Trophoblasts: Methods and Protocols (Methods in Molecular Biology, 2728) 1071634941, 9781071634943

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Methods in Molecular Biology 2728

Sandeep Raha  Editor

Trophoblasts Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

For further volumes: http://www.springer.com/series/7651

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-by step fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Trophoblasts Methods and Protocols

Edited by

Sandeep Raha Department of Pediatrics, McMaster University Medical Centre, Hamilton, ON, Canada

Editor Sandeep Raha Department of Pediatrics McMaster University Medical Centre Hamilton, ON, Canada

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3494-3 ISBN 978-1-0716-3495-0 (eBook) https://doi.org/10.1007/978-1-0716-3495-0 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A. Paper in this product is recyclable.

Preface The placenta is a fascinating organ that contributes to physiological changes in both the mother and fetus. Over the last two decades, our knowledge of placental physiology and contributions to maternal and fetal physiology has advanced significantly. While the placenta is a disposable organ, it undergoes dramatic changes from the time of implantation to parturition. Its structure and function have been shown to be impacted by maternal nutrition and lifestyle, environmental chemicals, and maternal immune status, along with a number of other factors. These factors can alter the function of trophoblasts, the stem cell lineage that comprises a significant portion of placental tissue. The study of trophoblasts, and the placenta, is challenging because these cells and tissues cannot be ethically sampled until the end of pregnancy or elective termination. Furthermore, tissue sampling also presents challenges as tissue functions deteriorate dramatically once removed from the uterine environment. Therefore, isolation of trophoblasts and their culture is a considerable methodological skill that enables the elucidation of mechanistic pathways ex vivo. In this volume, Chaps. 1, 2, and 3 outline various approaches to the study and manipulation of primary trophoblast cells. In addition to primary cells, there are many commercial sources available for trophoblast cells of various sublineages. These cell lines also expand the ability of researchers to ask a variety of mechanistic questions related to cell function. In this volume, we explore a number of approaches that are useful in assessing trophoblast angiogenesis, transport function, cellular respiration, migration, and invasion. An important challenge in addressing placental dysfunction is targeting the placenta and trophoblast cells in order to deliver therapeutic treatments. Chapters 14 and 15 demonstrate some strategies using liposomes to deliver biomolecules to trophoblasts. Along with targeted placental therapy, a focus of future research is to better understand the complex 3D interactions between trophoblast cells in the uterine environment. To address this, research focusing on developing advanced in vitro models of the placenta that emulate the cellular interactions in the 3D uterine environment is a valuable tool in the arsenal of the reproductive biologist. Such technologies are necessary to better understand cell-to-cell interactions between trophoblasts, as well as between trophoblasts and other cell types resident in the placenta. In this volume, Chaps. 16, 17, 18, and 19 address approaches to studying models comprised of heterogenous cell types and 3D-organoids structures that maybe more representative of cell-to-cell interactions in vivo. Taken together, this volume brings together a set of protocols that will facilitate research to better understand the function of the placenta and its complex contributions to maternal health and fetal development, and provide greater insights to the area of study known as the developmental origins of health and disease (DoHaD). I would like to express my gratitude to all the authors have made this book possible. I would also like to thank Professor John Walker, the Series Editor, for the invitation to edit this volume. Finally, I would like to extend a huge thanks to Amrita Debnath, for her excellent scientific editing skills in the preparation of this volume. Hamilton, ON, Canada

Sandeep Raha

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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

PART I

ISOLATION AND CULTURE METHODS

1 Isolation and Maintenance in Culture of Primary Human Trophoblast from Term Placentae. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yohanes N. S. Nursalim, Katie M. Groom, Cherie Blenkiron, and Lawrence W. Chamley 2 Isolation of Human Villous Cytotrophoblastic Cells from Term Placenta for Transfection. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Caroline Toudic and Benoit Barbeau 3 Enzymatic Digestion and Single Cell Isolation of Peri-implantation Stage Human Trophoblast Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Deirdre M. Logsdon, Toshihko Ezashi, and Ye Yuan 4 Precision-Cut Slice Culture Method for Rat Placenta . . . . . . . . . . . . . . . . . . . . . . . . Fusun Gundogan, Jeffrey Gilligan, and Suzanne de la Monte

PART II

v ix

3

13

25 35

ANALYSIS OF FUSION

5 Evaluation of Molecular Interactions and Cellular Dynamics at the Maternal-Fetal Interface During Placental Morphogenesis . . . . . . . . . . . . . 45 Madhurima Paul and Rupasri Ain 6 In Vivo Quantitative Assessment of Gestational Choriocarcinoma Development and Progression Using Luminescent Trophoblast Cells . . . . . . . . . 77 Wael Traboulsi, Deborah Reynaud, Roland Abi Nahed, Fre´de´ric Sergent, Nadia Alfaidy, and Mohamed Benharouga 7 Analyzing Trophoblast Fusion Using Immunofluorescence and Split Protein Complementation Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 Adam Jaremek and Stephen J. Renaud 8 Quantification of Trophoblast Syncytialization by Fluorescent Membrane Labeling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 Yang Zhang and Huanghe Yang 9 Assessment of Placental Sodium-Independent Leucine Uptake and Transfer in Trophoblast Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Jonas Zaugg and Christiane Albrecht 10 Assessing Cholesterol Efflux on Primary Human Trophoblast Cells . . . . . . . . . . . 123 Barbara Fuenzalida and Christiane Albrecht 11 RGD-Based Fluorescence to Assess Placental Angiogenesis . . . . . . . . . . . . . . . . . . 131 Veronique Josserand, Jonathan Lavaud, Michelle Keramidas, Constance Collet, Wael Traboulsi, Pascale Hoffmann, Jean-Jacques Feige, Mohamed Benharouga, Jean-Luc Coll, and Nadia Alfaidy

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12

Real-Time Assessment of Mitochondrial Function in Cytotrophoblast and Syncytialized Trophoblast Cells Using the Seahorse XFe24 Extracellular Flux Analyzer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137 O’Llenecia S. Walker, Linda L. May, and Sandeep Raha 13 Quantifying Trophoblast Function Using the xCELLigence System. . . . . . . . . . . 149 Rosemary J. Keogh and Padma Murthi

PART III

TARGETED THERAPY

14

Liposome-Encapsulated Anti-inflammatory Proteins for Targeted Delivery to the Placenta to Treat Fetal Growth Restriction. . . . . . . . . . . . . . . . . . . 165 Padma Murthi and Lynda K. Harris 15 Trophoblast-Targeted Liposomes for Placenta-Specific Drug Delivery . . . . . . . . . 173 Baozhen Zhang, Xiujun Fan, and Nihar R. Nayak

PART IV ADVANCED 3D MODELS 16

Methods for Co-culture of Primary Human Extravillous Trophoblast Cells and Uterine Natural Killer Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xiaowen Gan, Fen Ning, and Gendie E. Lash 17 Trophoblast Organoids as a Novel Tool to Study Human Placental Development and Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sandra Haider, Martin Kno¨fler, and Paulina A. Latos 18 A Three-Dimensional Trophoblast Invasion Microfluidic Platform for Toxicological Screening . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yong Pu and Almudena Veiga-Lopez 19 Three-Dimensional In Vitro Human Placental Organoids from Mononuclear Villous Trophoblasts or Trophoblast Stem Cells to Understand Trophoblast Dysfunction in Fetal Growth Restriction . . . . . . . . . . . . Cherry Sun, Joanna L. James, and Padma Murthi Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors ROLAND ABI NAHED • Institut National de la Sante´ et de la Recherche Me´dicale, Inserm U1292, Grenoble, France; University Grenoble-Alpes, Grenoble, France; Commissariat `a l’Energie Atomique et aux Energies Alternatives (CEA), Biosciences and Biotechnology Institute of Grenoble, Grenoble, France RUPASRI AIN • Division of Cell Biology and Physiology, CSIR-Indian Institute of Chemical Biology, Kolkata, India CHRISTIANE ALBRECHT • Institute of Biochemistry and Molecular Medicine, Faculty of Medicine, University of Bern, Bern, Switzerland; Swiss National Centre of Competence in Research (NCCR) TransCure, University of Bern, Bern, Switzerland NADIA ALFAIDY • Institut National de la Sante´ et de la Recherche Me´dicale, Inserm U1292, Biosante´, Grenoble, France; University Grenoble-Alpes, Grenoble, France; Commissariat `a l’Energie Atomique et aux Energies Alternatives (CEA), Biosciences and Biotechnology Institute of Grenoble, Grenoble, France; Centre Hospitalo-Universitaire Grenoble Alpes, Service Obste´trique, CS 10217, Grenoble Cedex 9, France BENOIT BARBEAU • De´partement des sciences biologiques, Centre d’excellence en recherche sur les maladies orphelines-Fondation Courtois, Universite´ du Que´bec `a Montre´al, Montreal, QC, Canada; Re´seau intersectoriel de recherche en sante´ de l’Universite´ du Que´bec, Montreal, QC, Canada MOHAMED BENHAROUGA • Institut National de la Sante´ et de la Recherche Me´dicale, Inserm U1292, Biosante´, Grenoble, France; University Grenoble-Alpes, Grenoble, France; Commissariat `a l’Energie Atomique et aux Energies Alternatives (CEA), Biosciences and Biotechnology Institute of Grenoble, Grenoble, France CHERIE BLENKIRON • Hub for Extracellular Vesicles Investigations (HEVI), University of Auckland, Auckland, New Zealand; Auckland Cancer Society Research Centre, School of Medical Sciences, University of Auckland, Auckland, New Zealand; Department of Molecular Medicine and Pathology, School of Medical Sciences, University of Auckland, Auckland, New Zealand LAWRENCE W. CHAMLEY • Department of Obstetrics and Gynaecology, School of Medicine, University of Auckland, Auckland, New Zealand; Hub for Extracellular Vesicles Investigations (HEVI), University of Auckland, Auckland, New Zealand JEAN-LUC COLL • Institute for Advanced Biosciences, INSERM-UGA U1209, CNRS UMR 5309, La Tronche, France CONSTANCE COLLET • Institut National de la Sante´ et de la Recherche Me´dicale, Inserm U1292, Biosante´, Grenoble, France; University Grenoble-Alpes, Grenoble, France; Commissariat `a l’Energie Atomique et aux Energies Alternatives (CEA), Biosciences and Biotechnology Institute of Grenoble, Grenoble, France SUZANNE DE LA MONTE • Regional Perinatal Pathology, Kaiser Permanente Oakland Medical Center, Oakland, CA, USA; Department of Pathology and Laboratory Medicine, The Warren Alpert Medical School of Brown University, Providence, RI, USA; Department of Pathology and Laboratory Medicine, Rhode Island Hospital, Providence, RI, USA TOSHIHKO EZASHI • Colorado Centre for Reproductive Medicine, Lone Tree, CO, USA XIUJUN FAN • Laboratory of Reproductive Health, Shenzhen Institute of Advanced Technology, Chinese Academy of Sciences, Shenzhen, China

