factor in determining

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There are three possible explanations for this result:

1) Time-related: Although the first seven days of culture represent the stage of anterior- intermediate mesoderm commitment, it is possible that to direct the cells to become an epithelium with UB identity they would need to receive the UB cues earlier than the day of aggregation.

2) Sub-optimal concentration of GFs: Even though we performed a titration of GDNF ligand using organ culture, and there was no apparent difference from 50 to 150 µM of GNDF in promoting branching and swollen UB tips, this alone was insufficient to mimic the environment within a developing kidney which still contains a metanephric mesenchyme.

3) Lack of other key components in the culture: Although GDNF and tRA are the two main ligands involve in ureteric bud pattering, there are other components and signalling pathways that have been described as playing an important role in ureteric bud morphogenesis which we might still be missing, such as 9-cis retinoic acid, canonical WNT/β-catenin, FGFs, WNT11, and TGF-β.

In summary, these studies offered us a preview of anterior-IM patterning, and pointed us towards investigating which other cues might turn on the expression of key genes associated with the RET signalling pathway.

Another factor to take into account when reporting a new methodology is transferability and reproducibility. As the success of this protocol relied not only on finding the right combination of chemicals and growth factors required to guide undifferentiated cells to a nephric duct/ureteric bud fate, but also on being transferable across distinct cell lines and laboratories we switched the culture method of iPSCs on MEFS into feeder free conditions. This was reinforced by the valuable opportunity we had to use reporter iPSC lines derived from this culture system, which allowed the standardization of different culture conditions and enabled us to monitor the expression of key genes regarded as promoting nephric duct/UB lineage. This switch in approach involved adapting the differentiation protocol to feeder-free conditions, to

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As mentioned above, we also had access to unique reporter lines generated using CRISPR/Cas9 gene editing, to track the expression in real time of key genes in response to different culture conditions and to further isolate and characterise specific cell types. Although all the reporter cell lines used in this thesis were generated by others, before using them for optimisation experiments we first had to validate their use. The gene chosen for identifying collecting duct cells was GATA3, as we assumed that we must first generate a correctly patterned nephric duct derived from a GATA3+PAX2+PAX8+ anterior intermediate mesoderm. Indeed, GATA3 together with PAX2/PAX8 and LIM1 comprised one of the earliest described transcriptional networks which stimulated the expression of RET, the gold standard gene defining the ureteric bud/tip+ population (Boualia et al., 2013). The first reporter line which we had access to, the GATA3 iPSC line (1502.3), unfortunately had a lack of clonality, which might have been caused by the sequential approach used in its derivation process. Hence a new GATA3 iPSC line (2429) was generated via a simultaneous approach, to reduce the chance of picking a mixed clone. We could then validate the use of GATA3 iPSC line (2429) by confirming adequate co- localization between native expression of the GATA3 protein and the fluorescently-tagged m-

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Cherry. This finding was also supported by evaluation of the m-RNA levels of GATA3 in the m-Cherry+ sorted fractions. These data conclusively validated the use of this cell line as a readout of differentiation cultures, and for further sorting of the GATA3+ expressing cell fraction.

Critically, the development of this protocol also relied upon our comprehensive understanding of kidney development. In the developing embryo, the nephric duct lineage derives from anterior-intermediate mesoderm (AIM), and cells migrating towards the rostral end of the mesoderm early in development are influenced by low levels of Wnt signalling and high concentrations of RA and FGF9 morphogens (Xu et al., 2014). Based on this knowledge, and having also gathered some evidence from our previous differentiation cultures suggesting patterning to anterior-intermediate mesoderm while still lacking UB/tip+ identity, we established new optimisation cultures. Because we thought the lack of evidence of RET expression might be due to the late addition of GDNF+ and tRA to our cultures, new variables and components were examined during the first seven days of the differentiation protocol. This included testing of new components and growth factors, increasing the concentration of key components, and testing different basal mediums. Since, to our knowledge, a balance between several signalling mechanisms is required for the successful patterning of undifferentiated cells to ureteric epithelium with tip+ identity, we first performed titration of WNT signalling (CHIR99021) from 6 to 10 µM. Our qPCR results demonstrated and reconfirmed that low CHIR directs the cells to an AIM fate, which was important to corroborate because of the switch in the culture system. Secondly, retinoic acid signalling has been also described as remarkably predominant in directing ureteric bud lineage. In fact, more recently, both retinoic acid isomers- All-trans-and 9-cis- have been shown to activate different nuclear receptors (RAR and RXR, respectively), potentially activating different subsets of genes. Hence, we added 9-cis-retinoic acid to our cultures. This approach was also reinforced by existing studies on the propagation of isolated ureteric epithelium, which show a requirement for both cis- and trans- retinoic acid, as well as GDNF, for ureteric branching in vitro in the absence of a surrounding mesenchyme (Rosellot et al., 2010). Thirdly, we examined the used of two basal mediums (STEMdiff™APEL™ Medium and STEMdiff™APEL™2 Medium-lacks protein-free hybridoma medium). Our data conclusively showed that when iPSCs were differentiated in monolayer cultures using STEMdiff™APEL™ supplemented with two days of 6 µM CHIR99021 followed by five days with FGF9, HA, RA-cis and trans- the expression of key genes associated with nephric duct/UB lineage (turned on. Conversely, genes regarded as metanephric mesenchymal lineage including, SIX2, EYA1, or HOXD11 were downregulated.

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Surprisingly, the use of APEL™ 2 as basal medium switched the gene expression pattern. This difference was related to the lack of protein-free hybridorma in this medium. Hence STEMdiff™ APEL™ was established as the basal medium for this differentiation protocol. While it is still difficult to distinguish between whether we were patterning to a nephric duct or ureteric epithelium what could be answered through a more comprehensive gene analysis, this data gave us great confidence that iPSCs were preferentially patterning to nephric duct/ureteric bud lineage.

Finally, and considering that the branching morphogenesis process is driven by positive and negative regulators which coordinate the initial budding (Costantini and Kopan, 2010) and the subsequent branching of the tree-like structure, we examined the action of GDNF, BMP7, and the combination of both growth factors in our cultures. Based on this paradigm, we added higher concentrations of GDNF as a stimulator, and BMP7 as inhibitor of the morphogenesis of the branched epithelia, from day two onwards in our differentiation cultures. BMP7 provoked a dramatic reduction in the expression of all the genes associated with UB lineage (GATA3, PAX8, WNT9B, GFRα1, and RET) even in combination with GDNF. While BMPs signalling is defined as a repressor of UB branching, its effect as a repressor has been described as playing an important role in the maintenance of an orchestrated equilibrium among the molecular signalling pathways governing the process, thereby collectively supporting branching morphogenesis. We interpreted this marked repressive effect as the result of an inadequate concentration of BMP7 in our cultures, but it may be that the concentrations used mimicked the suppression of branching induced by BMPs signalling. Conversely, when GDNF was added at a higher concentration and earlier in culture than in the first assays performed, the gene expression levels of RET and GATA3, the receptors responding to GDNF produced by nephron progenitors, were significantly upregulated by day 7 of the differentiation. Although RET and GFRα1 are expressed in the nephric duct long before there is evidence of ureteric bud budding, the expression of these two receptors is critical for promoting ureteric bud invasion, which leads to specification of the UB into tip and trunk segments and the subsequent branching morphogenesis process. We therefore concluded that when cells are cultured with two days of 6 µM CHIR9902, followed by five days of FGF9 200 ng/ml, heparin 1 µg/ml, 9-cisRA 0.2 µM, tRA 0.2 µM, and GDNF 100 µg/µl, they rapidly specify to nephric duct lineage with a strong UB identity, as demonstrated by peak expression of GATA3, PAX2 GFRα1, RET AND WNT9B at day seven of differentiation.

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Taken together, the data reported in this chapter demonstrate the successful generation of a methodology to differentiate iPSCs to nephric duct/ureteric bud lineage. This is a promising start, however all the markers of UB are also expressed but not necessarily critical for nephric duct formation. It is therefore difficult to distinguish between whether we have induced nephric duct or ureteric bud or even whether this matters with respect to the capacity of these cells to respond to a metanephric mesenchyme. Despite this, to our knowledge this is the first approach which has shown evidence of increasing expression of both RET and WNT9B, both key nephric duct/ureteric bud genes. This protocol was also adapted to feeder-free culture systems to ensure a fully defined, highly reproducible cell culture system which could operate across distinct cell lines and laboratories. Moreover, we also validated the use of a fluorescently-tagged GATA3 iPSC line as a tool in collecting duct differentiation. This protocol will be challenged in Chapter Four, using two fluorescently-tagged iPSC lines to thoroughly evaluate and characterise its capacity to derive epithelia with UB/CD identity.

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CHAPTER IV

Investigation into a directed differentiation protocol for the generation of nephric duct/ureteric bud epithelium from human pluripotent stem cells

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4.1 INTRODUCTION

While cellular therapy for the treatment of kidney injury is one approach, with advances in methods for the derivation and differentiation of stem cells a second option is stem-cell based regeneration. As described in Chapters One and Three, in 2015 our laboratory developed a detailed protocol for the formation of kidney organoids containing nephrons, stroma, vasculature, and evidence of GATA3/CDH1+ distal tubular segments assumed to represent collecting duct. However, the lack of sustained renal progenitor populations made this approach self-limiting for recreating kidney tissue. To overcome this limitation, we developed a new protocol to direct induced pluripotent stem cells to nephric duct/ureteric bud fate, as outlined in Chapter Three. Our major objective in Chapter Four was to thoroughly evaluate the capacity of this protocol to generate nephric duct/ureteric bud epithelium. This required a broad understanding of the stepwise formation of the collecting duct system. In Chapter One we reviewed the main basis of kidney morphogenesis, covering the different stages of the branching morphogenesis process, and in Chapter Three we covered the molecular basis of ureteric bud formation. Here we will comprehensively cover how patterning of CD systems occurs in a stepwise fashion across development, reviewing nephric duct lineage specification, segmentation of the ureteric bud epithelium into tip and stalk structures, and finally cell differentiation events that trigger formation of the mature collecting duct system.