ix

x

Contributors

JEAN-JACQUES FEIGE • Institut National de la Sante´ et de la Recherche Me´dicale, Inserm U1292, Biosante´, Grenoble, France; University Grenoble-Alpes, Grenoble, France; Centre Hospitalo-Universitaire Grenoble Alpes, Service Obste´trique, CS 10217, Grenoble Cedex 9, France BARBARA FUENZALIDA • Institute of Biochemistry and Molecular Medicine, Faculty of Medicine, University of Bern, Bern, Switzerland XIAOWEN GAN • Division of Uterine Vascular Biology, Guangzhou Institute of Pediatrics, Guangzhou Women and Children’s Medical Center, Guangzhou Medical University, Guangzhou, China JEFFREY GILLIGAN • Department of Neurosurgery, Mount Sinai Health System, New York, NY, USA KATIE M. GROOM • Liggins Institute, University of Auckland, Auckland, New Zealand FUSUN GUNDOGAN • Regional Perinatal Pathology, Kaiser Permanente Oakland Medical Center, Oakland, CA, USA; Department of Pathology and Laboratory Medicine, The Warren Alpert Medical School of Brown University, Providence, RI, USA SANDRA HAIDER • Department of Obstetrics and Gynecology, Reproductive Biology Unit, Placental Development Group, Medical University of Vienna, Vienna, Austria LYNDA K. HARRIS • Maternal and Fetal Health Research Centre, Division of Developmental Biology and Medicine, Faculty of Biology, Medicine and Health, University of Manchester, St Mary’s Hospital, Manchester, UK; St Mary’s Hospital, Manchester University NHS Foundation Trust, Manchester Academic Health Science Centre, Manchester, UK; Division of Pharmacy and Optometry, Faculty of Biology, Medicine and Health, University of Manchester, Manchester, UK PASCALE HOFFMANN • Institut National de la Sante´ et de la Recherche Me´dicale, Inserm U1292, Biosante´, Grenoble, France; University Grenoble-Alpes, Grenoble, France; Commissariat `a l’Energie Atomique et aux Energies Alternatives (CEA), Biosciences and Biotechnology Institute of Grenoble, Grenoble, France; Centre Hospitalo-Universitaire Grenoble Alpes, Service Obste´trique, CS 10217, Grenoble Cedex 9, France JOANNA L. JAMES • Department of Obstetrics and Gynaecology, Faculty of Medical and Health Sciences, The University of Auckland, Auckland, New Zealand ADAM JAREMEK • Department of Anatomy and Cell Biology, Schulich School of Medicine and Dentistry, University of Western Ontario, London, ON, Canada VERONIQUE JOSSERAND • Institute for Advanced Biosciences, INSERM-UGA U1209, CNRS UMR 5309, La Tronche, France ROSEMARY J. KEOGH • Department of Obstetrics & Gynaecology, Royal Women’s Hospital, University of Melbourne, Parkville, VIC, Australia MICHELLE KERAMIDAS • Institute for Advanced Biosciences, INSERM-UGA U1209, CNRS UMR 5309, La Tronche, France MARTIN KNO¨FLER • Department of Obstetrics and Gynecology, Reproductive Biology Unit, Placental Development Group, Medical University of Vienna, Vienna, Austria GENDIE E. LASH • Division of Uterine Vascular Biology, Guangzhou Institute of Pediatrics, Guangzhou Women and Children’s Medical Center, Guangzhou Medical University, Guangzhou, China PAULINA A. LATOS • Center for Anatomy and Cell Biology, Medical University of Vienna, Vienna, Austria JONATHAN LAVAUD • Institute for Advanced Biosciences, INSERM-UGA U1209, CNRS UMR 5309, La Tronche, France DEIRDRE M. LOGSDON • Colorado Centre for Reproductive Medicine, Lone Tree, CO, USA

Contributors

xi

LINDA L. MAY • Department of Pediatrics, McMaster University, Hamilton, ON, Canada PADMA MURTHI • Department of Pharmacology, Monash Biomedicine Discovery Institute, Monash University, Clayton, VIC, Australia; Department of Obstetrics and Gynaecology, University of Melbourne, Melbourne, VIC, Australia; Department of Maternal-Fetal Medicine, Pregnancy Research Centre, Royal Women’s Hospital, Parkville, VIC, Australia NIHAR R. NAYAK • Department of Obstetrics and Gynecology, UMKC School of Medicine, Kansas City, MO, USA FEN NING • Division of Uterine Vascular Biology, Guangzhou Institute of Pediatrics, Guangzhou Women and Children’s Medical Center, Guangzhou Medical University, Guangzhou, China YOHANES N. S. NURSALIM • Department of Obstetrics and Gynaecology, School of Medicine, University of Auckland, Auckland, New Zealand; Hub for Extracellular Vesicles Investigations (HEVI), University of Auckland, Auckland, New Zealand MADHURIMA PAUL • Division of Cell Biology and Physiology, CSIR-Indian Institute of Chemical Biology, Kolkata, India YONG PU • Department of Pathology, University of Illinois at Chicago, Chicago, IL, USA SANDEEP RAHA • Graduate Programme in Medical Sciences, McMaster University, Hamilton, ON, Canada; Department of Pediatrics, McMaster University Medical Centre, Hamilton, ON, Canada STEPHEN J. RENAUD • Department of Anatomy and Cell Biology, Schulich School of Medicine and Dentistry, University of Western Ontario, London, ON, Canada; Children’s Health Research Institute, London, ON, Canada; Lawson Health Research Institute, London, ON, Canada DEBORAH REYNAUD • Institut National de la Sante´ et de la Recherche Me´dicale, Inserm U1292, Grenoble, France; University Grenoble-Alpes, Grenoble, France; Commissariat `a l’Energie Atomique et aux Energies Alternatives (CEA), Biosciences and Biotechnology Institute of Grenoble, Grenoble, France FRE´DE´RIC SERGENT • Institut National de la Sante´ et de la Recherche Me´dicale, Inserm U1292, Grenoble, France; University Grenoble-Alpes, Grenoble, France; Commissariat `a l’Energie Atomique et aux Energies Alternatives (CEA), Biosciences and Biotechnology Institute of Grenoble, Grenoble, France CHERRY SUN • Department of Obstetrics and Gynaecology, Faculty of Medical and Health Sciences, The University of Auckland, Auckland, New Zealand CAROLINE TOUDIC • De´partement des sciences biologiques, Centre d’excellence en recherche sur les maladies orphelines-Fondation Courtois, Universite´ du Que´bec `a Montre´al, Montreal, QC, Canada WAEL TRABOULSI • Laboratory for Immuno-Oncology, Lombardi Comprehensive Cancer Center, Georgetown University Medical Center, Washington, DC, USA ALMUDENA VEIGA-LOPEZ • Department of Pathology, University of Illinois at Chicago, Chicago, IL, USA; The Chicago Center for Health and Environment, University of Illinois at Chicago, Chicago, IL, USA O’LLENECIA S. WALKER • Graduate Programme in Medical Sciences, McMaster University, Hamilton, ON, Canada HUANGHE YANG • Department of Biochemistry, Duke University Medical Center, Durham, NC, USA; Department of Neurobiology, Duke University Medical Center, Durham, NC, USA YE YUAN • Colorado Centre for Reproductive Medicine, Lone Tree, CO, USA

xii

Contributors

JONAS ZAUGG • Institute of Biochemistry and Molecular Medicine, Faculty of Medicine, University of Bern, Bern, Switzerland; Swiss National Centre of Competence in Research (NCCR) TransCure, University of Bern, Bern, Switzerland; Department of Physiology, Development and Neuroscience, University of Cambridge, Cambridge, UK BAOZHEN ZHANG • Department of Obstetrics and Gynecology, Women and Children’s Hospital of Chongqing Medical University, Chongqing, China YANG ZHANG • Department of Biochemistry, Duke University Medical Center, Durham, NC, USA

Part I Isolation and Culture Methods

Chapter 1 Isolation and Maintenance in Culture of Primary Human Trophoblast from Term Placentae Yohanes N. S. Nursalim, Katie M. Groom, Cherie Blenkiron, and Lawrence W. Chamley Abstract Trophoblasts are placenta-specific epithelial cells that play an essential role in conducting nutrient, gas, and waste exchange between the fetus and the mother. Primary culture of human trophoblasts from donated term placentae is an important tool to study placental functions. Currently, there is a lack of general consensus of the optimal culture conditions for maintaining term trophoblast cells in vitro. A key problem with culturing trophoblasts from term placentae is overgrowth of the trophoblasts by rapidly proliferating cellular contaminants. Recently we reported a system to culture trophoblasts from term placentae which differentiate into syncytiotrophoblast-like multinucleated cells that can be maintained in culture for at least 30 days with minimal contamination. This chapter details our optimized approach for long-term, contaminant-free in vitro culture of primary trophoblasts from term placentae. Key words Trophoblast, Syncytiotrophoblast, Cytotrophoblast, Primary culture, Long-term culture, Human term placenta

1

Introduction Trophoblasts are an epithelial cell-type specific to the placenta that play an essential role in placental function. Abnormalities in trophoblast function have direct consequences leading to pathologies associated with pregnancy. Culture of human primary trophoblasts provides an important model for placental research as animal models have limited utility due to the large differences between the anatomical structure of human and other placentae [1, 2]. In humans, the syncytiotrophoblast is a major trophoblast subtype that is in direct contact with the maternal blood, thus acting as a fetal-maternal interface that facilitates nutrient and oxygen transfer [3, 4]. Creating an in vitro syncytiotrophoblast model provides a valuable tool for the study of placental function but can be challenging. The syncytiotrophoblast is a large single cell arising from

Sandeep Raha (ed.), Trophoblasts: Methods and Protocols, Methods in Molecular Biology, vol. 2728, https://doi.org/10.1007/978-1-0716-3495-0_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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the fusion of progenitor mononuclear cytotrophoblasts and can grow to an area of 11–13 m2 [5]. Because of its size, it is difficult to directly isolate intact syncytiotrophoblast, and fragmented syncytiotrophoblast does not survive in prolonged culture. A common alternative to circumvent this problem is to culture the progenitor cytotrophoblasts, which have been shown to spontaneously fuse into syncytiotrophoblast-like multinucleated clusters when grown in vitro [6–11]. The method to maintain a naturally fused trophoblast culture is a fundamental principle in primary culture, employed as early as 1960 [6], with varying degrees of success and failure. A major cause of the failure of this technique is that freshly isolated cytotrophoblasts lose their capacity to proliferate in vitro [10]. Other non-trophoblastic contaminants, such as fibroblasts, maintain their ability to proliferate and frequently overtake the culture within 7–10 days [10, 11]. Currently, there are methods to reduce these contaminants from the trophoblast isolates using Percoll gradient separation or negative selection using antibody-coated magnetic beads [7, 8, 12], but cultures applying these additional steps have not extended beyond approximately 7 days in culture. In addition, the protocols described for successful primary trophoblast cultures lack consistency between different research groups [13]. Recently, a novel system to culture human trophoblasts from donated term placentae was established [9]. This method allows (1) the production of multinucleated syncytialized clusters from freshly isolated cytotrophoblast and (2) maintenance of the cultures in vitro for at least 30 days without the culture being overgrown by contaminating cells [9]. Since this system allows prolonged survival of trophoblasts in culture, the technique provides a new and useful platform for long-term in vitro study of the human term placenta.

2

Materials

2.1 Equipment and Materials

1. 15 mL centrifuge tube. 2. 50 mL centrifuge tube. 3. 50 mL sterile syringe. 4. 70 mL gamma sterile screw cap container (Thermo Fisher Scientific, cat#LBS30005X). 5. Cell Strainers 70 μm/White for use w/50 mL conical tubes, sterile (In Vitro Technologies, cat# FAL352350). 6. Class II laminar flow hood. 7. CO2 incubator. 8. Microcentrifuge. 9. Incontinence sheet (absorbent pads).

Primary Trophoblast Culture from Term Placentae

5

10. Neubauer hemocytometer. 11. Brightfield Microscope, inverted. 12. Brightfield Microscope, upright. 13. Controlled temperature water bath. 14. Sterile 6- or 24-well cell culture plates. 15. Sterile cheesecloth. 16. Sterile forceps. 17. Sterile Petri dish. 18. Sterile scalpels. 19. Sterile Schott bottle. 2.2 Chemical Reagents and Media

All chemicals should be of Laboratory or Analytical Grade. Unless stated otherwise, the water (distilled H2O) should be of ultrapure grade and autoclaved prior to use. 1. Advanced DMEM/F-12 cat#12634010).

(Thermo

Fisher

Scientific,

2. Corning® Matrigel® Basement Membrane Matrix, *LDEVFree (In Vitro Technologies, cat#354234). 3. Digestion buffer (1 mg/mL dispase; Roche, cat#4942078001 and 0.5 mg/mL DNase I; Sigma, cat# DN25, dissolved with 1× sterile PBS). 4. Dispase II dissolved with 1× PBS at 1 mg/mL, filtered sterile. 5. DNase I dissolved in 1× PBS at 0.5 mg/mL filtered sterile. 6. Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F12) with GlutaMAX (Thermo Fisher Scientific, cat#10565042). 7. Fetal bovine serum (FBS; New Zealand origin). 8. N-2-hydroxyethylpiperazine-N-2-ethane (HEPES) buffer.

sulfonic

acid

9. Newborn Calf Serum (NBCS; New Zealand origin). 10. Penicillin-Streptomycin (P/S). 11. Phosphate-buffered saline (PBS). In 1 L add 8.09 g NaCl, 0.2 g KCl, 0.2 g KH2PO4, 1.14 g Na2HPO4. Autoclave to sterilize. 12. Recombinant Human Epidermal Growth Factor (hEGF) Protein (In Vitro Technologies, cat#RDS236EG01M) reconstituted with sterile dH2O to a concentration of 0.5 mg/mL. 13. Plating medium (DMEM/F12, with 10% FBS, 5 ng/mL hEGF, 1% P/S, 15 mM HEPES). 14. Maintenance medium (Advanced DMEM/F12, with 2% FBS, 5 ng/mL hEGF, 1% P/S).

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15. Erythrocyte lysis buffer (0.802 g NH4Cl, 0.084 g NaHCO3, and 0.037 g EDTA, dissolved in 100 mL dH2O, filter using 0.2 μm filter to sterilize). 16. Trypan blue solution.

3

Methodology Human term placentae should be obtained from donors with written informed consent, undergoing normal, uncomplicated vaginal delivery or elective caesarean delivery. All samples should be processed within 2 h of birth. All procedures should be performed in sterile conditions inside a Class II laminar flow hood.

3.1 Isolation Protocol

1. Place the placenta on a paper incontinence sheet with the maternal side up, and remove the fetal membranes to expose the placental lobes. Choose a lobe of the placenta free from obvious damage or pathology, such as calcification (Fig. 1a). 2. Make a shallow (approximately 2–5 mm) rectangular (approximately 2 cm by 3 cm) shaped incision on the lobe (Fig. 1b). 3. Lift one edge of the rectangle with sterile forceps, and then dissect the rectangle parallel to the maternal surface (approximately 2–3 mm below the maternal surface), effectively peeling off the maternal interface, and discard this interface which is contaminated with maternal decidua and bacteria. The underlying villi will now be exposed and are sterile (Fig. 1c; see Note 1). 4. Using fresh sterile dissecting equipment, excise the exposed, underlying placental villi, taking care not to cut through the non-sterile amniotic surface of the placenta, and place the villi into a sterile, 70 mL gamma sterile screw cap container with sterile PBS solution (see Note 1). 5. Wash the villous tissue in PBS, replacing the PBS as needed, until maternal blood is no longer evident, and then transfer into a Petri dish containing a small amount of PBS (approximately 3–4 mL; Fig. 2a). 6. Remove vessels and connective tissues by gentle scraping from the villi using a scalpel (see Note 2, Fig. 2b). 7. Collect the processed tissues into a sterile container, at a weight needed for downstream use (we typically find 40 g of villous tissue yields approximately 1 × 107 mononuclear trophoblasts). Then transfer the tissues into a sterile Schott bottle or 50 mL centrifuge tubes (Fig. 3a), making sure to avoid adding any excess PBS (see Note 3).

Primary Trophoblast Culture from Term Placentae

7

Fig. 1 The dissection process used to isolate placental villi. Place the placenta on incontinence sheet with the maternal aspect facing upwards (a). Make a rectangular incision on a lobe of the placenta that is free from obvious damage or pathology (b, yellow box). The outer, non-sterile, maternal surface is peeled off (white arrow) and removed by making an incision parallel to the decidua and discarded (c)

8. Digest the tissue twice, using digestion buffer at a ratio of 3 mL per g of tissue for both digestion steps. For the first digest, add the digestion buffer into the container, and incubate in 37 °C water bath for 10 min with gentle agitation every 2 min. The solution should be cloudy indicating successful digestion of the tissue (see Fig. 3a). Discard this first digestion buffer, and wash the sedimented villous fragments with PBS until the solution is clear (see Note 3). 9. For the second digest, add the digestion buffer, and incubate the villous fragments in digestion buffer for 16 h at 4 °C without stirring.

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Fig. 2 Removal of connective tissue and blood vessels from villous tissue. (a) The villous tissues are kept in PBS to prevent them from drying out during processing. (b) The blood vessels (black arrowhead) and connective tissue are removed by manual dissection using a scalpel by scraping around the villi (white arrowhead)

Fig. 3 (a) Villous tissue fragments after the first digest, performed in a 50 mL centrifuge tube (suitable for 20 passages. 4. Complete growth medium: DMEM-low glucose, GlutaMAX™ Supplement, 1.0 mM pyruvate, containing 10–15% fetal bovine serum (FBS) and 1X Antibiotic-Antimycotic (100X contains: 10,000 units/mL penicillin, 10,000 μg/mL streptomycin and 25 μg/mL Amphotericin B). 5. Freezing medium: 10% DMSO/90% FBS. 6. 0.25% Trypsin-EDTA solution, suitable for cell culture. 7. Dulbecco’s Phosphate Buffered Saline (DBPS), without calcium chloride and magnesium chloride, liquid, sterile-filtered, suitable for cell culture.

2.2

Leucine Uptake

1. White-walled 96-well plates with clear bottom (see Note 1). 2. Na+-free Hank’s buffer: 5X stock solution (10X is not stable at 4 °C) in distilled water containing 125 mM choline chloride, 25 mM HEPES, 4.8 mM KCl, 1.2 mM MgSO4, 1.2 mM KH2PO4, 1.3 mM CaCl2, 5.6 mM glucose, adjust the pH to 7.4 with 5 M Tris-HCl (no Na+), fill up to final volume and sterile-filter to prolong storage time. The Hank’s 5X stock solution can be stored at 4 °C. Dilute the sterile-filtered stock solution shortly before starting the experiment.

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3. Leucine uptake solution: 150 μM L-Leucine uptake solution spiked with 2 μCi/mL L-[3,4,5-3H(N)]-leucine in Hank’s buffer. Use a 100 mM L-leucine stock solution prepared in Hank’s buffer and store at -20 °C. 4. Forskolin stock solution: 100 mM Forskolin in 95% ethanol = 1000x stock solution (store at -20 °C). 5. BCH (Inhibitor): Dissolve 0.186 g 2-Amino-2-norbornanecarboxylic acid (BCH) in 10 mL Hank’s buffer. 6. Pierce BCA Protein Assay Kit. 7. Scintillation cocktail (MicroScint-20). 8. Orbital shaker to mix 96-well plates. 9. TopCount® NXT™ Scintillation and Luminescence Counter. 2.3

Leucine Transfer

The following components are needed additionally to the material listed in Subheading 2.2: 1. 12-well plates (see Note 2). 2. Black-walled 96-well plates with clear bottom (see Note 1). 3. Transwell® inserts for 12-well plates (Note 2). 4. DMEM, low glucose, pyruvate, no glutamine, no phenol red. 5. Leucine transfer solution: Prepare a 150 μM L-Leucine uptake solution spiked with 2 μCi/mL L-[3,4,5-3H(N)]-leucine and 300 μM L-Glutamine (Gln) in Hank’s buffer. Use a 100 mM L-leucine and 100 mM L- glutamine stock solution prepared in Hank’s buffer and store in -20 °C. 6. 25 mM Lucifer yellow stock solution: Dissolve 25 mg Lucifer yellow (LY), 457 Da in 2.19 mL DPBS and keep at 4 °C in the dark. 7. Flex Station II fluorescence microplate reader (Molecular Devices, USA). 8. Tri-carb 2100TR Liquid Scintillation Counter (PerkinElmer, Waltham, US). 9. Millicell ERS-2 Volt-Ohm Meter (Millipore, MA, USA) (see Note 3). 10. cellZscope system with electrode set “12-well” type (nanoAnalytics, Mu¨nster, Germany).

3

Methods

3.1 Preparation of BeWo Cells

1. Remove the cryo-preserved BeWo cell tube from the liquid N2 store (see Note 4). 2. Add multiple times complete growth medium until all cells are thawed and transfer them to the 15 mL-centrifuge tube. Dilute cytotoxic DMSO in cryo-culture as fast as possible.

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3. Centrifugation for 5 min at 500 rcf. 4. Wash the cells twice with complete growth medium (see Note 5). 5. Resuspend the cell pellet in ca. 6 mL medium. 6. Spin cells down in centrifuge for 5 min with 500 rcf. 7. Resuspend the cell pellet in ca. 1 mL complete growth medium and transfer all cells into a T75-flask with 9 mL prewarmed complete growth medium. 8. Disperse the cells homogenously over the bottom of the flask. Smoothly move the flask in x-direction. Let the flask stay to calm the waves. Repeat this in y-direction. Observe the cells periodically under inverted microscope. 9. Incubate the cells for more than 2 days at 37 °C and 5% pCO2. Observe periodically (at least daily) to monitor the growth. 10. Change medium every 2–3 days at least. 3.2

Leucine Uptake

3.2.1 Preparation for Leucine Uptake DoseResponse Assay

1. Preparation of the uptake solution spiked with radioactive L-leucine in pre-warmed Hank’s buffer (150 μM cold Leu + 2 μCi/mL 3H-Leu in Na+-free Hank’s buffer). Example for leucine uptake solution pipetting for ten upper or six lower compartments: 15 μL

cold L-leucine (100 mM stock)

15 μL

cold L-glutamine (100 mM stock)

20 μL

radioactive L-3H-leucine (1 mCi/mL stock)

9950 μL

Hank’s buffer

2. Design the plate (see example for BCH dose-response assay in Table 2). 3. Wash BeWo cells cultured in T-75 flasks twice with 6 mL DPBS to remove FBS containing complete growth medium (FBS inactivates trypsin). 4. Add 3 mL 0.25% Trypsin-EDTA solution and distribute until all cells are covered. 5. Incubate the flask in 37 °C incubator and periodically check the cells for detachment (at least 2 min). Carefully tap the flask with your hand at the side. You can easily see the cells detaching by eye or under the inverted microscope. 6. As soon all cells are detached quickly add 7 mL of complete growth medium to inactivate the trypsin and count cell concentration using a coulter counter.

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Table 2 Pipetting scheme for leucine uptake dose-response assay in a 96-well plate 1

2

3

4

5

6

7

8

9

10

11

12

A

DC

B

DC

C

DC

D

DC

E

DC

Prt

Prt

Prt

Prt

Prt

Prt

Prt

NCB

NCB

NCB

NCB

NCB

NCB

NCB

NCB

NCB

0.0316

0.01 µM

0.001 µM

0.0001

0

6 min

7 min

8 min

9 min

0.316 µM

Prt

1 µM

NCB NCB

Prt

10 µM

Prt

100 µM

H

1000 µM

Prt

BCH conc.

G

0.1 µM

F

5 min

4 min

3 min

2 min

1 min

0.5 mi

0 min

Time

geometrical dilution series

Time course Legend: The main steps in the geometrical dilution series are suggested as 1:10 steps and half-logarithmic steps around the expected IC50. For time course experiments an observation time of max. 10 min with steps of 60 s are recommended. Shades of grey indicate inhibitor concentration; Prt wells for protein measurement after the uptake experiment, NCB no cell background control well (no cells seeded), DC dosage control wells (measurement of a tenth of the total dose)

3.2.2

Leucine Uptake

The previously described leucine uptake method was adapted to test the effect of different substrate concentrations on leucine uptake kinetics [20, 22]. Hank’s buffer is referring to Na+-free Hank’s buffer in the entire protocol. 1. Seed the cells 24 h before starting the experiment (T-24 h) into clear bottom/white wall 96-well plates at a density of 60,000 cells/100 μL/well (600,000 cells/mL) except in NCB (no cell background) control wells (see the example for a BCH doseresponse assay in Table 2). (a) To investigate differentiated BeWo cells in their syncytiotrophoblast stage (BeWo-STB) seed the BeWo cell with the same density as mentioned before and stimulate cells

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with 100 μM Forskolin (use a 1000X stock) in complete growth medium for at least 48 h (start experiment at T-48 h). (b) One BeWo T-75 flask with an 80–90% confluency contains approximately 5–6 × 106 cells. 2. Wash twice with 120 μL Hank’s buffer and starve the cells with 100 μL/well Hank’s buffer for 30 min (start at T-0.5h) at 37 °C with 5% pCO2. 3. Continue with step 5, if no dose-response assay is applied (see Note 6). Otherwise, remove the Hank’s buffer using suction and add 100 μL of pre-diluted candidate inhibitor in Hank’s buffer (see Table 2). 4. Incubate cells and NCB with uptake solution in absence or presence of inhibitors at 37 °C with 5% pCO2 for 3 min (start at T0). 5. Uptake-Stop: Wash 3 times with cold Hank’s buffer and don’t add buffer after the last washing step. For time courses: Stop the leucine uptake after predefined time points. An observation time of 10 min, maximum, is recommended with steps of 60 s, maximum (see Table 2). 6. Add 10 μL of the corresponding uptake solution (cold leucine/ radioactive 3H-leucine mix) to Dosage Control (DC) wells (see Table 2). 7. Add 100 μL MicroScint-20 solution to all sample wells, 90 μL MicroScint-20 solution into DC wells, 55 μL radioimmunoassay precipitation (RIPA) buffer into wells for protein measurement wells for total protein determination using BSA Pierce protein measurement kit and cover the plate with plate sealer (see Table 2). 8. Lyse the cells in MicroScint™-20 (PerkinElmer, Waltham, MA, USA) by shaking the plate at 250 rpm for >1.5 h. 9. Measure scintillation with scintillation counter (3H program; 20 min/plate) using a TopCount Scintillation Counter. 10. Transfer of the counts into a spreadsheet for calculation of leucine uptake. (a) Leucine uptake is calculated per well using the following equations: Scintillation count ½cpm - mean of NCB ½cpm mean of DC × 10 ½cpm = Leu uptake ½% of total dosage

ð1Þ

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Leu uptake ½% ×

L - leucine conc: in uptake solution ½nmol=mL Total protein conc: determined by BSA Pierce ½mg=mL

= Leu uptake ½nmol=mg protein ð2Þ 3.3 Leucine Transfer Across the Placental Barrier Using Transwell® Technique

The leucine transfer method is based on previously published experiments using primary trophoblasts [23]. Hank’s buffer is referring to Na+-free Hank’s buffer in the entire protocol. To stimulate the transition of undifferentiated BeWo cells into the syncytiotrophoblast stage (BeWo-STB), the BeWo (clone b30) cells were treated with 100 μM Forskolin for at least 48 h (start experiment at T-48 h).