4.1.1 Anterior intermediate mesoderm and its derivatives

4.1.1.2 Nephric duct lineage specification

The formation of excretory organs is mediated by induction of the intermediate mesoderm to undergo lineage specification, with the nephric duct (Wolffian duct) primordium being the first structure to appear (Figure 4.1A, B). Initially, paired ductal structures are derived simultaneously with the nephric duct primordium (ND) and the resulting nephric cord (NC) elongating caudally as the trunk of the embryo does (Figure 4.1C, D, E). Although it was initially postulated that ND extension was mediated by recruitment of new cells, mice studies support the current understanding that cell migration and proliferation within the duct epithelium itself are the main biological processes involved (Mugford et al., 2008). Nevertheless, little is known at the molecular level. Here we will not discuss the processes underlying morphogenesis and derivation of the metanephric kidney, as they were extensively

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The nephric duct arises from the collaborative action of mediolateral gradients of BMP/Activin, in combination with the high retinoic acid gradient existing in the anterior portion of IM, thus defining the expression of the HOX family of genes along the anterior-posterior axis. Indeed, the correct assembly of the Hox domain is essential for correct patterning, localization, and orientation of renal derivatives. This process also requires co-expression of the gene core composed of LIM1/PAX2/PAX8 (Bouchard et al., 2002, Caroll et al., 1999).

The normal extension of ND involves the establishment of gene networks which subsequently activate downstream regulators modulating tissue morphogenesis and cellular specialization. In this context, PAX2 AND PAX8 are shown as essential regulators of nephric duct lineage. Mice lacking Pax2 and Pax8 exhibit failed pronephros formation, as well as loss of all subsequent renal structure, caused by inability to undergo mesenchymal to epithelial transition (Bouchard et al., 2002). These two transcription factors function jointly to maintain LHX1 expression and trigger the expression of the transcription factor GATA3. (Bouchard et al., 2002; Grote et al., 2006). Even though mice lacking Lhx1 are able to form ND, its caudal portion subsequently degenerates (Perdersen et al., 2005, Tsang et al., 2005). In contrast, loss of GATA3 expression results in ductal swelling of the ND, which then fails to fuse with the cloaca (Grote et al., 2006, Lim et al., 2000). Although they initially form distinctive phenotypes, defective mice of either Lhx1 or Gata3 are unable to form the permanent metanephric kidney, indicating that these factors are critical regulators of nephric duct morphogenesis. While there are many unknown molecular aspects of the genetic circuit composed by LIM1/PAX2/PAX8/GATA3, it constitutes the most important gene network known to date which regulates ND lineage.

As the nephric duct emerges from the primordium to develop into the ureteric bud, its epithelium also responds to surrounding changes. Evidence from chick studies have reported BMP4 as one of the candidates which could influence the initial conversion from a mesenchymal cord to an epithelial tube (Obara-Ishihara et al., 1999). The ND corresponds to a simple cuboidal epithelium until it reaches the cloaca at E.9.5 (Figure 4.1E, F). Once the ND starts swelling to form the ureteric bud, the increase in cell proliferation changes the nature of the epithelium to a pseudostratified epithelium which persists throughout ureteric bud

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Figure 4.1. Stepwise formation of the nephric duct and renal collecting duct. A) Schematic diagram depicting E.9.5 embryo; left, presentation nephric duct (green, ND) and nephrogenic cord (blue, NC); right, cross section of the embryo showing the ND, NC, neural tube (NT), somites (S), and lateral plate mesoderm (LPM). B) Intermediate mesoderm at E.8.5. C) Nephric duct (ND) and nephrogenic cord (NC). D) Nephric duct elongation. E- F) Epithelialisation of ND, and formation of MM. G) The rostral part of ND maintains a cuboidal epithelium, but the caudal ND forms pseudostratified epithelium in the zone where MM interacts with UB. H) Outgrowth of the UB. I) First branching event of UB within MM. J) Representation of the UB after 3 to 4 rounds of branching. K) Ureteric bud after several rounds of branching. L) E.18.5 showing elongation of the ducts to form the medulla. M) Intense elongation of the collecting ducts to form the papilla. (Extracted from Constantini and Kopan, 2010).

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4.1.2 Segmentation of the ureteric epithelium

As the metanephric kidney arises, the UB epithelium is defined by two segments, the ureteric tip and stalk. While the tip region contains the proliferative population, which degenerates shortly after birth (post-natal day 3 in mice) (Hartman et al., 2007), the stalk differentiates and undergoes specification and segmentation to form the mature collecting duct system (CD). Although there are still many unknowns about how this occurs, the collecting duct is composed of different cell types. Anatomically, the CD can be categorized into four segments, the tip, the cortical collecting duct (CCD), the medullary collecting duct (MDC), and the urothelium. The mature CD is also defined by four different cell types, principal cells, and intercalated cells types A, B and non-A or B. Hence, eight different cells types can be identified as populating the collecting duct structure of the developing/mature kidney (Figure.4.2)

B) C)

Figure 4.2. Ureteric bud segmentation and cell types. A) A cross-section of embryonic kidney at E.15.5 showing segmentation of the ureteric tree into tip, cortical collecting, medullary collecting duct, and B) Urothelium. Specific markers have been identified for each region, including Slco4c1 (tip-specific), Vldlr (cortical enriched), AI836003 (medullary-specific), and Upk3a (urothelium). C) Representation of the nephron, highlighting the collecting duct system and distribution of different cell types. PCs: principal cells, ICs-type A: Intercalated cells type A, ICs-type B: Intercalated cells type B, ICs- type non A-B: Intercalated cells type non A-B. (Adapted from 38-Thiagarajan et al., 2010)

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Many efforts have been made to characterise the gene expression related to specific segments, which would allow us to understand differentiation along the kidney. Although the molecular mechanisms involved in patterning ureteric bud tip and stalk are not well understood, gene knockout studies in mice, microarray, RNAseq, and more recently single cell RNA-Seq analyses, have been important tools to characterise cell diversity and heterogeneity within a compartment. While the canonical Wnt signalling is widely understood to maintain UB cells in the precursor state (Marose et al., 2008), the protein heregulin alpha (HRGA) has been shown to specifically promote expression of stalk and repress tip markers (Sakurau et al., 2015). Similarly, sonic hedgehog pathway is active in the stalk population and is postulated to play a role in cell proliferation via cyclin B1 activity (Jenkins et al., 2007). In the mature ductal structure, while Foxi1 stimulates intercalated cell fate (Blomqvist et al., 2004), NOTCH signalling promotes principal cell specification (Guo et al., 2014, Jeong et al., 2009).

The activation of different signalling pathways in the CD leads to the expression of widespread and/or segment-specific genes. As outlined in Chapter 3, retinoic acid plays a vital role not only in CD formation, but also in its determination, by promoting specific expression of CD markers such as Scnn1b (encoding βENAC) and Elf5 (Takayama et al., 2014). While some genes, such as Hoxb7 and Calb1, have been described to be ubiquitously expressed in the CD (Srinivas et al., 1999, Georgas et al., 2008), others such as Wnt11, Ret, and Gfrα1 are enriched in the tip population (Majumdar et al., 2003). Similarly, Upk3a, encoding uroplakin-3a, an essential protein regulated by BMP4, forms the urothelial plaque and is specifically expressed within the medullary collecting duct (Thiagarajan et al., 2011, Jenkins et al., 2005). Moreover, in the last decade microarray analysis in early developing mouse has revealed new UB anchor genes for two compartments, the UB-tip (Slco41) and MCD (Gsdmc4, Clmn, AI83600, Fam129a, and Upk3a) (Thiagarajan et al., 2011). Taken together, the temporo-spatial activation of different molecular pathways and genes along the CD is essential for formation of a proper and functional segmented ductal system, allowing us also to detect distinctive populations within heterogeneous samples.

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4.1.3 Collecting duct epithelial cell differentiation

Having been correctly differentiated and determined, the resulting stalk epithelium develops into the mature collecting duct network formed by two distinctive cellular types, principal (PCs) and intercalated cells (ICs). While intercalated cells are more abundant in the renal cortex, principal cells are more abundant in the medulla/papilla region (Guo et al., 2014). PCs are involved in the water and ion transport effected by specific transporters, Aquaporin-2 (AQP2) and sodium channel (ENAC) respectively (Pearce et al., 2014). ICs maintain acid-base homeostasis by regulating protein and bicarbonate secretion. Intercalated cells are sub- categorized into three types, according to the expression of specific transporters- type A, type B and type non-A or B cells (Alper et al., 1989, Nakai et al., 2003). In fact, type A cells perform their function via apical exchanges of H+-ATPASE and H+/K+, via which proteins are secreted, while type B cells secrete bicarbonate through apical exchanges of Cl−/HCO3− (Kim et al., 1999). Although ICs and PCs perform specialised functions, some authors have reported evidence that they come from a common progenitor. For example, in vitro studies have shown that type B-ICs have a capacity to turn into type A-ICs, principal cells, and hybrid cells (Fejes el al., 1992, Fejes et al., 1993). Another study also showed cell conversion from type B-ICs to A-ICs, without changing the total number of intercalated cells, by managing metabolic acid in vitro (Schwartz et al., 1985). On the other hand, it is notable that the renal papilla, anatomically located within the inner medullary collecting duct (IMCD) region, has cytostructure and function mostly made up of a distinct cellular type (Madesen et al., 1998). IMCD cells are modulated by anti-diureteric hormones and play a vital role in urine concentration and acidification via active reabsorption of urea, water, and sodium chloride, as well as in the fine tuning of the final pH (Madsen et al., 1998). These cells also produce nitric oxide in the kidney, promoting natriuresis and diuresis and thus regulating the renal response to salt intake (Mount et al., 2006).

As mentioned earlier, at the molecular level the fate of ICs and PCs is positively modulated by the transcription factor Foxi1 and the NOTCH signalling pathway, respectively. Loss of Foxi1 leads to defective intercalated cells lacking the vacuolar-H+-ATPase AE1 and the anion exchanger AE4, resulting in mice with renal tubular acidosis (Blomqvist et al., 2004). Moreover, in 2006, Kurth and colleagues demonstrated that Slc4a9, encoding the apical transporter Cl−/HCO3−, is a direct target of Foxi1 and thus demonstrated the stimulative action

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In summary, although the renal collecting duct does not appear to be a complex system formed by different structures, its correct differentiation, patterning, and anatomical segmentation are key to forming a sophisticated tubular network with a heterogeneous cellular composition. That network is vital for its physiological function of regulating the fluid/electrolyte balance in the body. We have reviewed here the molecular mechanisms, anatomical and phenotypical segmentation critical to determine the formation of the collecting duct system. This understanding was applied in this Chapter to examine and challenge the capacity of the methodology reported in Chapter Three to direct iPSCs towards nephric duct, or its derivative ureteric epithelium.

4.1.4 Hypothesis and aims

Hypothesis:

The ND/UB protocol promotes specific derivation of nephric duct/ureteric bud epithelium in kidney organoids

AIM 1: Examination of the capacity of the ND/UB protocol to trigger collecting duct formation

AIM 2: Evaluation of the response of a RET+ reporter cell line to the ND/UB protocol

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4.2 RESULTS

Aim 1. Examination of the capacity of the ND/UB protocol to trigger collecting duct formation

Having assessed the clonality of the iPSC 2429 GATA3: m-Cherry clone 26, this line was chosen for more extensive differentiation experiments. To this end, cells were plated at day 1 using 5µM of rock inhibitor-Y27632 for 24 hours to obtain a density of 20 x 103 cells per cm2. Next, differentiation cultures were exposed to 2 days of 6 µM CHIR99021 to pattern the cells towards an anterior intermediate mesoderm (AIM), followed by 5 days of FGF9 200 ng/ml, heparin 1 µg/ml, 9-cisRA 0.2 µM, tRA 0.2 µM, and GDNF 100 µg/ul. During the first seven days of culture, cells behaved as a monolayer which then became a multilayer with some areas of higher cell density. This is the typical phenotype when driving cells towards an AIM differentiation (Figure 4.3). After this, cells were used to form organoids to promote better 3D epithelial organization. Organoids were kept in culture for the following five days with the same cocktail of growth factors (day 7+5), all of which are essential to derive the UB/tip+ population by induction of cell proliferation and the expression of genes regarded as key markers of the progenitor UB population (Figure 4.3). We hypothesised that following this methodology we might be able to manipulate cells to fate-switch to nephric duct or its derivative, ureteric bud.

4.2.1 Assessment of the capacity of the protocol to trigger epithelia formation in two- and three-dimensional approaches.

The cellular composition and epithelial organization within organoids was then evaluated using immunofluorescence of aggregates at two different time-points. Surprisingly, organoids generated with this protocol formed robust epithelial structures which were Cherry+GATA3+ECAD+KRT8+, which is consistent with an enriched nephric duct/ureteric bud epithelium (Figure 4.4 and 4.5). Organoids collected at day 7+5 showed a dense epithelium characterized by small and abundant structures in the centre which seemed to organise around the edges to form a main connected epithelium.

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Figure 4.3. Bright-field images of differentiation of GATA3/m-Cherry+ reporter line to kidney organoids across the ND/UB protocol. First step (A-D): Series of bright-field images showing the features of anterior-intermediate mesoderm cultures using the GATA3/m-Cherry+ reporter line during the first seven days. The basal medium supplemented with 6 µM CHIR99021 for two days, and then replaced for 200 ng/ml FGF9, 1 µg/ml Heparin, 0.2 µM tRA, 0.2 µM 9-cisRA and 100 ng/ml GDNF for five days. Second step (E-H): Bright-field and confocal images of kidney organoids derived from GATA3/m- Cherry+ iPSCs showing the morphological features and activation of the m-Cherry+ reporter. At day seven, cells were used to make aggregates which were cultured for five days with the same cocktail of grow factors, and then with for 0.2 µM tRA and 9-cisRA, and 100 ng/ml GDNF for six days.

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Figure 4.4. Kidney organoids generated by ND/UB protocol contain robust collecting duct structures. Morphological features of kidney organoids generated from GATA3/m- Cherry+ iPSCs were analysed at two times d7+5 and d7+11. A-K: Organoids collected for both time points showed improved and positive GATA3+CHERRY+ECAD+ epithelium. I-K: Large tubular structures with well-pronounced lumen were restrictive for d7+11. Perfect co-localization between the reporter m-CHERRY+ and the native GATA3 protein was also detected. Scale bar: A-B: 200 µm, C-D: 50µm and E: 100 µm.

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Interestingly, epithelial structures resembling ureteric tips were detected around the edge of day 7+5 organoids, just as occurs in vivo during budding of the ureteric bud (Figure 4.4 C-E). At the last collection point, day 7+11, we found large connected epithelial ducts with well- defined lumens, suggesting a more mature epithelium (Figure 4.4 I-K). Moreover, the capacity of this protocol to elicit structure formation was also examined in two-dimensional cultures up to 20 days, using the same reporter line as a readout of ND/UB formation (Figure 4.6). While GATA3 mCherry/+ cells were initially sparse throughout the well, as the culture progressed a dense mesh of Cherry+ cells was detected from day 7 to 9, which then organised as a well- structured epithelial network from day 9 onwards (Figure 4.7). Indeed, immunofluorescence assays demonstrated that Cherry+ structures co-localised with ECAD+GATA3+PAX2+KRT8+ (Figure 4.8). Even more interestingly, these structures were also immunopositive for ETV5 (Figure 4.8I), a downstream transcription factor of RET signalling, which is widely described as a ureteric bud marker (Lu et al., 2009).

Figure 4.5. Immunofluorscence of GATA3/m-Cherry+ iPSCs-derived organoids. D7+11 organoids formed epithelial structures positive for cherry+ and KRT8. Scale bars A: 200 µm, B-C: 50µm

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Figure 4.6. Extended two-dimensional culture of GATA3/m-Cherry+ iPSCs following UB protocol. Series of Bright-field images of extended differentiation cultures for 20 days following the UB protocol. At day 2, cells become spikey and triangular after 6 µM CHIR treatment. Between days 6-8, high cell density characterizes the culture. Between days 9- 20, structure formation become visible, forming connecting networks.

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Figure 4.7. Live expression of m-Cherry+/expressing cells. Series of confocal images following the organization of GATA3/m-Cherry+ cells between day 8 to 20 in monolayer cultures differentiated with the UB protocol. m-Cherry+/expressing cells form initially a mesh of cells(day8-day12), which then organized in more complex connected networks. Scale bar, 100 µm.

Figure 4.8.6. Protein Protein assessment assessment in in structures structures formed formed in in two two-dimensional-dimensional extended extended cultures with UB UB protocol. protocol. GATA3/mGATA3/m--Cherry+Cherry+ reporterreporter line line after after 18 18 days days of of differentiation following the UB protocol. Immunofluorescence confocal microscopy differentiation following the UB protocol. Immunofluorescence confocal microscopy assay assay showing formation of epithelial positive structures for m- Cherry+/GATA3+/ETV5+/ECAD/PAX2+KRT8+.showing formation of epithelial po Scalesitive bar, 20 structures and 100 µm. for m- Cherry+/GATA3+/ETV5+/ECAD/PAX2+KRT8+. Scale bar, 100 µm (A-H) 20 µm (I-L).

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Given the fact that the protocol previously reported by our laboratory showed formation of epithelial structures positive for ECAD+GATA3+PAX2+, we evaluated its capacity in triggering CD formation. To this end, extended monolayer cultures of the GATA3mCHERRY/+ cell line were established using the protocol reported by Takasato et al., 2016. During the first seven days of differentiation the cells primordially displayed a pattern of balling up; subsequently some structures were visible via bright-field microscope after day 8 in a two-dimensional culture (Figure 4.9). Immunofluorescence of these cultures revealed that, while GATA3+ECAD+ structures were detectable (Figure 4.10), they were clearly less abundant and less complex than the epithelial network obtained after differentiating the same iPSC line with the ND/UB protocol.

Figure 4.9. Extended two-dimensional culture of GATA3/mCherry+ iPSCs following standard protocol (ST). Series of bright-field images of differentiation cultures after 18 days following the protocol reported by Takasato et al., 2016. These differentiation cultures show a tendency to ball up during the first seven day of culture. Between days 8 to 18, some structures become visible over differentiated cells (circular patches). The area surrounded by a dashed line is magnified in the next right picture.

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Figure 4.10. Evaluation of the capacity of the standard protocol in generating UB/ collecting duct structures. Immunofluorescence assay of monolayer cultures after subjecting GATA3/m-Cherry+ reporter line to the differentiation protocol reported by Takasato et al., 2016 showing evidence of Cheery+/ECAD+, indicating collecting duct. A) While collecting structures arise after subjecting the cell to the standard protocol, low- magnification view of culture revealed less complex and abundant structures. B) and C) represent close-up images of the areas indicated by a dashed line. Scale bars are 500 (A), 100 (C) and (B) 50 µm.

We then also addressed the question of whether the ND/UB protocol might be inducing cells towards other types of renal structures unrelated to the nephric duct lineage. To examine this, organoid culture as well as extended monolayer culture were carried out to evaluate protein expression. Using this protocol, no other nephron segments were detected, as evidenced by an absence of staining for markers of glomeruli (NEPHRIN+) or proximal tubules (LTL+). This validated the specificity of the ND/UB protocol to generate renal structures derived from the anterior intermediate mesoderm (Figure 4.11).

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Figure 4.11. Absence of nephrons in organoids and 2D-structures generated by the UB protocol. Immunofluorescence of A) kidney organoids at d7+11, and B) extended monolayer cultures (18 days) did not show evidence of glomeruli (evaluated by NEPHRIN) as well as proximal tubule (evaluated by LTL). Scale bars are 200 (A), and (B) 20 µm.

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4.2.2 Evaluation of the molecular signature of Cherry+-expressing cells.

At present, the ND/UB protocol described in this thesis has resulted in patterning towards an anterior intermediate mesoderm. To characterise the molecular signature of this GATA3mCherry/+ population, FACS and gene expression studies were also carried out. Given that previous evidence indicated that Cherry+ cells turn on within the first seven days using the ND/UB differentiation protocol, differentiation cultures and organoid cultures were carried out to perform time-course collection of Cherry+ cells, as indicated in figure 4.12A. At day 7, 18.2% of cells became Cherry+, which then increased to 40.6% for day 7+5, and 34.3% for day 7+11. (Figure 4.12B). QPCR analysis of the positive sorted cells across the three time- points evaluated revealed high expression of the core ND/UB genes PAX2, PAX8 and GATA3 which are necessary for morphogenesis and guidance of the nephric duct in the developing kidney (Figure 4.12C). A significant enrichment of tip-specific markers, including RET, GFRα1, and WNT11, was detected at day 7 and day 7+5, with a minor signature of stalk/trunk genes at day 7. Conversely, expression of stalk+ markers reached its highest expression level at day 7+11. Interestingly, the WNT9B stalk+ marker had a significantly increased expression at the last time-point, suggesting a shift in the epithelial pattern towards a more differentiated CD epithelium (Figure 4.12C).

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Figure 4.12. Enriched expression of key genes of nephric duct/ureteric lineage. GATA3/m-Cherry reporter line was subjected to the ND/UB differentiation protocol for further FACS to isolate m-Cherry + cells and gene evaluation. A) Schematic representation depicting strategy for different collection times of Cherry-expressing cells. B) Flow cytometric analysis of GATA3/m-Cherry reporter line at different times. Day7 (monolayer), 18.2 % of the cells become m-Cherry+; Day7+5 (organoids), 40.6 % of the cells become m-Cherry+; Day7+11 (organoids), 34.3 % of the cells become m-Cherry+. C) A stringent gene evaluation of the Cherry-expressing cells was carried out to examine different stages throughout collecting duct development including nephric duct (PAX2, PAX8 and GATA3), segmentation of ureteric epithelium (tip: RET, GFRα1, WNT11 and stalk markers: WNT7B, TACSTD2, WNT9B) and mature collecting duct (AQP2 and UPK3A). All the genes were normalized to GAPDH and then compared with levels in undifferentiated iPSCs.