3.3.1 Preparation for Leucine Transport Across a Placental Barrier Model

1. Seed BeWo cells at the density of 140,000 cells/cm2 (160,000 BeWo cells/12-well insert) in 800 μL of pre-warmed complete growth medium onto the insert (upper compartment). (a) NEVER touch the Transwell® membrane! (b) Try to plate the cells gently and drop by drop uniformly over the entire area of the membrane. (c) Include wells without seeding cells as no cell background control (NCB). 2. Add 1.5 mL pre-warmed complete growth medium into the well (lower compartment). 3. Let the cells settle down on the membrane by leaving the seeded plates covered in sterile laminar flow hood for 15 min without any disturbance. 4. Gently move the plate to the incubator and incubate at 37 °C with 5% pCO2. (a) Start the trophoblast differentiation by exchanging the medium in both compartments with pre-warmed complete growth medium including 100 μM Forskolin (use a 1000X stock) for at least 48 h (start experiment at T-48 h). Include no-Forskolin control wells at least in the pilot experiments. (b) Monitor the tightness for 5–8 days until both TEER and the Papp reach a plateau. (c) The approximate plateau levels of TEER and the Papp should be evaluated in a pilot experiment (see Note 7).

3.3.2 Trans-epithelial Electrical Resistance (TEER)

The barrier-forming capacity of the trophoblasts was evaluated by measuring TEER of the cell layer using a Millicell ERS-2 Volt-Ohm Meter each day for 2 h [23] or using the cellZscope system

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(nanoAnalytics, Mu¨nster, Germany) according to the manufacturer’s instructions [21]. Millicell ERS-2 Volt-Ohm Meter 1. Exchange of the medium with fresh pre-warmed complete growth medium. Avoid the formation of foam: even little bubbles generate big variance in resistance measurements (see Note 8). 2. Place the electrodes on both sides of the insert (see Note 9). 3. Keep electrode arms vertical and measure at three different insert positions (approximately 120 degree turns around the insert). 4. Resistance is analyzed over time as means of the three positions and corrected for the surface area to obtain Ω* cm2. cellZscope System The formation of a tight trophoblast monolayer by BeWo cells can be monitored by measuring transepithelial electrical resistance (TEER, [Ω*cm2]) and cellular capacitance (Ccl, [μF/cm2]) according to the manufacturer’s instructions. 3.3.3 Apparent Permeability Coefficient ( Papp)

Lucifer yellow (LY), a low molecular weight substance (size 444 g·mol-1), is used to determine the passive paracellular transfer across the cell monolayer. 1. Equilibrate cells in pre-warmed phenol red-free DMEM medium for 30 min. 2. Dilute the 25 mM LY stock to a 50 μM working solution (1: 500). (a) For example: 30 μL LY + 15 mL phenol red-free DMEM medium. 3. Add LY working solution into the donor compartment, basal (1.5 mL) or apical (800 μL) and add phenol red-free DMEM medium into the receiver compartments with the respective volume; incubate 30 min at 37 °C with 5% pCO2. 4. Prepare standard curve with LY in phenol red-free DMEM medium with a range from 50 μM to 0.05 μM (see Table 3). 5. Take 120 μL aliquots in 30 min intervals only from the receiver chamber for 2 h. Always remove and discard the same volume of 120 μL from the corresponding (not analyzed) compartment to equilibrate hydrostatic pressure differences, while keeping the samples in the dark. 6. Spin down any debris at 1000 rcf for 5 min and transfer 100 μL of each standard and sample collected at the different time

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Table 3 Template dilution series of Lucifer Yellow (LY) in phenol red-free DMEM medium in a 96-well plate Well

LY conc. [μM]

Transfer volume [μL]

Diluent volume [μL]

Total volume [μL]

1

50

1000

2

25

500

500

500

3

12.5

500

500

500

4

6.25

500

500

500

5

3.125

500

500

500

6

1.5625

500

500

500

7

0.7813

500

500

500

8

0.3906

500

500

500

9

0.1953

500

500

500

10

0.09766

500

500

500

11

0.04883

500

500

1000

12

0

500

500

500

intervals into a 96-well plate (preferentially use black-walled 96-well plates with clear bottom for fluorescent detection). 7. After finishing the LY diffusion experiment, immediately replace the media with pre-warmed complete growth medium. 8. Read fluorescence at Ex/Em = 450/530 nm by a Flex Station II fluorescence microplate reader (Molecular Devices, USA). Lucifer yellow has excitation/emission peaks of 428/536 nm. 9. Papp is calculated per well using the following equation and compared to the NCB wells. Apparent Permeability Coefficient (Papp) Calculation P app ½cm=s = ðdQ=dt Þ ÷ ðA × C 0 Þ

ð3Þ

dQ/dt: rate of the substrate appearance in the receiver chamber (μmol/s); A: surface area of the filter membrane (cm2); C0: initial concentration of the substrate in the donor chamber (μmol/mL). 3.3.4

Leucine Transfer

Before starting the transfer experiment, the formation of a tight trophoblast monolayer in BeWo cells has to be assayed. By measuring both the TEER and Papp values as reported previously [20] and as shown in the supporting information (Suppl. Fig. 1 online on Zenodo, https://doi.org/10.5281/zenodo.4384654), the achievement of the plateau phase can be monitored.

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Fig. 1 Assessment of placental nutrient transport. Forskolin stimulation of the BeWo clone b30 trophoblast cell line allows (i) to assess leucine uptake, if cultured in conventional well plates and (ii) to measure leucine transfer, if using the Transwell® system. Characterization of the SLC7-mediated leucine transport is performed under sodium-free conditions

1. Select Transwell® inserts with verified tightness and randomly assign them for Leu transfer time course experiments. (a) Include NCB Transwell® inserts as controls. (b) Use at least triplicates for each experimental condition. 2. Preparation of the transfer solution spiked with radioactive L-leucine and glutamine (L-Gln) as exchange substrate in pre-warmed Hank’s buffer (150 μM cold Leu, 300 μM Gln + 2 μCi/mL 3H-Leu in Na+-free Hank’s buffer). See example in Note 10. 3. Wash all Transwell® inserts 3-times with Hank’s buffer (900 μL in upper- and 1.7 mL in lower compartment) and starve the cells with 800 μL in upper- and 1.5 mL Na+-free Hank’s buffer (pH 7.4) in lower compartment for 30 min (start at T-0.5 h) at 37 °C with 5% pCO2. 4. Start the Leu transfer from the upper (maternal, 800 μL) toward the lower (fetal, 1500 μL) compartment or vice versa by simultaneously replacing the Hank’s buffer in donor compartments with transfer solution (T0). 5. Take 50 μL samples from 0–6 h simultaneously from the donating and receiving compartment. The dose control (DC) is a 50 μL sample taken directly from the spiked transfer solution (see Note 11). 6. At the end of the transfer experiment, wash all membranes of the Transwell® inserts twice with DPBS and cut them out. 7. Collect all the medium and membrane samples including non-cell controls (NCB) in 3 mL of scintillation cocktail.

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8. Lyse the cells on the membrane samples and homogenize all scintillation samples by shaking the plate with 250 rpm for >1.5 h. 9. Measure scintillation using a Tri-carb 2100TR Liquid Scintillation Counter (PerkinElmer, Waltham, US). 10. Transfer of the counts into a spreadsheet for calculation of leucine transfer from the donor to the receiver compartment across the trophoblast cell layer. (b) Leucine transfer is calculated per well using the following equations: Scintillation count ½cpm - mean of NCB ½cpm mean of DC ½cpm = Leu transfer ½%of total dosage Leu transfer ½% × L - leucine conc: in transfer solution ½μmol=L = Leu transfer ½μM

4

ð4Þ

ð5Þ

Notes 1. It is recommended to use white-walled 96-well plates with flat clear bottom for luminescence-based quantification of radioactivity and black wall plates for fluorescence-based quantification of transmembrane diffusion in the LY measurements. 2. Not all commercially available 12-well plates are suitable for all 12-well inserts. Make sure that the Transwell® inserts are not touching the bottom in the 12-well plates. 3. Keep the electrodes dry or in 70% ethanol during a running experiment. 4. Treat tube with 70% EtOH, wipe with a paper towel and bring tube carefully to the laminar flow hood. 5. Use 15% FBS in the complete growth medium for 1–2 passages to accelerate cryo-recovery. 6. A pre-inhibition step can be applied by adding the inhibitor solutions for 30 min (T-0.5 h) at 37 °C before starting the uptake, if higher inhibition efficiency should be achieved. 7. Usually the TEER values first decrease after starting the stimulation with Forskolin, before they increase and even exceed the TEER values of the no-Forskolin control. 8. It is recommended that only one experimenter acquires all the measurements per experiment, since there is relatively large inter-individual variability.

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9. The sensor is designed with electrodes on two arms. One of them is longer and needs to be inserted in the lower compartment at the bottom, while the other is shorter and should be dipped into the medium of the upper compartment without touching the membrane of the insert. 10. Example for leucine uptake solution pipetting for ten upper or six lower compartments: (i) 15 μL L-leucine (100 mM stock) (ii) 15 μL L-glutamine (100 mM stock) (iii) 20 μL L-3H-leucine (1 mCi/mL stock) (radioactive) (iv) 9950 μL Hank’s buffer 11. There should be a remaining volume of at least 30% of the starting volume until the end of the transfer experiment. Recommended time points are: 0 min (take this sample before starting the transfer), 5 min, 30 min, 60 min, 90 min, 120 min, 180 min, 240 min, 300 min, 360 min. References 1. Roos S, Powell TL, Jansson T (2009) Placental mTOR links maternal nutrient availability to fetal growth. Biochem Soc Trans 37:295– 298. https://doi.org/10.1042/BST0370295 2. Burton GJ, Fowden AL, Thornburg KL (2016) Placental origins of chronic disease. Physiol Rev 96:1509–1565. https://doi.org/ 10.1152/physrev.00029.2015 3. Faichney GJ, White GA (1987) Effects of maternal nutritional status on fetal and placental growth and on fetal urea synthesis in sheep. Aust J Biol Sci 40:365–377. https://doi.org/ 10.1071/BI9870365 4. Paolini CL, Marconi AM, Ronzoni S, Noio MDI, Fennessey P, Pardi G, Battaglia FC (2001) Placental transport of leucine, phenylalanine, glycine, and proline in intrauterine growth-restricted pregnancies. J Clin Endocrinol Metabol 86:5427–5432. https://doi.org/ 10.1210/jcem.86.11.8036 5. Lager S, Powell TL (2012) Regulation of nutrient transport across the placenta. J Pregnancy 2012:179827. https://doi.org/10. 1155/2012/179827 6. Jansson T, Ekstrand Y, Bjo¨rn C, Wennergren M, Powell TL (2002) Alterations in the activity of placental amino acid transporters in pregnancies complicated by diabetes. Diabetes 51:2214–2219. https://doi.org/10. 2337/diabetes.51.7.2214 7. Harder T, Rodekamp E, Schellong K, Dudenhausen JW, Plagemann A (2007) Birth weight