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Aim2. Evaluation of the response of an iPSC RET/ tdTomato line to the ND/UB protocol

4.2.1 Characterization of the response RET/ tdTomato line to the ND/UB protocol

As has been extensively reviewed and discussed in this thesis, one of key markers of the nephric duct/ureteric tip+ epithelium is RET, the receptor responding to GDNF produced by nephron progenitors (Costantini and Kopan, 2010). Because we have seen robust evidence indicating that the ND/UB protocol reported here generates a correctly patterned nephric duct (derived from a PAX2+PAX8+GATA3+), and because the QPCR data revealed increased expression of several key ND/UB tip+ genes, a new fluorescently tagged induced human pluripotent stem cell line reporting for RET expression (RET/tdTomato+ iPSCs), obtained from a collaboration with Dr. Sanjay Jain at Washington University, was subjected to the same directed differentiation protocol. During the first seven days of differentiation the RET/tdTomato+ cell line differentiated with an initial sheet-like appearance which then became very dense, in line with what we previously described as an anterior differentiation pattern (Figure 4.13).

A) B)

Figure 4.13. Schematic representation and differentiation process of the RET/tdTOMATO (GCamp6) iPSCs reporter line. A) Diagram of the fluorescently tagged RET/tdTOMATO (GCamp6) iPSCs line which allows the identification and isolation of the nephric duct/tip+ cell fraction. B) Bright-field images of the cells during differentiation following the UB protocol.

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Notably, activation of the reporter tdTomato+ was first detected between days 5 and 6 with red fluorescence persisting into the next stages of culture (Figure 4.14). The appropriate response of this fluorescently tagged iPSC line validates the data generated using the GATA3-mCherry reporter line. At the organoid level, the iPSC RET/tdTomato+ line also displayed the capacity to form organoids with visible structures which appeared at day 7+2 and which were also positive for tdTomato+ expression. The epithelial nature of tdTomato+ structures was validated by performing immunofluorescence for ECAD and GATA3. Epithelial structure present in these cultures were ECAD+/tdTomato+/GATA3+ (Figure 4.15).

Figure 4.14. Live expression of the tdTomato+-expressing cells. Demostration of succesful response of RET/tdTOMATO+ (GCamp6) reporter line to the ureteric bud differentiation protocol in monolayer culture. Initially, tdTomato+-expressing cells are spread out through the culture plate (upper panel), forming later organized structures. Scale bars are 100 (upper panel), and 200 µm (below panel).

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Figure 4.15. Bright-field and confocal images of the developing RET/tdTOMATO+ organoids across a time series. A) RET/tdTOMATO+ iPSCs line showed ability to differentiate into kidney organoids, forming visible structures at day7+2. B) Immunofluorescence of d7+5 organoids revealing RET-tdTomato+/GATA3+/ECAD+ structures, scale bars 50 µm.

While RET expression is detected as early as E.8.5-10.5 (Pachnis et al., 1993), and thus expressed throughout the nephric duct, after commencement of ureteric budding its expression becomes restricted to the UB/tip+ population, being able to upregulate the expression of other downstream markers including WNT11 (Majumdar et al., 2003), ETV4, and ETV5 (61-Lu et al., 2009). Therefore, to provide further insights into the molecular signature of the RET/tdTomato+ population derived from ureteric-tip optimised cultures, organoids were enzymatically dissociated, as described in ‘Materials and Methods’ chapter, and then FACS- sorted. At day 7+5, 17.8% of cells became tdTomato+. Subsequently, RNA was extracted from the sorted populations to evaluate expression levels of a sub-set of genes distinguishably expressed in the tip+ and stalk+ segments. Strikingly, the expression levels of genes associated with cells of the ND/UB tip+ were significantly increased, including RET, GFRα1, and ETV4 (Figure 4.16). Conversely, evaluation of the gene set of UB/stalk markers (WNT7B, WNT9B, and TACSTD2) did not show either increase or equivalence of RNA levels when compared to human foetal kidney expression (HFK) (Figure 4.16).

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Figure 4.16. Gene expression analysis of RET/tdTomato+ expressing cells. A) FACS analysis of day 7+5 organoids derived from RET/tdTomato+ reporter line, 17.8% of the cells become tdTomato+. B) QPCR analysis of FACS-purified tdTomato+ cells indicate enrichment for key UB/tip+ genes. All the samples were normalized to GAPDH and then were then compared with levels in human fetal kidney RNA (HFK).

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4.2.2 Dissecting cell composition in organoids derived from the iPSC RET/tdTomato+ line

To more comprehensively evaluate the gene expression of the cells present within these cultures, organoids generated using the iPSC RET/tdTomato+ line were dissociated into a single cell suspension using Liberase TM (Sigma). The protocol involved dissociation and digestion of organoids with the enzyme for 10-20 minutes at 40 for 20 min by mixing at every 5 minutes using a Gilson pipette, followed by two passages through a 30 µM filter. The entire procedure was carried out at 40C. Dissociated viable single cells were subjected to barcoding and transcriptional library preparation using the 10x Chromium approach to be finally subjected to Illumina sequencing (performed by the Australian Genome Research Facility).

Resulting data was mapped to the hg38 human reference genome and processed initially using the Cell Ranger pipeline (v1.3.1). Quality control of the resulting expression matrix was performed using the Scater package giving a filtered dataset with 2,207 cells and 11,972 genes. A median of 2,382 genes were expressed in each cell. Normalisation, clustering analysis and visualisation were then performed using Seurat (Satija et al., 2017). tSNA plots identified a total of 7 unique cell clusters as identified by differential gene expression (DE genes) (Figure 4.17A). GO analysis of each cluster was performed using the ToppFun tool within the Toppgene suite (https://toppgene.cchmc.org/). Analysis of the DE genes identified a strong similarity between the third largest cell cluster, Cluster 2, and the ureteric epithelium. Notably, DE expression was seen in this cluster for genes such as LHX1, PAX2, PAX8, RSPO3 and GFRα1 (Figure 4.17B). Of 155 differentially upregulated genes in cluster 2, 49 of these genes are present within the top 2500 genes expressed in the mouse E15.5 collecting duct epithelium, and 43 DE genes within the top 2500 genes expressed in the mouse E.11.5 ureteric tips (see Table 1 and 2). The expression of key genes across all clusters was also examined. Of note, despite the presence of clear red fluorescence in cultures, there was very little expression of RET within any cells (Figure 4.17B). GFRa1 expression was localised to cluster 2 but was also detectable in a number of other cell clusters. We assume this reflects a level of RET expression below detectable using single cell profiling. However, this does suggest that the culture protocol is not yet generating high levels of RET expression. The identity of the other clusters suggested the presence of a number of distinct surrounding stromal populations (clusters 1, 3, 4), two potential vascular populations (clusters 0 and 6) and one cluster associated with a cell

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Figure 4.17. Dissecting human RET/tdTomato+ organoids using single-cell RNA sequencing. Whole organoids were dissociated at d7+5. Single-cell RNA sequencing data for 2,207 cells was generated with unsupervised clustering carried out with Seurat package. A) Unbiased clustering using t-SNE revealed seven clusters. Cluster 2 showed strong nephric duct/UB tip+ identity. B) Marker genes regarded with nephric duct/UB tip+. Cells are coloured based on expression level.

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Top 2500 DE Collecting Duct Genes Top 2500 DE Ureteric Tip Genes

Cluster ID # Entrez Gene Gene Symbol Cluster ID # Entrez Gene Gene Symbol 2 1 2066 ERBB4 2 1 51201 ZDHHC2 2 2 8727 CTNNAL1 2 2 8727 CTNNAL1 2 3 22822 PHLDA1 2 3 387882 C12orf75 2 4 9254 CACNA2D2 2 4 157506 RDH10 2 5 3880 KRT19 2 5 9806 SPOCK2 2 6 387882 C12orf75 2 6 5455 POU3F3 2 7 8496 PPFIBP1 2 7 347733 TUBB2B 2 8 55353 LAPTM4B 2 8 2644 GCHFR 2 9 55107 ANO1 2 9 6492 SIM1 2 10 3655 ITGA6 2 10 6750 SST 2 11 9806 SPOCK2 2 11 7020 TFAP2A 2 12 5455 POU3F3 2 12 2674 GFRA1 2 13 2644 GCHFR 2 13 8828 NRP2 2 14 79191 IRX3 2 14 8829 NRP1 2 15 6492 SIM1 2 15 81539 SLC38A1 2 16 7020 TFAP2A 2 16 3975 LHX1 2 17 2674 GFRA1 2 17 84870 RSPO3 2 18 8829 NRP1 2 18 159371 SLC35G1 2 19 26751 SH3YL1 2 19 80781 COL18A1 2 20 81539 SLC38A1 2 20 3213 HOXB3 2 21 3975 LHX1 2 21 3215 HOXB5 2 22 159371 SLC35G1 2 22 3216 HOXB6 2 23 80781 COL18A1 2 23 23705 CADM1 2 24 3213 HOXB3 2 24 1946 EFNA5 2 25 3215 HOXB5 2 25 10653 SPINT2 2 26 3216 HOXB6 2 26 2719 GPC3 2 27 6546 SLC8A1 2 27 4257 MGST1 2 28 23705 CADM1 2 28 10406 WFDC2 2 29 10653 SPINT2 2 29 7849 PAX8 2 30 4257 MGST1 2 30 6319 SCD 2 31 10406 WFDC2 2 31 7345 UCHL1 2 32 56999 ADAMTS9 2 32 154043 CNKSR3 2 33 7849 PAX8 2 33 5315 PKM 2 34 6319 SCD 2 34 7108 TM7SF2 2 35 7345 UCHL1 2 35 441027 TMEM150C 2 36 7351 UCP2 2 36 9421 HAND1 2 37 116159 CYYR1 2 37 5076 PAX2 2 38 441027 TMEM150C 2 38 8660 IRS2 2 39 81618 ITM2C 2 39 216 ALDH1A1 2 40 5076 PAX2 2 40 57568 SIPA1L2 2 41 8660 IRS2 2 41 8673 VAMP8 2 42 216 ALDH1A1 2 42 5092 PCBD1 2 43 476 ATP1A1 2 43 51176 LEF1 2 44 8673 VAMP8 2 44 5629 PROX1 2 45 481 ATP1B1 2 46 5092 PCBD1 2 47 51176 LEF1 2 48 1525 CXADR 2 49 7164 TPD52L1 Table 1. Top 2500 differentially Table 2. Top 2500 differentially expressed (DE) collecting duct genes. expressed (DE) ureteric tip genes Forty- Forty-nine DE genes in the cluster 2 four DE genes in the cluster 2 matched with matched with the mouse E.15.5 collecting the mouse E.11.5 ureteric tip epithelium. duct epithelium.