and subsequent risk of type 2 diabetes: a metaanalysis. Am J Epidemiol 165:849–857. https://doi.org/10.1093/aje/kwk071 8. Eriksson J, Forsen T, Osmond C, Barker D (2003) Obesity from cradle to grave. Int J Obes Relat Metab Disord 27:722–727. https://doi.org/10.1038/sj.ijo.0802278 9. Boney CM, Verma A, Tucker R, Vohr BR (2005) Metabolic syndrome in childhood: association with birth weight, maternal obesity, and gestational diabetes mellitus. Pediatrics 115:e290–e296. https://doi.org/10.1542/ peds.2004-1808 10. Leon D a, Lithell HO, Vaˆgero¨ D, Koupilova´ I, Mohsen R, Berglund L, Lithell UB, PM MK (1998) Reduced fetal growth rate and increased risk of death from ischaemic heart disease: cohort study of 15 000 Swedish men and women born 1915–29. BMJ (Clinical Research ed) 317:241–245. https://doi.org/ 10.1136/bmj.317.7153.241 11. Sadovsky Y, Jansson T (2015) Placenta and placental transport function. In: Knobil and Neill’s physiology of reproduction: two-volume set 2, pp 1741–1782. https:// doi.org/10.1016/B978-0-12-397175-3. 00039-9 12. Fotiadis D, Kanai Y, Palacı´n M (2013) The SLC3 and SLC7 families of amino acid transporters. Mol Asp Med 34:139–158. https:// doi.org/10.1016/j.mam.2012.10.007

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13. Bodoy S, Fotiadis D, Stoeger C, Kanai Y, Palacı´n M (2013) The small SLC43 family: facilitator system l amino acid transporters and the orphan EEG1. Mol Asp Med 34:638–645. https://doi.org/10.1016/j.mam.2012. 12.006 14. Cleal JK, Glazier JD, Ntani G, Crozier SR, Day PE, Harvey NC, Robinson SM, Cooper C, Godfrey KM, Hanson MA, Lewis RM (2011) Facilitated transporters mediate net efflux of amino acids to the fetus across the basal membrane of the placental syncytiotrophoblast. J Physiol 589:987–997. https://doi.org/10. 1113/jphysiol.2010.198549 15. Gaccioli F, Aye ILMH, Roos S, Lager S, Ramirez VI, Kanai Y, Powell TL, Jansson T (2015) Expression and functional characterisation of system L amino acid transporters in the human term placenta. Reprod Biol Endocrinol 13:57. https://doi.org/10.1186/s12958015-0054-8 16. Kudo Y, Boyd CAR (2001) Characterisation of L-tryptophan transporters in human placenta: a comparison of brush border and basal membrane vesicles. J Physiol 531:405–416. https:// doi.org/10.1111/j.1469-7793.2001.0405i.x 17. Lewis RM, Glazier J, Greenwood SL, Bennett EJ, Godfrey KM, Jackson a. a., Sibley CP, Cameron IT, Hanson M a. (2007) L-serine uptake by human placental microvillous membrane vesicles. Placenta 28:445–452. https:// doi.org/10.1016/j.placenta.2006.06.014 18. Cleal JK, Brownbill P, Godfrey KM, Jackson JM, Jackson AA, Sibley CP, Hanson MA, Lewis RM (2007) Modification of fetal plasma amino acid composition by placental amino acid exchangers in vitro. J Physiol 582:871– 882. https://doi.org/10.1113/jphysiol. 2007.130690 19. Orendi K, Gauster M, Moser G, Meiri H, Huppertz B (2010) The choriocarcinoma cell line BeWo: syncytial fusion and expression of syncytium-specific proteins. Reproduction 140:759–766. https://doi.org/10.1530/ REP-10-0221 20. Zaugg J, Ziegler F, Nuoffer J-M, MoserH€assig R, Albrecht C (2021) Counter-directed leucine gradient promotes amino acid transfer across the human placenta. J Nutr Biochem 96: 108760. https://doi.org/10.1016/j.jnutbio. 2021.108760 21. Zaugg J, Huang X, Ziegler F, Rubin M, Graff J, Mu¨ller J, Moser-H€assig R, Powell T, Gertsch J, Altmann KH, Albrecht C (2020) Small molecule inhibitors provide insights into the relevance of LAT1 and LAT2 in materno-foetal amino acid transport. J Cell

Mol Med 24:12681–12693. https://doi.org/ 10.1111/jcmm.15840 22. H€afliger P, Graff J, Rubin M, Stooss A, Dettmer MS, Altmann KH, Gertsch J, Charles RP (2018) The LAT1 inhibitor JPH203 reduces growth of thyroid carcinoma in a fully immunocompetent mouse model. J Exp Clin Canc Res 37. https://doi.org/10.1186/s13046018-0907-z 23. Huang X, Lu¨thi M, Ontsouka EC, Kallol S, Baumann MU, Surbek D, Albrecht C (2016) Establishment of a confluent monolayer model with human primary trophoblast cells: novel insights into placental glucose transport. Mol Hum Reprod 0:gaw018. https://doi.org/10. 1093/molehr/gaw018 24. Johnson LW, Smith CH (1988) Neutral amino acid transport systems of microvillous membrane of human placenta. Am J Physiol Cell Physiol 254. https://doi.org/10.1152/ ajpcell.1988.254.6.c773 25. Desforges M, Mynett KJ, Jones RL, Greenwood SL, Westwood M, Sibley CP, Glazier JD (2009) The SNAT4 isoform of the system A amino acid transporter is functional in human placental microvillous plasma membrane. J Physiol 587:61–72. https://doi.org/10. 1113/jphysiol.2008.161331 26. Hoeltzli SD, Smith CH (1989) Alanine transport systems in isolated basal plasma membrane of human placenta Am J Physiol Cell Physiol 256 https://doi.org/10.1152/ajpcell.1989. 256.3.c630 27. Schio¨th HB, Roshanbin S, H€agglund MGA, Fredriksson R (2013) Evolutionary origin of amino acid transporter families SLC32, SLC36 and SLC38 and physiological, pathological and therapeutic aspects. Mol Asp Med 34:571–585. https://doi.org/10.1016/j. mam.2012.07.012 28. Pramod AB, Foster J, Carvelli L, Henry LK (2013) SLC6 transporters: structure, function, regulation, disease association and therapeutics. Mol Asp Med 34:197–219. https://doi. org/10.1016/j.mam.2012.07.002 29. Lassance L, Haghiac M, Leahy P, Basu S, Minium J, Zhou J, Reider M, Catalano PM, Hauguel-De Mouzon S (2015) Identification of early transcriptome signatures in placenta exposed to insulin and obesity. Am J Obstet Gynecol 212:647.e1–647.e11. https://doi. org/10.1016/j.ajog.2015.02.026 30. Karl PI, Tkaczevski H, Fisher SE (1989) Characteristics of histidine uptake by human placental microvillous membrane vesicles. Pediatr Res 2 5: 19–2 6. h ttps://doi.o rg/10 .1 203/ 00006450-198901000-00005

Placental Amino Acid Uptake and Transfer Assay 31. Novak DA, Beveridge MJ (1997) Glutamine transport in human and rat placenta. Placenta 18:379–386. https://doi.org/10.1016/ S0143-4004(97)80037-9 32. Dicke JM, Verges D, Kelley LK, Smith CH (1993) Glycine uptake by microvillous and basal plasma membrane vesicles from term human placentae. Placenta 14:85–92. https:// doi.org/10.1016/S0143-4004(05)80251-6 33. Miyamoto Y, Balkovetz DF, Leibach FH, Mahesh VB, Ganapathy V (1988) Na+ + Cl-gradient-driven, high-affinity, uphill transport of taurine in human placental brush-border membrane vesicles. FEBS Lett 231:263–267. https://doi.org/10.1016/0014-5793(88) 80744-0 34. Norberg S, Powell TL, Jansson T (1998) Intrauterine growth restriction is associated with a reduced activity of placental taurine transporters. Pediatr Res 44:233–238. https://doi. org/10.1203/00006450-199808000-00016

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35. Furesz TC, Moe AJ, Smith CH (1995) Lysine uptake by human placental microvillous membrane: comparison of system y+ with basal membrane. Am J Phys 268:C755–C761 36. Ayuk PT, Sibley CP, Donnai P, D’Souza S, Glazier JD (2000) Development and polarization of cationic amino acid transporters and regulators in the human placenta. Am J Physiol Cell Physiol 278:C1162–C1171 37. Hoeltzli SD, Kelley LK, Moe AJ, Smith CH (1990) Anionic amino acid transport systems in isolated basal plasma membrane of human placenta Am J Physiol Cell Physiol 259 https:// doi.org/10.1152/ajpcell.1990.259.1.c47 38. Moe AJ, Smith CH (1989) Anionic amino acid uptake by microvillous membrane vesicles from human placenta. Am J Phys 257. https://doi. org/10.1152/ajpcell.1989.257.5.c1005

Chapter 10 Assessing Cholesterol Efflux on Primary Human Trophoblast Cells Barbara Fuenzalida and Christiane Albrecht Abstract Cholesterol transport across the placenta must be tightly regulated to avoid a deficiency or an oversupply of cholesterol which is transferred from the mother to the fetus. In trophoblasts, the transport of cholesterol across the cell membrane is mainly mediated by the ATP-binding transporters, ABCA1 and ABCG1. The localization of the transporters at the apical and basal sides of syncytiotrophoblasts has been described. A frequently used method to quantify the amount of cholesterol that cells are capable of exporting is the cholesterol efflux assay. The principle of this assay is that when exogenous [3H]-labeled cholesterol is provided to cultured cells, the efflux of the radioactive cholesterol toward different acceptors in the culture medium is evaluated. Then, the percentage of cholesterol efflux from the cells to the acceptors is calculated. The present work gives an overview on the principle of this assay and a detailed protocol of this technique performed in primary trophoblasts isolated from human term placentas. Key words Cholesterol, Cholesterol efflux, Lipoproteins, Transporters, Primary cells, Trophoblast cells

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Introduction Cholesterol is required for the synthesis of cell membranes and signaling molecules, and serves as a precursor for steroid hormones [1, 2]. The developing fetus requires cholesterol and obtains it from endogenous synthesis and maternal lipoproteins [3, 4]. Although the developing fetus requires high amounts of cholesterol, its intracellular accumulation can be lethal for cells, so there are protective mechanisms to prevent its toxic effects [5]. In general, the cellular concentration of cholesterol is regulated by transcription factors that respond to lipids, and post-transcriptional and post-translational mechanisms that control biosynthesis, uptake, efflux, oxidative metabolism, and storage of this sterol [6]. The efflux of cholesterol toward different extracellular acceptors occurs through transporters that belong to the super-family of ATP-binding cassette (ABC) transporters. ABCA1 mediates the

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efflux of cholesterol from the plasma membrane to apolipoprotein A-I (apoA-I) [7]; ABCG1 is responsible for the efflux of cholesterol toward lipid-loaded spherical high-density lipoprotein (HDL) particles (also called mature HDL) [8]. Syncytiotrophoblasts are polarized cells responsible for the exchange of nutrients and other molecules between the maternal and fetal circulation. These cells have an apical membrane in contact with maternal blood and a basal membrane in contact with placental endothelial cells [9]. Due to the polarized nature of syncytiotrophoblasts, the expression of cholesterol transporters at either side of the cell membrane could be related to the flow of cholesterol to the fetal circulation (if expression is located at the basal membrane) or toward the maternal circulation (if expressed at the apical membrane). Cholesterol efflux into the maternal blood could help to protect the fetus against high levels of maternal cholesterol [10]. Various studies of cholesterol efflux have been carried out in trophoblast cells, such as the BeWo cell line, or cells at the cytotrophoblast and syncytiotrophoblast stage [10–14]. However, immortalized cells have been cultured for many passages and some of their original properties may have significantly changed over time. In this respect, the use of primary cultures presents a great advantage since they show a higher degree of physiology than immortalized cells. The cholesterol efflux assay is designed to quantify the rate of cholesterol efflux from cells. It reflects the capacity of cells to maintain cholesterol efflux and the capacity of plasma proteins to accept cholesterol released from cells. The cholesterol efflux assay is frequently used to determine the activity of cholesterol efflux transporters when affected by a treatment, disease, or different stimuli. The present work provides a detailed protocol of the cholesterol efflux assay in primary trophoblasts isolated from human term placenta. This technique consists of the following steps: (1) seeding cells, (2) radioactive cholesterol incubation, (3) equilibration, (4) acceptor incubation, (5) processing of samples, and (6) analysis of results.

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Materials Be sure to have all media and solutions prepared when starting the assay. 1. DMEM-HG + FBS (complete medium). Dulbecco’s modified Eagle’s medium containing 4.5 g/L glucose (DMEM-HG) including 10% Fetal bovine serum (FBS) and 1× antibioticantimycotic.

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2. Phosphate buffered saline (PBS): NaCl 136 mM, KCl 2.7 mM, Na2HPO4 7.8 mM, KH2PO4 1.5 mM, 37 °C, pH 7.4 or Dulbecco’s phosphate buffered saline (DPBS). 3. [3H]-Cholesterol in complete medium. Resuspend [3H]-Cholesterol in complete medium to a final concentration of 0.5 μCi/mL. Vortex and mix well. 4. DPBS + Bovine Serum Albumin - Fatty Acid Free (DPBS + BSA-FA-Free; dissolve BSA-FA-Free to a final concentration of 0.2 g/mL in DPBS. 5. DMEM-HG + BSA-FA-Free. Dissolve BSA-FA-Free to a final concentration of 0.2 g/mL in DMEM-HG serum free. 6. Scintillation liquid cocktail. 7. 1 N NaOH.

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Methods

3.1 Seeding the Cells and Labeling Cellular Cholesterol

Cytotrophoblasts are isolated from term placentas as previously described [13–16], by three trypsin/DNase enzymatic digestions for 30 min, at 37 °C, (trypsin: total activity: 615,000, 410,000, and 315,000 BAEE units, respectively, for each digestion; DNase: total activity: 60,000, 40,000, and 30,000 kilounits, respectively, for each digestion). The cellular pellets are resuspended in DMEM and separated by centrifugations (1500 RCF, 20 °C, 20 min) in a Percoll gradient (10–70%). Cytotrophoblast cells are obtained from gradient fractions between 35% and 55%. The following protocol can be performed for both cytotrophoblasts (24 h after seeding) or syncytiotrophoblasts (60 h after seeding) (see Note 1). 1. Resuspend primary cytotrophoblast cells and count them. 2. Seed the cells into 24-well plate (see Note 2) at a final density of 0.4 × 106 cells per well in 0.5 mL complete medium. The number of wells should be calculated for at least triplicate determinations for each experimental condition. 3. 12–24 h after seeding, cells are mainly at the cytotrophoblast stage, while usually after 60 h they have formed syncytiotrophoblasts. At the respective cells stage remove the medium and add the medium containing [3H]-Cholesterol to the wells with cells (final volume per well is 0.3 mL). 4. Incubate cells for 24 h in cell culture incubator (37 °C, 5% CO2) (see Note 3) (Fig. 1).

3.2

Equilibration

1. After 24 h of incubation with [3H]-Cholesterol-DMEM-HG, observe cells under microscope to ensure that they are alive and healthy.

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Fig. 1 Cholesterol efflux assay in primary trophoblast cells. The cholesterol efflux assay is a technique to evaluate the ability of the cell to remove cholesterol and/or the ability of extracellular acceptors to bind cholesterol. Trophoblast cells are (1) seeded at a defined density for a specific time, (2) incubated with [3H]Cholesterol for 24 h, (3) then washed and equilibrated with serum-free culture medium containing fatty acid (FA)-free BSA, (4) incubated with defined acceptors. (5) The samples are processed by lysing the cells and recovering the culture medium. (6) The counts are measured in a scintillation counter and results are calculated to obtain the % cholesterol efflux mediated by a specific acceptor

2. Wash cells with DPBS BSA-FA-free 3 times to remove media containing [3H]-Cholesterol (see Note 4). 3. Add 0.3 mL of DMEM-HG + BSA FA-free and serum free to each well. 4. Incubate for 4 h in the cell culture incubator at 37 °C and 5% CO2 (Fig. 1). 3.3 Acceptor Incubation

1. Prepare solutions of cholesterol efflux acceptors in DMEMHG + BSA-FA-free and serum free: ApoA-I (final concentration 10 μg/mL); HDL (final concentration 50 μg/mL) or serum (final concentration 5%) (see Notes 5 and 6). 2. Remove the medium that was added at the equilibration step. 3. Add 0.3 mL of medium containing the acceptors to each well and to the control (without acceptors) (see Note 6). 4. Incubate cells for 6 h in cell culture incubator at 37 °C and 5% CO2. Efflux time can vary between 1 and 8 h (see Note 7) (Fig. 1).

3.4 Sample Processing

1. Prepare scintillation vials (5 mL), add 2 mL of scintillation liquid to each vial. 2 vials per well are needed (one for the medium, one for the lysed cells) (see Note 8).

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2. After 6 h incubations with acceptors, observe cells under microscope, and ensure that the cells are alive. 3. Collect the medium (0.3 mL) and transfer it into the scintillation vial. 4. Wash cells with 0.3 mL of DPBS BSA-FA-Free and transfer the wash buffer in the same vial of step 3 in Subheading 3.4 (see Note 9). 5. Add 0.3 mL of 1 N NaOH to lyse the cells and incubate for 30 min at room temperature. 6. Once the cells have been released from the plate (check under the microscope), transfer the entire volume to a new scintillation vial and vortex it. Keep the sample at 4 °C until measurement. 7. Measure the samples in the scintillation counter: configure counter to count [3H] in CMPA units (Fig. 1). 3.5 Analysis of Results and Calculations

1. The radioactivity is determined both in the culture medium and in the cell lysates. The efflux is estimated as the fraction of radioactive signal [3H] in the medium compared to the total signal in the medium and the cells, according to the following formula (formula 1): %Cholesterol efflux =

CPM medium × 100% CPM medium þ CPM cells

CPM: counts per minute 2. Acceptor-mediated efflux is calculated by subtracting efflux without acceptor (% Cholesterol efflux basal) from efflux with acceptor (% Cholesterol efflux acceptor), according to the following formula (formula 2): %Cholesterol effluxðmediated by

acceptor Þ

= %Cholesterol effluxacceptor –%Cholesterol effluxbasal

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Notes 1. Using cytotrophoblasts or syncytiotrophoblasts depends on the objective of the study. 2. Cholesterol efflux in a 24-well plate is feasible. If it is performed in a 12- or 6-well plate, the cell density must be calculated with respect to the diameter of the well. 3. [3H]-Cholesterol is added to medium containing serum. It is assumed that cholesterol is incorporated into serum lipoproteins and these are taken up by the cells. 24 h labeling is

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sufficient time for lipoproteins to be taken up and for cholesterol to move from lipoproteins to cellular membranes. 4. The purity of albumin is an important factor to consider since it directly alters the binding and availability of lipids. It is imperative to use BSA without FA because FA present in less purified BSA preparations can induce biological effects by themselves [17]. Labeled cholesterol is balanced between various intracellular pools, and there are literature reports where the equilibrium time was prolonged to 24 h when no BSA was added. In general, the equilibrium time depends on the experiment and the cell type, and can be adjusted. 5. The type of acceptor determines specific routes of cholesterol efflux. For the determination of ABCA1-dependent cholesterol efflux apoA-I is used as acceptor while HDL is applied if ABCG1 and SR-BI-dependent pathways are being investigated. It is important to include a “blank” sample, which contains no acceptor, to measure the basal cholesterol efflux. 6. It is important to leave one set of wells (triplicates) without addition of acceptors to determine background efflux. 7. Both the concentration of the acceptors and the efflux time can be adapted to answer specific questions or to perform kinetic studies. Extra wells with cells must be included if more time points or different acceptor concentrations are tested. Generally, 3–6 h efflux time in presence of the acceptors is recommended. Incubations longer than 24 h could reflect a state of equilibrium indicating the acceptor’s capacity to accumulate cholesterol rather than the transporters’ activity to mediate cholesterol efflux. 8. The tubes with the scintillation liquid must be prepared before the end of the assay to avoid exceeding times for the cholesterol efflux with the acceptors. 9. The main transporters for cholesterol efflux are ABCA1 and ABCG1, and both are regulated by liver X Receptor (LXR). Cells can be pre-incubated with LXR agonists to increase cholesterol efflux activity.

Acknowledgments This work was supported by the Swiss National Science Foundation (SNSF) (Grant No. 310030_149958) and the Stiftung Lindenhof Bern (Grant No. 17-15-F).

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References 1. Cortes VA, Busso D, Maiz A et al (2014) Physiological and pathological implications of cholesterol. Front Biosci (Landmark Ed) 19: 416–428 2. Wild R, Weedin EA, Wilson D (2015) Dyslipidemia in pregnancy. Cardiol Clin 33:209–215 3. Palinski W (2009) Maternal-fetal cholesterol transport in the placenta: good, bad, and target for modulation. Circ Res 104:569–571 4. Woollett LA (2011) Review: transport of maternal cholesterol to the fetal circulation. Placenta 32:S218–S221 5. Tabas I (2002) Consequences of cellular cholesterol accumulation: basic concepts and physiological implications. J Clin Invest 110:905– 911 6. Soffientini U, Graham A (2016) Intracellular cholesterol transport proteins: roles in health and disease. Clin Sci (Lond) 130:1843–1859 7. Wang N, Silver DL, Costet P et al (2000) Specific binding of ApoA-I, enhanced cholesterol efflux, and altered plasma membrane morphology in cells expressing ABC1. J Biol Chem 275:33053–33058 8. Wang N, Lan D, Chen W et al (2004) ATP-binding cassette transporters G1 and G4 mediate cellular cholesterol efflux to highdensity lipoproteins. Proc Natl Acad Sci U S A 101:9774–9779 9. Jansson T, Myatt L, Powell TL (2009) The role of trophoblast nutrient and ion transporters in the development of pregnancy complications and adult disease. Curr Vasc Pharmacol 7: 521–533 10. Aye IL, Waddell BJ, Mark PJ et al (2010) Placental ABCA1 and ABCG1 transporters efflux cholesterol and protect trophoblasts from

oxysterol induced toxicity. Biochim Biophys Acta 1801:1013–1024 11. Jenkins KT, Merkens LS, Tubb MR et al (2008) Enhanced placental cholesterol efflux by fetal HDL in Smith -Lmli-Opitz syndrome. Mol Genet Metab 94:240–247 12. Sreckovic I, Birner-Gruenberger R, Besenboeck C et al (2014) Gestational diabetes mellitus modulates neonatal high-density lipoprotein composition and its functional heterogeneity. Biochim Biophys Acta 1841:1619– 1627 13. Kallol S, Huang X, Mu¨ller S et al (2018) Novel insights into concepts and directionality of maternal-fetal cholesterol transfer across the human placenta. Int J Mol Sci 19:2334 14. Fuenzalida B, Cantin C, Kallol S et al (2020) Cholesterol uptake and efflux are impaired in human trophoblast cells from pregnancies with maternal supraphysiological hypercholesterolemia. Sci Rep 10:5264 15. Huang X, Lu¨thi M, Ontsouka EC et al (2016) Establishment of a confluent monolayer model with human primary trophoblast cells: novel insights into placental glucose transport. Mol Hum Reprod 22:442–456 16. Fuenzalida B, Kallol S, Lu¨thi M et al (2021) The polarized localization of lipoprotein receptors and cholesterol transporters in the syncytiotrophoblast of the placenta is reproducible in a monolayer of primary human trophoblast. Placenta 105:50–60 17. Alsabeeh N, Chausse B, Kakimoto P et al (2018) Cell culture models of fatty acid overload: problems and solutions. Biochim Biophys Acta Mol Cell Lipids 1863:143–151