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4.3 DISCUSSION

In this chapter, the capacity of the differentiation methodology described in Chapter 3 to differentiate human iPSCs towards nephric duct/UB tip lineage was further investigated. While a protocol for generating UB-like cells was previously reported by Xia and colleagues in 2013, that study had many limitations. For example, a very limited spectrum of ureteric bud genes was assessed (RET and GFRα1), no study of UB-protein markers in differentiation cultures was made, there was no evidence of a capacity of the differentiated cells to form epithelial structures, and there were no three-dimensional in vitro assays of spheroid growth/branching to prove genuine UB lineage derivation. As such, the study failed to provide evidence of many functional aspects of ureteric bud epithelium. We therefore addressed the challenge of generating a new improve methodology to derive iPSCs towards nephric duct/UB lineage. Our data is based on stringent evaluation of differentiation cultures throughout the critical embryonic stages involved in CD derivation, including anterior intermediate mesoderm, nephric duct, and its derivative ureteric epithelium. The cells subjected to this differentiation protocol showed proof of epithelialization by a three-dimensional approach, evaluation of protein markers by antibodies, as well the product of expression of two reporter lines regarded to CD lineage, and finally, by a more comprehensive gene evaluation. Although it was difficult to distinguish between whether we were patterning to a nephric duct or ureteric epithelium, and we still need to apply functional analysis to prove the identity of ND/UB lineage, the findings reported here are undoubtedly the strongest evidence to date of the directed differentiation to this embryonic renal lineage. Therefore, our results represent an important contribution to not only dissecting the lineage relationship of the nephric niche during kidney development but also recreating kidney tissue to begin the process of scaling-up one of the two renal progenitor cell types which are critical to ensuring ongoing morphogenesis.

The use of fluorescently tagged human induced pluripotent stem cells (iPSCs) provided a solid platform to standardise and evaluate this protocol by allowing us to monitor in real time the specification of the nephric duct/ureteric bud lineage through the differentiation and organoid maturation stages, and to isolate specific cell types for molecular characterisation. Since our initial assumption was that we must first generate a correctly patterned nephric duct derived from a GATA3+PAX2+PAX8+ anterior intermediate mesoderm, we initially tracked the response of the GATA3/mCherry+ iPSCs to the ND/UB protocol because of its role in forming one of the earliest core regulatory genes stimulating the expression of RET (Costantini and

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Kopan, 2010). Using this cell line, the phenotypical features of these cultures were mainly characterized by a sheet appearance which then becomes highly confluent with many multilayer zones. We also proved that cells subjected to this protocol were able to form epithelial structures by themselves in extended two-dimensional cultures, or by forming tridimensional aggregates referred to as organoids. Kidney organoids generated from this cell line display faithful reporter gene expression in live cultures subjected to fluorescence microscopy. The presence of GATA3+Cherry+KRT8+ECAD+PAX2+ETV4+ epithelial segments within organoids and two-dimensional monolayers was interpreted as an epithelium derived from the nephric duct lineage.

Organoids derived from this line were also examined at two time-points d7+5 and d7+11. Interestingly, when organoids were maintained in culture for five days with a cocktail of growth factors (FGF9/HA/tRA/9-cisRA/GDNF), they were morphologically characterised by small and abundant tubular structures within the aggregate, whereas the removal from the cocktail of FGF9 and HA at day 7+5 generated more complex ducts with well pronounced lumens, suggesting the possibility of a more mature epithelium. The withdrawal window of FGF9/HA provoked a consequent reduction in proliferation which may have then favoured convergence of different minor epithelial structures into large ducts with more mature features. Although this data was promising, and established that this was an interesting technique to encourage the derivation of a more mature epithelium, it only represents preliminary evidence which was not further examined because our major aim was to pattern to nephric duct or its derivative UB. However, upon establishment of correct ND/UB cultures, it will be interesting to attempt further modification of these cultures to direct the ND committed cells towards a more mature epithelium. We could, for example, activate sonic hedgehog and/or NOTCH pathways in our cultures. The former is believed to play a role in cell proliferation via cyclin B1 activity in the stalk+ cell population (Jenkins et al., 2007), while NOTCH signalling is critical for determination of principal cells (Guo et al., 2014, Jeong et al., 2009).

To identify the molecular signature of differentiated m-Cherry+/GATA3 cells, we examined them according to the UB/ND protocol at three time-points (day 7, day7+5 and day 7+11). To do this, we performed a time-course collection of FACS-sorted Cherry+ cells to specifically evaluate the expression of key genes which allow us to distinguish different developmental stages during collecting duct formation. Notably, our gene expression data showed high expression of genes regarded as nephric duct genes (PAX2, PAX8, GATA3) across all the time-

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While the differentiation protocol reported here did not show enough evidence to support formation of a mature collecting duct epithelium, our results have provided strong evidence supporting a clearly patterned nephric duct resembling an epithelium with ureteric bud identity. Then, to test transferability, as well as to challenge the use of the differentiation methodology reported here, a new human iPSCs line reporting for RET/tdTomato+ was subjected to the same optimisation protocol. This cell line offered the advantage of monitoring in real-time the expression of RET, widely regarded as the gold standard gene for both outgrowth and subsequent branching of the ureteric epithelium and whose expression becomes specifically restricted to the UB/tip+ region. Significantly, the ND/UB protocol showed evidence of transferability from one iPSCs (GATA3/Cherry+) to another (RET/tdTomato+), reproducing the phenotypical features of an anterior-IM differentiation defined in the results section, and also turning the expression of tdTomato+ reporter on during the first seven days of

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Future Directions

While we were not able to properly distinguish whether we were generating nephric duct or ureteric bud epithelium, essentially because there is not a clear molecular boundary between them and most of the markers identifying UB are also expressed during nephric duct formation. What is most critical with respect to kidney regeneration will be to generate an epithelium with a capacity to respond to a metanephric mesenchyme by branching. The characteristic features of a ureteric bud is its capacity to form a branching epithelium and to induce the NP to drive nephron formation. We will therefore evaluate the presumptive potential of the differentiated nephric duct/ureteric bud cells to further mature into ureteric bud structures using a three- dimensional culture system. One approach to this would be to FACS-sort cells (RET/tdTomato+ and GATA3/m-Cherry+) from cultures following the ND/UB protocol described here and seed these cells into Matrigel for culture in growth factor combinations recently reported to support the growth of freshly isolated ureteric buds from E11.5 mouse kidney (Yuri et al., 2017). These include recombinant growth factor FGF2, GDNF, RA, +/- RSPO1, and a Rho-kinase inhibitor (Yuri et al., 2017). Here we anticipate formation of epithelial structures or a branched epithelium with persistence of RET and WNT9B expression. Morphology will be assessed by immunofluorescence against the tip markers ETV4 and ETV5 (Lu et al., 2010), to discriminate between nephric duct and ureteric bud. In addition, segmentation of the ureteric epithelium will be evaluated by QPCR using the set of primers defined here (tip: RET, GFRα1, WNT11, ETV4 and Stalk: WNT9B, WNT7B, TCSTD2). Secondly, we will also evaluate the functional potential of the FACS purified presumptive ureteric bud to induce nephrogenesis. To investigate this further, we will use a re-aggregation culture, a widely described assay to assess either of two renal progenitor populations (NP or UB) (Hendry et al., 2013). Briefly, E.12.5 kidneys will be dissociated to single cell suspension, combined with either RET/tdTomato+ and GATA3/m-Cherry+, and cultured for up to 10 days. As previously reported, the recombined cells self-organize to form the ureteric bud and the nephric niche with an appropriate spatial arrangement (Hendry et al., 2013). If presumptive ureteric bud cells show the ability to incorporate into the self-organized ureteric bud, this would represent a stringent assay of ureteric bud competence. Finally, because the current objective of our laboratory is to improve our capacity to form the key cellular components to recreate kidney tissue, we also have access to presumptive nephron progenitor cells (NPs) generated from fluorescently tagged iPSCs. Taking advantage of this collaborative work, co-cultures from FACS-enriched presumptive UB and NPs will be also established by combining both

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Altogether, our research here provides an improved protocol to generate one of the two renal progenitor populations. The use of fluorescently tagged iPSCs provided a solid platform to validate the expression of RET, which is the gold standard gene restricted to ND/UB tip+, accurately allowing the isolation of a pure RET+ fraction and giving us now great confidence in using them to test the recreation of an epithelium to extensively branch when surrounded by the cap or supplemented with the right cues. Ultimately, this represents a unique opportunity to improve our understanding of human embryogenesis, as well as to generate large scale cultures of progenitor populations for tissue engineering applications.

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CHAPTER V

Concluding Remarks

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CONCLUDING REMARKS

Kidney failure constitutes a major public health issue worldwide. Diseases affecting the collecting duct system, congenital abnormalities of the kidney and urinary tract (CAKUTs), are the most frequent cause of End-Stage Renal Disease (ESRD) in children (Limwongse et al., 1999). Given that children with ESRD have a low survival rate and require renal replacement therapy, the study of the mechanisms underlying repair and regeneration of the collecting duct system will facilitate new therapeutic strategies for treating kidney disease. In recent years, enormous advances have been made in the kidney field towards the concept of cellular therapies. These include studies based on the use of stem cells to either ameliorate injury or improve repair or to recreate kidney tissue. These approaches have been discussed in this thesis. Based on the knowledge gained through advances in regenerative medicine, we published two reviews on this area entitled ‘Regenerative medicine in kidney disease’, and ‘Does renal repair recapitulate kidney development?’.

While it is well known that nephrons are not able to regenerate after birth, the kidney is able to boost proliferation and repair in response to injury. One cell-based therapy is treatment with mesenchymal stem cells, which have been reported to ameliorate damage and reduce allograft rejection via their immunomodulatory capacity and their release of paracrine factors (Imberti et al., 2007, Aggarwal et al., 2005, Krampera et al., 2003, Di Nicola et al., 2002, Stagg et al., 2007). However, no evidence of a structural contribution of. Previous work from this laboratory has reported a capacity for kidney-derived MSC-like cells to both arise from the collecting duct epithelium and then reinsert into this segment of the renal epithelium (Li et al., 2015). The functional value of this observation was yet to be demonstrated. In the first part of this thesis we therefore developed a model in which to test the putative functional integration of endogenous kidney MSCs isolated from the adult murine kidney. Integration in this instance should have recapitulated autosomal dominant polycystic kidney disease (ADPKD). To this end, k-MSC lines defective for the Pkd1 gene were generated. While the deletion between exon 2-4 in the Pkd1 gene did not produce apparent defects in MSC properties, the mutation did lead to a detectable reduction in the number of primary cilia. Given that micro-injection of double transgenic k-MSCsPkd1del2-4/TMTO+ did not produce significant integration in the recipient mice, our research ruled out the possibility that k-MSCs represent a preferable population for effecting repair. However, we showed the capacity to generate k-MSCs with mutant genes,

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Additionally, with the discovery of induced pluripotent stem cells and the rapid evolution of techniques for their directed differentiation, this provides the prospect of regrowing of a whole new organ. Since 2013, we have seen significant advances in kidney regeneration, with a variety of studies reporting the generation of kidney cell types from pluripotent stem cell sources. Among these, our laboratory pioneered the generation of kidney organoids from hPSCs. It is because of the potential importance of this approach for therapeutic uses that in this thesis we have addressed some of the limitations of this technique. We have therefore developed an improved protocol to direct the differentiation of iPSCs towards nephric duct/ureteric bud lineage, with the long-term objective of generating epithelium which responds to a metanephric mesenchyme with branching activity. As we had access to unique reporter lines, using CRISPR/Cas9 gene editing of hPSCs as a readout of differentiation cultures, and because the success of any new protocol relies on being transferable to different cell lines and laboratories, we successfully adapted the protocol reported in Chapter Three to feeder-free conditions. This new protocol showed robust evidence of patterning to early nephric duct. Using the GATA3/mCherry+ reporter line, our QPCR data showed robust expression of PAX2, PAX8, GATA3, GFRα1, and RET alongside down-regulation of metanephric mesenchymal (MM) markers, which demonstrated its specificity for ND/UB lineage specification. The cells subjected to this differentiation protocol showed proof of epithelialisation using both two- and three-dimensional immunofluorescence, clearly showing that this methodology does promote formation of early nephric duct epithelium and no other nephron segments. Given the promise of this data, and because it involved the development of new techniques, this protocol is currently being evaluated for patenting as an invention.

We then also proved the transferability of the ND/UB protocol using another reporter cell line, the RET/tdTomato+ iPSC line. Of significance, and in line with our previous data, the phenotypic features of an anterior-IM differentiation defined for the GATA3/mCherry+ reporter line were reproducible, as was the response of the reporter tdTomato+, which turned on during the monolayer-phase of the protocol between days 5 and 6. The RET/tdTomato+ line also displayed the ability to grow organoids which form RET+ECAD+GATA3+ epithelial structures. This cell line offered the advantage of real-time monitoring of RET expression, which is detected throughout the nephric duct as early as E.9.5, and then becomes specifically

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Finally, it is also worth mentioning that, at the time of finishing writing up this thesis, Taguchi et al. (2017) reported a multi-step protocol to direct mESCs/iPSCs towards UB lineage, and they claimed that specification for each progenitor lineage (MM and UB) requires distinct cues. Interestingly, they first investigated the cues required to commit undifferentiated cells to UB lineage using Wolffian Duct (WD) sorted precursor cells from E8.75 to E11.5. In their study they established the combination of factors, including RA, Wnt, FG9, and GDNF, as optimal for induction of Pax2, Ret, Hnf1b, Wnt9, Calb1, and Wnt11 expression. Based on this, the authors established a multi-step protocol using mESCs which uses most, but not all, of the growth factors used in our protocol. However, the exposure time and concentrations are quite different. The protocol of Taguchi et al. specifies sorting of the committed WD cells at day 6.5 by Cxcr4 /Kit, based on their mouse data (E8.75 WD). It then used differentiated mESCs for organoid reconstruction, using iUB, iNP, and embryonic NP. This procedure provided the first strong evidence of successful interaction between both progenitor populations, evidence of high levels of organisation within the reconstructed organoids, and evidence of branching of the iUB. This protocol was next transferred to the differentiation of human iPSCs. While

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Taken together, we concluded that the methodology reported here does promote the differentiation of human iPSCs towards cell types in which the nephric duct/ureteric bud genes have not only switched on, but are even organizing themselves into unique epithelial structures resembling the nephric duct epithelium of a developing human kidney. The use of fluorescently tagged human iPSCs provided a solid platform to validate the response of ND/UB genes to this protocol, accurately allowing the isolation of pure fractions. This study therefore has established a basis to begin the study of the capacity of this epithelium to respond to a metanephric mesenchyme with branching activity. This will in the long term contribute to our capacity to recreate kidney tissue and develop cellular therapies, as well as assisting disease modelling and drug screening.

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CHAPTER VI

Material and Methods

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6.1 MATERIALS AND METHODS

6.1.1 Mice

The care and experimental use of mice conformed to the guidelines of the Animal Ethics Committee at the University of Queensland, and was then re-assessed and approved by the Animal Ethics Committee at Murdoch Research Institute.

Genotyping of Rosa26TdTomatoFlox/Flox (Stock number: 007909) and Pkd1Flox/ Flox (Stock number: 010671) mice was performed according to the Jackson Laboratory website protocols (https://www.jax.org/strain/007909) and (https://www.jax.org/strain/010671) respectively.

6.1.2 Neonatal Injection Model

To determine the capacity of different k-MSCs lines to integrate into kidney compartments, microinjection experiments were carried out. The neonatal injections were given into the nephrogenic zone of the neonates at post-natal day 1 (PD1) of outbred CD1 (Chapter2, AIM1), SWISS (Chapter2, AIM3) and C57BL/6 (Chapter2, AIM3), using a microinjection pipette and following a protocol previously described by Li et al., 2015. Briefly, neonates were anaesthetized and a small incision in the skin was made. Cells were mixed with Fluoresbrite Yellow Green microspheres (2.0 mm; Polysciences Inc.) at a ratio of 1:20, and then resuspended at 0.5-1x107/ml in PBS. Using an Eppendorf microinjector ~ 5,000-7000 cells were microinjected into the kidney. Finally, kidneys were harvested at three different times- 4, 6, and 10 weeks for AIM1, and then 1, 6, and 12 months for AIM2 post-injection. In all the harvested mice, successfully injected kidneys were selected based on the presence of the coinjected fluorescent microspheres.

6.1.3 Quantification of cell integration:

For quantification of the proportion of medullary collecting duct cells derived from injected kidney MSC–like cells, kidney samples were examined from three independent neonatal injection experiments in which cells of interest were delivered into CD1 neonates at PND1 and then collected at 4, 6, and 10 weeks post-delivery for analysis. Two to three sections were chosen from each sample and stained with antibody to detect injected cells (anti- GFP) and 160

Commercial in Confidence collecting duct epithelial cells (anti-Aqp2). All images (300 dpi) were captured with a fluorescence microscope (Olympus BX-51) and uploaded into Imaris software (version 7.2; BitPlane AG) for quantification. Imaris software was then used to create masks on the basis of specific staining (Aqp2, GFP), and spot counting was performed. The number of nuclei positive for GFP+ staining was counted separately and used to calculate integration efficiency (GFP+/Aqp2+).

6.1.4 Isolating and culturing the intact metanephric kidney

Pregnant females from the Hoxb7/myr-Venus strain were humanely sacrificed according to the guidelines of the Animal Ethics Committee at MCRI. After mice dissections, metanephric kidneys at 11.5 dpc were placed in a 60-mm petri dish with plain CO2-independent media. Only the kidneys positive for Hoxb7 were selected for further culturing, using a fluorescent dissection microscope. The branching morphogenesis processes of these kidneys were followed up until 96 hours. Kidneys were cultured using the basal medium DMEM serum free, 0.5 % P/S supplemented with different concentrations of growth factors. Two different concentrations of GDNF- 50µg/ml and 150 µg/ml- were assessed with and without supplementation of 2.5 µM AGN.

6.1.5 Cell culture

All cells were grown and maintained in a 5% CO2-controlled incubator at 37°C. Culture conditions, media, and special requirements for each cell line are detailed below.

6.1.5.1 Cell Isolation from adult kidney tissue and maintenance of k-MSCs lines

Kidney MSCs-like cells were isolated from Pkd1Flox/Flox and Pkd1Flox/ Flox; Rosa26TdTomatoFlox/Flox mice. Male mice at 8 weeks of age were used for both isolations from two biological replicates, where one biological replicate represents isolation from a single animal (two kidneys). The Pkd1 mice were obtained from a collaboration with Dr Ben Hogan from the Institute of Molecular Bioscience (IMB at University of Queensland). Kidney MSC-like cells were derived as bulk cultures and cells were then grown as standard cultures, as previously described (Fiedenstein et al., 1974). Briefly, kidneys were dissected and subjected to enzymatic digestion (Collagenase B, 1mg/ml; Dispase II, 1.2 unit/ml), followed by two rounds of filtration with a

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6.1.5.2 iPSC and culture on mouse embryonic feeders (MEFS) iPS cell line CRL1502 (clone C32)

The wild-type human iPSC line CRL1502 (clone C32) was generated using episomal reprogramming, and tested for the mycoplasma infection (Briggs, J. A. et al. 28). Undifferentiated human iPSCs were maintained on the mouse embryonic fibroblasts (MEFs) (Millipore) as a feeder layer with human ES cell (hESC) medium. Human iPSCs were plated on a Matrigel-coated (Millipore) culture dish and cultured in MEF-conditioned hES medium (MEFCM) until reaching 60–100% confluence. After this, cells were detached using Tryple Select and plated on Matrigel for differentiation in feeder-free condition, with MEF- conditioned medium to avoid any unknown effect of MEFs during differentiation.

6.1.5.3 Generation of iPSC reporter lines and culture on a feeder layer-free culture

The generation of kidney organoids previously reported by our laboratory was based on iPSCs lines derived and maintained on mouse embryonic feeders (MEFs). However, because our current goal- optimization of the key component cell types required for nephric niche formation- requires transferability between cell lines, the two iPSC reporter lines generated for this thesis are on a feeder layer-free system. The generation of these lines was carried out by a postdoctoral researcher in our laboratory, Dr. Sara Howeden.

1502.3 GATA3: m-Cherry Clone 60 and 2429 GATA3: m-Cherry clone 26

In order to improve, manipulate, and monitor collecting development in kidney organoids, two iPSC lines for the GATA3 gene were developed, using CRISPR/Cas9 gene editing on hPSCs. These lines were generated by the same target gene but through different approaches. Firstly, 1502.2 GATA3: m-Cherry clone 60 was generated by a sequential approach, and later the GATA3: m-Cherry clone 26 iPSC line was derived from a normal foreskin fibroblast line

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(ATCC# CRL-2429) through a simultaneous reprograming and gene editing approach described by Howden et al. 2015. Briefly, it is a two-step approach: firstly, episomes encoding reprogramming factors are transfected into fibroblasts to create iPSCs, which are then transfected with the homologous reporter cassette and CRISPR/Cas9 RNA; and secondly, in a simple one-step procedure, episomal vectors encoding reprogramming factors, plasmids encoding guide RNA for the target gene, the homologous reporter cassette, and RNA encoding CRISPR were simultaneously transfected into fibroblasts via electroporation (Figure 6.1). Two to three weeks after transfection, individually induced pluripotent stem cell (iPSC) colonies were picked and passaged separately. Correctly targeted clones were identified by PCR analysis and confirmed by Sanger sequencing. Molecular karyotyping of targeted iPSC lines was performed using Illumina SNP array to assess genomic integrity. These lines were also checked for mycoplasma.