Chapter 11 RGD-Based Fluorescence to Assess Placental Angiogenesis Veronique Josserand, Jonathan Lavaud, Michelle Keramidas, Constance Collet, Wael Traboulsi, Pascale Hoffmann, Jean-Jacques Feige, Mohamed Benharouga, Jean-Luc Coll, and Nadia Alfaidy Abstract Normal fetal growth and placental development depend on active angiogenesis occurring at the fetomaternal interface throughout pregnancy. Nevertheless, reliable in vivo methods to assess placental angiogenesis are still missing. Here, we describe a quantitative and noninvasive in vivo method to specifically measure placental neovascularization in the gravid mouse. This method uses a technique based on the measurement of a fluorescent molecule Angiostamp700 that targets the alpha v beta 3 (αvβ3) integrin, a protein that is highly expressed by endothelial cells during the neovascularization and by trophoblast cells during invasion of the maternal decidua. Due to this noninvasive method, quantification of the fetomaternal angiogenic activity and information regarding the outcome of pregnancy are now possible. Key words In vivo angiogenesis, Fluorescence imaging, Angiostamp700, Neovascularization, Placental development, Fetomaternal interface

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Introduction Placental vascularization and subsequent angiogenesis are key processes that occur within the placental villi and at the fetomaternal interface to allow placental growth and establishment of the fetomaternal circulation, respectively [1–3]. While numerous in vitro methods have been established to assess placental vascularization and angiogenic activity, relatively few consistent and quantitative methods are available in vivo to assess this activity at the fetomaternal interface [4]. The available approaches are mainly based on color Doppler blood flow visualization and micro-ultrasound analyses [5]. The main weakness of these techniques remains their inability to provide quantitative information. Among proteins that contribute to the angiogenic processes at the fetomaternal interface are the integrins. In the placenta,

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integrins are known to participate in a number of key functions including cell migration, invasion, and adhesion [6–8]. Integrins comprise non-covalently bound α and β subunits that participate in cell-to-cell and cell-to-substratum adhesion [9]. Previous reports in the literature have shown that αv and β3 proteins are highly expressed in the mouse placenta with specific localizations to the endothelial and trophoblast cells present at the fetomaternal interface [10, 11]. Using 3D optical imaging of fluorescent probes, we have described a noninvasive and quantitative assessment of in vivo angiogenesis of subcutaneous sponges [12]. The probe used in this study is Angiostamp700, a fluorescent molecule that targets the αvβ3 integrin, which allows quantitative determinations of the angiogenic activity in vivo. Here, we describe a method that uses the same probe to assess the angiogenic activity at the fetomaternal interface in the gravid mouse in vivo.

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Materials 1. Female mice nulliparous around 8–12 weeks of age. 2. Surgical kit (clamps, scissors, and surgical suture). 3. Inhalant anesthesia device based on isoflurane vaporization. 4. Chemical anesthesia: 100 μg/kg medetomidine and 100 mg/ kg ketamine. 5. 25 G needle. 6. 1 mL syringes. 7. Ethylenediaminetetraacetic acid (EDTA). 8. Angiostamp700 is a cyclo decapeptide platform presenting 4 copies of the cyclic arginine-glycine-aspartic acid (cRGD) sequence known to target the αvβ3 integrin [13, 14]. 9. AlexaFluor®700 fluorescent dye was linked to the peptide platform to convert this reagent into an optical imaging probe called Angiostamp700 able to target αvβ3-expressing cells [15–17]. 10. 2D NIR fluorescence imaging device (Aequoria, Hamamatsu) with 660 nm light-emitting diodes equipped with interference filters for fluorescence excitation and a back-thinned charge-coupled device (CCD) camera cooled at -80 °C (ORCAII-BT-512G; Hamamatsu, Massy, France) and fitted with a high-pass RG 9 filter (Schott, Clichy, France) for fluorescence collection [15]. 11. Continuous-wave fluorescence diffuse optical tomography (fDOT) optical imaging system for deeper abdominal area exploration was used for noninvasive 3D fluorescence imaging

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of the neo-formed blood capillaries labeled with Angiostamp700. As previously described by Koenig et al. [16, 17], fDOT consists of a 690-nm laser source, a CCD camera, and a set of filters. The light source is a 35-mW compact laser diode (Power technology) equipped with a bandpass interference filter (Melles Griot 685AF30OD6). The emitted fluorescence is filtered by two 700-nm high-pass colored glass filters (Schott RG9 OD5) placed in front of a NIR-sensitive CCD camera (Hamamatsu ORCA AG) mounted with a f/15-mm objective (Schneider Kreutznach).

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Methods

3.1 αvb3 Labeling with Angiostamp700

1. Anesthetize gravid mice 24 h before fluorescence imaging.

3.2 2D-Fluorescence In Vivo Imaging

1. For 2D reflectance imaging, anesthetize mice with 4% air/isoflurane for induction and 1.5% thereafter.

2. Inject mice intravenously with 200 μL of 50 μM Angiostamp700 (Fluoptics, Grenoble, France) into the tail vein (Fig. 1) (see Note 1).

2. Perform whole-body 2D fluorescence imaging on the 4 faces of the mouse by positioning the mouse in ventral, dorsal, and lateral position (see Note 2). 3.3 3D-Fluorescence and microCT In Vivo Imaging

1. After whole-body 2D fluorescence imaging, place anesthetized mice (2.5% isoflurane/air) in a homemade mobile animal holder (Fig. 2) for bimodal microCT/fDOT imaging. 2. Perform 3D fluorescence acquisition and quantification using the fDOT system. The excitation sources are set as a regular 26 × 30-mm spaced grid over the abdominal area of the mouse, where the embryos are present. Two scans are successively performed for tissue attenuation evaluation and fluorescence measurement. The exposure time is automatically computed at each laser position to use the entire dynamic range of the

Fig. 1 Protocol of gravid mouse treatment. Angiostamp700 was injected 1 day before imaging at days 6.5 and 11.5 dpc. Mice were analyzed at days 7.5 and 1.25 dpc, respectively

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Fig. 2 Homemade mobile animal holder compatible for both fDOT and microCT. The anesthetized mouse is positioned between 4 Plexiglas plates so that it can be moved from the fDOT to the microCT without the animal moving at all. Small ceramic beads are embedded in the edges of the plates to serve as landmarks for the resizing and alignment of the respective reconstructed volumes from fDOT and microCT

camera. The two stacks of diffusion and fluorescence images are compiled by the reconstruction algorithm to generate a 3D image (see Notes 3 and 4). 3. Perform 3D reconstruction. The fDOT principle lies in the ability to both reconstruct fluorescence, even in highly heterogeneous attenuating media, and handle complex geometries. The results are presented as a 3D view of the reconstructed area. The reconstructed area is a volume meshed with a 2-mm sample rate in the x and y directions and 1 mm in the z direction (depth) that yields a size of approximately 8 × 10 × 15 voxels and may vary slightly depending on animal thickness. Fluorescence was reconstructed in z cross-sections. The cross-sections are presented from bottom to top for z = 0 (ventral side) and z = 15 (dorsal side). The superimposition of the reconstructed volumes viewed as a smooth interpolation perspective and positioned on top of the white-light image of the animal allowed for the generation of the final image. The procedure time on a 3-GHz intel Xeon was 10 min to reconstruct the fluorescence distribution. Each fluorescence reconstruction is presented with the same color scale to allow for visual comparison (see Note 5).

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1. Sacrifice mice by intravenous injection of 400 mg/kg Exagon and harvest uterine horns and placentas for ex vivo fluorescence imaging. 2. Perform fluorescence imaging using a NIR 2D-fluorescence reflectance imaging device (Fluobeam®700, Fluoptics, France). Excitation was achieved using a class 1 expanded laser source at 680 nm and the fluorescence signal was collected using a CCD equipped with a high pass filter >700 nm. Expose tissues from non-injected control mice to the camera to set the autofluorescence level. 3. For image analyses, adjust region of interests (ROIs) on each tissue sample. Express as mean fluorescence in the ROIs (relative light unit per pixel per 100 milliseconds: RLU/pixel/ 100 ms).

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Notes 1. Intravenous injection of Angiostamp700 is performed in the tail vein after mild inhalant anesthesia (4% air/isoflurane for induction and 1.5% thereafter). A compress moistened with warm water may be applied on the tail for few seconds to slightly vasodilate the vein and facilitate the intravenous injection. 2. For noninvasive fluorescence imaging, mice need to be carefully depilated using a clipper and then applying depilatory cream for no more than 1 min before rinsing carefully with warm water. 3. Mice should be thoroughly anesthetized during 3D imaging to reduce fetal motion and associated artifacts in image reconstruction. 4. In combination with sustained anesthesia, body temperature should be carefully controlled. The mice need to be warmed using a heated animal holder for the entire duration of image acquisition. 5. To better locate the 3D fluorescence signal within the wholebody mouse anatomy, we performed a medium resolution microCT (VivaCT 40 ScancoMedical) with a 42 μm isotropic voxel size, a voltage of 45 kV, and a current of 114 mA. The 3D fluorescence volume is merged with the mice skeleton 3D view in order to replace the fluorescent signal in an anatomical context.

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Acknowledgments We acknowledge the following sources of funding: INSERM (U1292), University Grenoble-Alpes Fourier ANR-17-EURE0003, Commissariat a` l’Energie Atomique (IRIG/DS/Biosante´) and Institute for Advanced Biosciences, INSERM-UGA U1209, CNRS UMR 5309. The Optimal imaging platform is supported by France Life Imaging (French program “Investissement d’Avenir” grant; “Infrastructure d’avenir en Biologie Sante´”, ANR-11-INBS0006) and the IBISA French consortium “Infrastructures en Biologie Sante´ et Agronomie”. Imaging systems used in this study were purchased thanks to France Life Imaging (French program “Investissement d’Avenir” grant; “Infrastructure d’avenir en Biologie Sante”, ANR-11-INBS-0006) and the Contrat Plan EtatRe´gion Auvergne-Rhone-Alpes. References 1. Alfaidy N et al (2020) The emerging role of the prokineticins and Homeobox genes in the vascularization of the placenta: physiological and pathological aspects. Front Physiol 11:591850 2. Bardin N, Murthi P, Alfaidy N (2015) Normal and pathological placental angiogenesis. Biomed Res Int 2015:354359 3. Brouillet S et al (2010) Molecular characterization of EG-VEGF-mediated angiogenesis: differential effects on microvascular and macrovascular endothelial cells. Mol Biol Cell 21(16):2832–2843 4. van Oppenraaij RH et al (2009) Vasculogenesis and angiogenesis in the first trimester human placenta: an innovative 3D study using an immersive virtual reality system. Placenta 30(3):220–222 5. Stevenson GN et al (2018) Automated visualization and quantification of spiral artery blood flow entering the first-trimester placenta, using 3-D power Doppler ultrasound. Ultrasound Med Biol 44(3):522–531 6. Cross JC, Werb Z, Fisher SJ (1994) Implantation and the placenta: key pieces of the development puzzle. Science 266(5190): 1508–1518 7. Damsky C, Sutherland A, Fisher S (1993) Extracellular matrix 5: adhesive interactions in early mammalian embryogenesis, implantation, and placentation. FASEB J 7(14):1320–1329 8. Zhou Y et al (1997) Human cytotrophoblasts adopt a vascular phenotype as they differentiate. A strategy for successful endovascular invasion? J Clin Invest 99(9):2139–2151 9. Gurtner GC et al (1995) Targeted disruption of the murine VCAM1 gene: essential role of

VCAM-1 in chorioallantoic fusion and placentation. Genes Dev 9(1):1–14 10. Bowen JA, Hunt JS (1999) Expression of cell adhesion molecules in murine placentas and a placental cell line. Biol Reprod 60(2):428–434 11. Sutherland AE, Calarco PG, Damsky CH (1993) Developmental regulation of integrin expression at the time of implantation in the mouse embryo. Development 119(4): 1175–1186 12. Keramidas M et al (2013) Noninvasive and quantitative assessment of in vivo angiogenesis using RGD-based fluorescence imaging of subcutaneous sponges. Mol Imaging Biol 15(3): 239–244 13. Aumailley M et al (1991) Arg-Gly-Asp constrained within cyclic pentapeptides. Strong and selective inhibitors of cell adhesion to vitronectin and laminin fragment P1. FEBS Lett 291(1):50–54 14. Boturyn D et al (2004) Template assembled cyclopeptides as multimeric system for integrin targeting and endocytosis. J Am Chem Soc 126(18):5730–5739 15. Jin ZH et al (2007) In vivo optical imaging of integrin alphaV-beta3 in mice using multivalent or monovalent cRGD targeting vectors. Mol Cancer 6:41 16. Koenig A et al (2010) Fluorescence diffuse optical tomography for free-space and multifluorophore studies. J Biomed Opt 15(1): 016016 17. Koenig A et al (2008) In vivo mice lung tumor follow-up with fluorescence diffuse optical tomography. J Biomed Opt 13(1):011008