Figure 6.1: Reprogramming and gene targeting of human fibroblasts to generate reporter lines. Orange dashed arrows denote the two-step approach; episomes encoding reprogramming factors are transfected into fibroblasts to create iPSCs, which are then transfected with the homologous reporter cassette and CRISPR/Cas9 RNA. Blue solid arrows denote the simultaneous approach; reprogramming episomes, the homologous cassette, and CRISPR/Cas9 are transfected into fibroblasts simultaneously. Both methods create a mix of targeted clones, with the reporter, and untargeted clones, illustrated here with constitutive eGFP expression. The simultaneous approach makes it easier to select clonal populations for screening, as shown by the distribution of eGFP+ cells in the iPSCs produced by the simultaneous approach, compared to the two-step approach. (Howden, S. et al., 2015)

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The iPSC line h-RET-F7 was obtained from a collaboration with Dr. Sanjay Jain at Washington University. This reporter iPSC line was derived from an archived parent WTC11 iPSC cell line expressing the red fluorescent protein from the RET locus. This cell line has no cytopathic effects or signs of any infectious microorganism, and is also mycoplasma free.

6.1.5.4 iPSC culture on a feeder layer-free culture iPSC reporter lines were plated on vessels pre-coated with Geltrex LDEV-Free hESC-qualified (Life Technologies), and grown in Essential 8TM medium with daily media changes. Cells were typically passaged every 3 days with EDTA 0.5 mM in PBS, as previously described (Chen et al., 2011).

6.1.5.5 Standard differentiation protocol of iPSC lines to form kidney organoids.

In 2016, our research group reported a successful stepwise protocol for the differentiation of iPSC to form kidney organoids (Taksato et al., 2016). Briefly, undifferentiated iPSCs were seeded onto Matrigel-coated vessels at the cell density of 15x10 3 cells/cm2 in condition medium containing bFGF10 ng/ml for cell line cultures on MEFS or grown in Essential 8TM medium with 5 μM Y27632 for cells on a feeder layer-free culture system for 24 hours. The next day, cells were incubated with 8μM CHIR99021 in APEL (STEMCELL Technologies) for 4 days. Then the medium was aspirated, and cells were treated with FGF9 (200ng/mL) and heparin (1μg/mL) in the same basal medium for five days, refreshing the media every two days. After seven days of differentiation, the cells were detached using Trypsin EDTA (0.05%) at 37°C for 3 minutes to make organoids according to Taksato et al., 2016. Aggregates were made by centrifuging three times at 400xg for 3min, and were gently picked up using a wide-bore pipette tip. They were placed onto a transwell 0.4μm pore polyester membrane (Corning) and pulsed with 8μM CHIR99021 in the basal medium for 1 hour. The medium was then changed to APEL with FGF9 (200ng/mL) and heparin (1μg/mL) for five days, refreshing the medium every two days. Finally, organoids at d7+5 were cultured in the basal medium APEL until harvest.

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6.1.6 Transfection and Antibiotic Selection

K-MSCs were plated on a 35-mm tissue culture plate and cultured until confluent. In order to induce ex vivo CRE recombination, a transfection protocol was then implemented using a plasmid containing CMV-CRE-IRES-PUROMICIN gene cassette and lipofectamine LTX (Invitrogen). Three different conditions of DNA/lipofectamine ratio were tested: condition1, 1% lipofectamine + 15ugDNA; condition 2, 0.5 % lipofectamine + 15ugDNA; and condition3, 0.5 % lipofectamine + 15ugDNA. The transfection media were replaced after three days by αMEM supplemented with puromycin at 2ug/ml for four days. Cells were then expanded in α- MEM with 20% FBS 2 % P/S.

6.1.7Clonal Selection and Expansion of k-MSCs Mixed Population

In order to obtain k-MSCs mutant for both alleles of the Pkd1 gene, mixed populations of k- MSCs were subjected to clonal isolation by manual isolation, and single cells were seeded to 100 mm petri dishes by limited dilution seed (1x103-1x104). After eight days, clones were isolated via ring cloning and transferred to a 24-well and a 6-well plate. After four weeks of culture, each clone was screened by PCR using primers F4 and R4, as previously described.

6.1.8 Cilia induction

Ciliation was induced on k-MSCs lines by reducing the amount of serum in the basal medium to 0.05 % for 24 hours. Serum starvation prompts the cells to exit the cell cycle and extend a primary cilium.

6.1.9 MRI

MRI scans were carried out at The Florey Institute of Neuroscience and Mental Health. Mice were anaesthetised with isoflurane 3-5% and transferred to a specialist MRI bed with integrated circulating heating and anaesthetic nose cone. Isoflurane was then reduced for maintenance to 0.5-2.5 %. Body temperature, breathing rate, and heart rate were monitored throughout the procedure. The acquisition consisted of T2-weighted RARE sequence and T1-integrated FLASH sequence. A matrix of 256 x 256, and 0.5 mm and interslice gap of 0.5 mm coronal slices, a TR/TE of 80/1.905 ms a FOV 3x3. Cyst formation was also confirmed by histology.

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6.1.10 Cystic index

Kidneys were harvested and then embedded in OCT. Samples were frozen and kept at -80°C. 10 µm sections were cut and stained with haematoxylin eosin. Two sections were analysed using Image J software to calculate the percentage of the cystic kidney area compared to the total kidney area.

6.1.11 Statistical analysis

All statistical analyses were performed using the GraphPad Prism software package. Statistical significance was assessed using an unpaired (two-tailed) Student's t-test. P-values are represented in the Figures as *= P ≤ 0.05, **= P ≤ 0.01, ***= P ≤ 0.001, and ****= P ≤ 0.0001.

6.1.12 Imaging

Imaging was performed using a variety of microscopes. A Leica M165 FC stereomicroscope was used for isolation and fluorescent imaging of metanephric kidneys. A Leica M165 FC was used for brightfield images of kidney tissue. A Nikon eclipse Ts2 was used for brightfield images of cell monolayer and organoids. An LSM 780 confocal was widely used for immunofluorescence of adherent cells, kidney organoids, and sections. ZEN and IMARIS software packages were used for exporting images into TIFF documents.

6.1.13 Immunofluorescence:

Details of all antibodies and dilutions used are presented in Table X.

Immunofluorescence of tissue cryosections

To confirm that the integration of k-MSCs is still detectable and restricted to collecting duct structures, immunofluorescence analyses were performed. Kidneys were fixed in 4% PFA, dehydrated in 30% sucrose/PBS overnight, and then embedded in OCT medium. Samples were frozen in liquid nitrogen and stored at 80°C. Ten m sections were cut and then post-fixed in 4% PFA for 10 minutes at room temperature, washed three times (5 minutes each) with PBS 1X, and incubated in blocking buffer (10% donkey serum/0.3% Triton-X/PBS) for 1 hour at 166

Commercial in Confidence room temperature. Primary antibodies were prepared in blocking buffer. Sections were then incubated in primary antibodies for 2 hours, and then washed in PBS 1X three times for 10 minutes each. Secondary antibodies were diluted and blocking buffer and tissue sections were incubated in a darkened staining chamber for 1 hour at room temperature. The secondary solution was aspirated and washed three times for 10 minutes each time. Sections were then stained with 4', 6-diamidino-2-phenylindole (DAPI), 0.25μg/mL diluted in PBS, for 10 minutes. Finally, sections were washed with 1x PBS, rinsed with water, and mounted with Dako Fluorescent Mounting Medium (Dako).

Immunostaining of adherent cells

After culture, cells were fixed in 4% PFA for 10 minutes at 4°C. Fixative was removed and cells washed three times with PBS 1X, 10 minutes each time. Cells were then incubated with blocking buffer (10% donkey serum/0.3% Triton-X/PBS) for 1 hour. Primary antibodies were diluted in the same blocking buffer and applied for 2 hours at room temperature, or overnight at 4°C. Cells were then washed in 0.1% PBTX three times, 10 minutes each, followed by incubation with secondary antibodies for 1 hour in the dark at room temperature. Cells were then incubated with 4', 6-diamidino-2-phenylindole (DAPI), 0.25μg/mL diluted in PBS, for 10 minutes to stain nuclei, and subsequently washed with PBS 1X three times for 10 minutes each time.

Organoids on tranwell filters

Immunofluorescence assay was performed according to Takasato et al., 2016. Briefly, once organoids were collected, they were fixed with paraformaldehyde 2% at 4°C for 20 minutes and subsequently washed in PBS 1X three times. Organoids were then removed from the plate by cutting the surrounding transwell filter with a scalpel and then using tweezers to remove them to a 24-well plate and submerge them in blocking buffer (10% donkey serum/0.3% Triton-X/PBS) for 3 hours. Organoids are incubated in primary antibodies, and prepared in blocking buffer at 4°C overnight. The next day, this solution is removed, and organoids are washed in 0.1% Triton-X100 or Tween/PBS six times, for 10 minutes each time. Organoids are then incubated with secondary antibodies of choice in PBTX + DAPI (1:000) on a rocking platform at 4°C overnight, and subsequently washed 6 times, 10 minutes each time, with PBS

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1X. After this, organoids are carefully transferred onto MatTek Glass bottom dishes (MatTek Corporation) for imaging.

Antibody Catalogue # Source Host Dilution Primary Antibodies anti-NPHS1 AF4269 R&D Systems Sheep 1/300 anti-E-Cadherin 610181 BD Biosciences Mouse 1/300 anti-Pax2 71-6000 Zymed Laboratories Inc Rabbit 1/300 anti GATA3 AF2605 R&D Systems Goat 1/300 anti GATA3 5852S Cell signalling technology Rabbit 1/300 anti-LTL-biotin B-1325 Vector Laboratories NA 1/300 anti-ARL13b N295B/66 Neuromab, Antibodies Inc Mouse 1/300 anti-Aqp2 AB3274 Millipore Australia Rabbit 1/300 anti-Vimentin V2258 Sigma Mouse 1/300 anti-DBA B-1035 Vector laboratories NA 1/300 anti-GFP Ab13970 Sapphire Bioscience Chicken 1/300 Developmental studies anti-KRT8 TROMA-1-S Rat 1/300 hybridoma bank anti-Laminin 5 Ab77175 abcam Mouse 1/300 Anti-RFP M208-3 MBL (Jomar life science) mouse 1/400 MBL Medical & Biological Anti-RFP PM005 Rabbit 1/400 laboratories Secondary Antibodies Alexa 405 Streptavidin S32351 Invitrogen Life Technology Donkey 1/400 Alexa 488-anti Mouse A21202 Invitrogen Life Technology Donkey 1/400 Alexa 488-anti Goat A11055 Invitrogen Life Technology Donkey 1/400 Alexa 488-anti Rat A21208 Invitrogen Life Technology Donkey 1/400 Alexa 488-anti Chicken A21208 Invitrogen Life Technology Goat 1/400 Alexa 488-anti Rabbit A11008 Invitrogen Life Technology Donkey 1/400 Alexa 568-anti Rabbit A10042 Invitrogen Life Technology Donkey 1/400 Alexa 568-anti Mouse A10037 Invitrogen Life Technology Donkey 1/400 Alexa 647-anti Mouse A31571 Invitrogen Life Technology Donkey 1/400 Alexa 647-anti Goat A21447 Invitrogen Life Technology Donkey 1/400 Alexa 647-anti Sheep A21448 Invitrogen Life Technology Donkey 1/400 Alexa 647-anti Rabbit AB175704 Abcam Donkey 1/400 Table 3. List of antibodies used for immunostaining.

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6.1.14 Flow cytometry and cell sorting

6.1.14.1 Fluorescent activated cell sorting (FACS)

Kidney tissue, organoids, and cells cultured in monolayer were subjected to fluorescent activated cell sorting using a BD FACS Aria cell sorter. Here we describe the dissociation protocol for each sample.

Kidney samples

Kidneys were dissected and subjected to enzymatic digestion (Collagenase B, 1mg/ml;Dispase II, 1.2 unit/ml) followed by two rounds of filtration with a cell strainer (70 and 40 µm; BD Falcon) to produce single-cell yields. FACS for tdTOMATO+ sorting was performed at the Queensland Brain Institute for isolation of cells from double transgenic mice Pkd1Flox/ Flox; Rosa26TdTomatoFlox/Flox (University of Queensland) (FACS Aria, BD). Therefore, the tdTOMATO+ population was selected based on stringent criteria of size and granularity, as well as upon single cell discrimination. Cells were then collected and were kept at 4°C until seeding.

Kidney organoids

For dissociation of whole organoids, Accutase was added to the organoids, pipetting a few times to begin dissociation, followed by incubation at 37°C for 10 min, and pipetting every 3 mins. This cell suspension was then passed through a 40 µm filter two times, transferred into a 15-ml conical tube, and pelleted by centrifugation at 1500 rpm for 3 minutes. Supernatant was discarded, and the pellet was washed with cold FACS buffer (1% FBS in PBS). Cells were finally spin down again, resuspended in ~ 500μL FACS buffer, and kept at 4°C until RNA extraction. Sorting for this protocol was performed for m-Cherry GATA3 and RET- tdTOMATO (GCamp6) iPSC lines in FACS Aria, BD. Cell suspension of non-fluorescent organoids derived from the same parental line (2429) were used as negative control to get out the events which did not represent a real signal of m-Cherry+. Dead cells were removed by setting up the forward and scatter gates prior sorting of the m-Cherry+ fraction as well as by staining with propidium iodide.

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Cells in monolayers

Cells were initially washed two times with PBS 1X and then incubated in 0.05% trypsin/EDTA (Gibco) for 2 min at 37°C. Trypsin was inactivated with DMEM-FBS10% and the cell suspension was passed through a 30 µm filter. Cells were pelleted by centrifugation at 1500 rpm for 3 minutes. Supernatant was discarded, and the pellet was washed with cold FACS buffer (1% FBS in PBS). Cells were spin down again, resuspended in ~ 500μL FACS buffer, and kept at 4°C until RNA extraction. Sorting for this protocol was performed for m-Cherry GATA3 clone 26 iPSC cell line in FACS Aria, BD.

6.1.14.2 Flow cytometry k-MSCs lines were subjected to flow cytometry at passage 11. Briefly, freshly isolated kidney- derived MSCs were fixed in 1 ml 4% PFA for 5 min at 4C, washed with FACS buffer (1% FBS in PBS), and then stained with directly conjugated antibodies for 30 minutes at 4C (PE- CD24, APC-CD44, PE-CD49e, PE-CD51, PE-CD81, PE-CD140a, PE-CD140b, and PE-Sca- 1; BD Falcon). Cells were then washed with FACS buffer two times, centrifuged at 1500 rpm for 3 minutes, and then analysed on FACSCantoII (BDFalcon). Data were collected and analysed using FACSDiva software (BD Falcon) and FlowJo (version 7.6.5; Tree Star).

6.1.15 RNA extraction, cDNA preparation, qRT-PCR and RT-PCR.

RNA isolation and reverse transcription

Total RNA was extracted using either PureLink RNA mini kit (cat # 12183018A, Life Technologies) or Isolate II RNA micro kit (cat # BIO-52075, Bioline) as per manufacturer’s instructions. RNA was quantified using a NanoDrop 1000 spectrophotometer (Thermo Fisher Scientific). Equal amounts of RNA were used to make cDNA using the GoScript reverse transcription system (Promega), following manufacturer’s instructions.

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Quantitative PCR were performed using GoTaq qPCR master mix (cat # A6001, Promega). The reaction was conducted on an ABI Real-time PCR (Applied Biosystems) machine. Amplification was performed in triplicate in a MicroAmp 96-well optical reaction plate, with 20 μl reaction samples. Primer sequences are listed in table X. cDNA levels of target genes were analysed using comparative Ct levels, normalized to glyceraldehyde-3-phosphate dehydrogenase and expressed as relative transcript abundance (ΔCt or 2−ΔCT), as well as fold change relative to control (ΔΔCT or 2–ΔΔCT) (Schmittgen et al., 2008). Error bars represent S.E.M. calculated from independent biological replicates; statistical significance was assessed using unpaired (two-tailed) Student's t-test. P-values are represented in the Figures as *= P ≤ 0.05, **= P ≤ 0.01, ***= P ≤ 0.001 and ****= P ≤ 0.0001. Standard error of the mean is represented in error bars. Conventional RT-PCR analysis was performed using One Taq DNA polymerase (NEB) as per the manufacturer’s instructions. Primer sequences for qRT-PCR and RT-PCR are supplied in table X and for in Table Y respectively.

Primer Forward (5'-3') Reverse (5'-3') GAPDH AGCCACATCGCTCAGACAC GCCCAATACGACCAAATCC RET TAACCGGAGCCTGGACCATA GAAGGGATGTGGGTGACAGG PAX2 GCAACCCCGCCTTACTAAT AACTAGTGGCGGTCATAGGC WNT11 TGAAGGACTCGGAACTCGTC CGCTTCCGTTGGATGTCTTG HOXB7 CGACACTAAAACGTCCCTGC GCGAAAACCGAACTTGAGGC GATA3 GCCCCTCATTAAGCCCAAG TTGTGGTGGTCTGACAGTTCG SIX2 AGGCCAAGGAAAGGGAGAAC AGCTGCCTAACACCGACTTG EYA1 ATCTAACCAGCCCGCATAGC GTGCCATTGGGAGTCATGGA HOXD11 GCCAGTGTGCTGTCGTTCCC CTTCCTACAGACCCCGCCGT ETV4 AACAGACGGACTTCGCCTAC TCGCAGAGGTTTCTCATAGCC ETV5 TCGGGATTACTGCGTCGATTC AAAAACCTTCATGGCTGCTGG GFRA1 CCACTCATGTTTTGCCACCG ACAGAGGTGTGTATTGCCCG PAX8 CGATGCCTCACAACTCCATC GAGGTCTGCCATTCACAAAGG WNT9B GCTTGAGTGCCAGTTTCAGTT AGGAAAGCTGTCTCTTTGAAGC WNT7B TCCCTGGATCATGCACAGAAAC ATGACAGTGCTCCGAGCTTCAC TACSTD2 TCACGCTTCCTGATTCCTCG ACCCTGAGGCCAGGATCTAT AQP2 ATCCATTACACCGGCTGCTC TCCAGAAGACCCAGTGGTCA UPK3A GACTTTCGCCACCAACAACC TTCCTGGAAATGGCTGAGTCG Table 4. List of primers used for qRT-PCRin this thesis.

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Primer Forward (5'-3') Reverse (5'-3') Pkd1 Fr4-Rv4 CCT GCC TTG CTC TTT CC CCT GCC TTG CTC TTT CC tdTMTO Wt AAGGGAGCTGCAGTGGAGTA CCGAAAATCTGTGGGAAGTC tdTMTO Mt GGCATTAAAGCAGCGTATCC CTGTTCCTGTACGGCATGG Table 5. List of primers used for genotyping by RT-PCR.

6.1.16 Data generation and analysis for single-cell RNA sequencing

Files returned from the sequencing facility were processed using the Cell Ranger pipeline (v1.3.1). The resulting expression matrix was loaded into the R statistical programming language using the scater package (McCarty et al., 2017). Quality control was performed using scater and low-quality cells were filtered out based on how many genes were expressed and the percentage of counts assigned to mitochondrial and ribosomal genes. Genes were removed if they had less than two counts in total, were expressed in less than two cells or did not have an associated HGNC symbol. Mitochondrial genes, ribosomal genes and genes associated with the cell cycle were also removed. The final filtered dataset contained 11972 genes and 2207 cells.

The filtered dataset was analysed using the Seurat package (Satija et al., 2015). The genes used in the analysis were chosen based on their expression and variability with 1554 genes being selected. Factors associated with the total counts in each cell, percentage of genes with zero expression and percentage of counts assigned to mitochondrial or ribosomal genes were regressed out of the dataset and the first 15 principal components selected to use for clustering. Cells were clustered at a range of resolutions but a resolution of 0.5 was chosen as it was the lowest resolution after which the number of clusters did not change. Marker genes for each cluster using a likelihood-ratio test to test for differential expression between one cluster and all other cells. Gene ontology terms associated with the top 100 positive marker genes for each cluster were tested using the goana function (Young et al., 2010) in the limma package (Ritchie et al., 2015).

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Minerva Access is the Institutional Repository of The University of Melbourne

Author/s: Kairath Oliva, Pamela Andrea

Title: Unveiling the biology of collecting duct epithelium repair and regeneration

Date: 2017

Persistent Link: http://hdl.handle.net/11343/213434

File Description: Unveiling the biology of collecting duct epithelium repair and regeneration

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