Chapter 12 Real-Time Assessment of Mitochondrial Function in Cytotrophoblast and Syncytialized Trophoblast Cells Using the Seahorse XFe24 Extracellular Flux Analyzer O’Llenecia S. Walker, Linda L. May, and Sandeep Raha Abstract Physiological processes utilize variable amounts of energy required for optimal growth, development, and survival. This energy is supplied by total intracellular adenosine triphosphate (ATP) and is mainly generated by mitochondrial oxidative phosphorylation and to a lesser extent via glycolysis. Here, we provide a detailed protocol for obtaining measurements of energy metabolism using the Seahorse XFe24 Extracellular Flux Analyzer. Specifically, this assay measures mitochondrial oxidative phosphorylation based on oxygen consumption rate (OCR) and glycolysis by analyzing the extracellular acidification rate (ECAR) via realtime live cell analysis. Using trophoblast cell lines, this protocol focuses on analyzing mitochondrial respiration for both cytotrophoblasts and syncytiotrophoblasts. Key words OCR, Mitochondria, Oxidative phosphorylation, Seahorse XF24 Extracellular Flux Analyzer, Trophoblasts

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Introduction Several investigations have linked mitochondrial dysfunction to adverse health outcomes and drug-induced toxicity [1–4]. Consequently, there is a need for high-throughput screening methods for assessing the impact of various drugs on mitochondrial function [5]. The extracellular flux (XF) assay described herein is a platebased method in which trophoblast cell lines HTR8/SVneo and BeWo were acutely exposed to test compounds, then real-time changes in oxygen consumption rate (OCR) were measured in live cells using a Seahorse XFe24 Extracellular Flux Analyzer. The Seahorse XFe24 Extracellular Flux Analyzer continuously measures oxygen concentration and proton flux in the supernatant over time [6]. Those measurements are converted to OCR and extracellular acidification rate (ECAR) values that enable a direct quantification of mitochondrial respiration and glycolysis,

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respectively. This protocol focuses on measuring OCR as most of the intracellular ATP is generated from mitochondrial respiration. Using this protocol, we sought to quantify basal mitochondrial function and mitochondrial stress on two different trophoblast cell lines in response to tetrahydrocannabinol (THC), the psychoactive component of cannabis [7, 8] to delineate its mechanism of action. Cells were seeded in 24-well microtiter XFe24 culture plates and maintained for 48 h with THC treatment. Prior to analysis, the treated culture media was replaced with XF base medium, and cells were equilibrated in a non-CO2 incubator immediately before executing the metabolic flux analysis to ensure precise measurements of values that are pH sensitive. OCR was measured under basal conditions and after injection of compounds through drug injection ports. This protocol allowed for the assessment of the basal metabolic profiles of the two trophoblast cell lines as well as key parameters of mitochondrial function in response to THC and by sequential addition of mitochondrial perturbing agents oligomycin, carbonyl cyanide-4-(trifuoromethoxy) phenylhydrazone (FCCP), and rotenone/antimycin A. The OCR values were normalized to total cell protein content from each well using the bicinchoninic assay (BCA) [7, 8].

2

Materials Prepare all solutions at room temperature in a cell culture hood and store in the refrigerator at 4 °C (or as otherwise indicated). Equipment 1. XFe24 Extracellular Flux Analyzer (Agilent) 2. CO2 incubator 3. Heated/humidified incubator set to 37 °C; not a CO2 supplied incubator Solutions 1. Mitochondrial stress test medium: 100 mL XF base medium supplemented with 5 mL sodium pyruvate (100 mM), 5 mL Lglutamine (200 mM), 5 mL of 45% glucose solution (see Note 1). Adjust the pH of the mitochondrial stress test media to 7.4, sterile filter using a 0.2 micron filter and store at 4 °C. Warm media to 37 °C and re-adjust the pH at 37 °C prior to use (see Note 2). 2. Oligomycin: Prepare 100 μM stock solution in assay medium (see Note 3). 3. FCCP: Prepare 100 μM stock solution in assay medium (see Note 3). 4. Rotenone/antimycin A: Prepare 50 μM stock solution in assay medium (see Note 3).

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3

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Methods The protocol outlined here is for a 24-well microtiter plate. Volumes will need to be adjusted if another plate format is used. Seeding of Cells 1. Optimize cell density to achieve a confluent monolayer after 24–48 h of culture (see Note 4). 2. Seed cells in 250 μL of media (see Note 5). 3. Do not seed blank wells with cells. Ensure there are a minimum of 3 wells as blanks to which cell culture media will be added (see Note 6). 4. Place the plate in CO2 incubator overnight. Day Before Assay 1. Open XFe24 Flux Assay Kit. The kit contains the following: • Cartridge Lid (clear) • Sensor Cartridge (green) • Hydro Booster (pink) • Utility Plate 2. Place the Sensor Cartridge next to the Utility Plate (Fig. 1). 3. Fill each well of the Utility Plate with 1 mL of sterile water (DNase and RNase free). 4. Place the Hydro Booster on top of the Utility Plate. 5. Lower the Sensor Cartridge through the openings on the Hydro Booster plate, into the Utility Plate, submerging the sensors in the sterile water (see Note 7).

Fig. 1 Utility cartridge set up. (a) Place the sensor cartridge (green) next to the utility plate. Fill each well of the utility plate with 1 mL of sterile water. (b) Place the hydro booster (pink) on top of the utility plate. Lower the sensor cartridge through the openings on the hydro booster plate, into the utility plate, submerging the sensors in the sterile water. The underside of the sensor cartridge shows the sensors with the embedded fluorophores

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6. Place the cartridge assembly in a non-CO2 37 °C incubator overnight (see Note 8). Day of Assay 1. Following the overnight incubation, remove the cartridge assembly from the incubator. Lift the sensor cartridge completely out of the water and utility plate. 2. Discard the water from the utility plate and replace with 1 mL of pre-warmed (37 °C) Seahorse calibrant. 3. Lower the sensor cartridge into the utility plate submerging the sensors into the calibrant. 4. Place the assembled sensor cartridge with utility plate in a non-CO2 37 °C incubator for 1 h prior to loading drug ports of the sensor cartridge (see Note 9). 3.1 Washing the Cells

1. Warm the pre-made assay medium to 37 °C. 2. Remove the treated cells from the incubator and check under a microscope for cell health, morphology, and contamination. 3. Using a p1000 pipet remove all but 50 μL of growth media (set p1000 to 200 μL) and carefully wash once (this is to remove the FBS) with 1 mL of pre-warmed assay media (see Note 10). Remove 1 mL of wash media leaving 50 μL behind (total volume remaining in the wells is now 50 μL). 4. Add 475 μL of the warmed assay media to all wells including blank wells and check under the microscope to ensure that no cells were washed away (see Note 11). 5. Place plate in a non-CO2 37 °C incubator for 1 h prior to commencing assay. 6. Keep the assay media warm at 37 °C.

3.2 Preparing Stock Compounds (See Note 12)

1. Remove compounds from the foil pack and let equilibrate to room temperature for 15 min, and prepare stock compounds (see Note 13). 2. Prepare stocks by adding assay medium (Table 1) and pipet up and down gently exactly 10 times (see Note 14).

Table 1 Preparation of stock compounds Volume of assay medium

Final stock concentration

Oligomycin

630 μL

100 μM

FCCP

720 μL

100 μM

Rotenone/antimycin A

540 μL

50 μM

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A Seahorse XFe24

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Constant Loading Volume/Variable Compound Concentration. Starting well volume: 525 µL assay medium Final well concentration (µM)

Stock volume (µL)

Medium volume (µL)

Total volume in each port (µL)

Port A: Oligomycin

1.0

240

2760

3000

Port B: FCCP

2.0

540

2460

3000

Port C: Rotenone/Antimycin A

0.5

300

2700

3000

B A

B

Injection port Fluorescent sensors

C

D

Fig. 2 Diluting and loading uncoupling compounds into sensor cartridge. (a) Uncoupler concentrations used in trophoblast cells. (b) Orientation of the ports in assembled cartridge. This is the order of the ports. Load all the “A” ports first, then all the “B” ports, etc.

3.3 Loading Compounds into Sensor Cartridge (See Note 15)

1. Retrieve the utility plate from non-CO2 incubator. To each port, add 75 μL (see Notes 16 and 17; Fig. 2a, b). Load all the “A” ports first, then all the “B” ports, etc. (Fig. 2b). Once completed, the Seahorse assay can begin. If using Seahorse Desktop Software: 2. Turn on Wave software and ensure the heater is on. Let it warm up at 37 °C for at least 5 h prior to running assay. If the assay is to be run the following morning, warm up Wave overnight as this will save time. 3. Browse for and open the saved design file as follows: (a) Start Wave (b) Select Open (c) Select Browse (d) Locate the template file (.asyt) or design file (.asyd) (e) Select the template/design file (f) Click “Open”

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(g) Select the “Review and Run” tab, and then click “Start Run”. (If starting with a new experiment/design, see Note 18.) 4. Remove cartridge lid and hydro boost plate (pink plate) BEFORE loading it into the Seahorse and verify correct plate orientation: i.e., A1 must be aligned with the bottom left for loading. 5. When prompted, place the loaded sensor cartridge with the calibrant plate into the instrument, then click “I’m ready”. Calibration will take approximately 15–30 min (see Note 19). 6. Following calibration and equilibration of the cell culture microplate, when prompted, click “I’m ready”. Load the cell culture microplate and click “I’m ready” to run the assay (see Note 20). 7. When the run is complete, remove the plate and carefully wash once with ice-cold 1X phosphate buffered saline (PBS; see Note 21). Proceed to perform BCA analysis. (a) If it is not convenient to begin the BCA assay right after the Seahorse protocol, remove as much PBS as possible and wrap the plate and lid in parafilm and place in a -20 °C freezer upside down until the BCA can be performed (see Note 22). 3.4

BCA Analysis

The BCA Pierce kit from Thermo-Fisher Scientific was used in this protocol 1. If the plate was frozen following the Seahorse protocol, take the plate out of freezer and place face down on ice lined with paper towel without lid (see Note 23). 2. Using a pipet, carefully remove residual PBS from the wells. Add 100 μL of RIPA to each well containing cells (see Note 24). 3. Add the lid to the plate and agitate on a plate shaker in the cold room for 15 minutes (400 rpm). 4. Turn on and set up the plate reader. Name the protocol. 5. Make the standards from the BCA kit as serial dilutions (see Pierce protocol). 6. Make the BCA reagent as per the Pierce protocol (see Note 25). 7. Add 25 μL of samples and standards in duplicate (see Note 26). 8. Add 200 μL of working reagent to each well with a multichannel pipet and seal plate with plate sealer. Agitate the plate on a plate shaker (~400 rpm) for 30 s at room temperature. Incubate the plate for 30 min at 37 °C degrees.

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9. Allow the plate to cool to room temperature and then place it in the plate reader and read in “end point” at 562 nm. 10. Plot the standard curve and do calculations to obtain protein concentration in μg/μL. 11. From the y = mx + b equation, solve for “x”. 12. In a separate Excel sheet, normalize the Seahorse data well-forwell with the protein concentration obtained in the above calculations. 13. Calculate OCR (pmoles/min/μg/mL). 14. Do this for all the time points. OCR is obtained from the excel data retrieved from the Seahorse instrument. 15. Once this is completed for all the wells, average the experimental replicates (i.e., quadruplicates) and this is the data that should be copied into the graphing and statistics software (i.e., GraphPad Prism).

4

Notes 1. Correct the pH of this Assay Media with 0.1 N NaOH to pH 7.4 at 37 °C. This is critical because the assay is temperature and pH sensitive. 2. It is important to use XF medium which lacks bicarbonate. Bicarbonate will buffer the media as it will dissociate to form CO2 and water. CO2 will degas during the assay, causing intracellular shifts in pH that will interfere with the OCR readings [9]. 3. Stocks are to be used the day they are reconstituted only. It takes 10–15 min to prepare the stock solutions. 4. The optimizing of seeding density is especially important if one is culturing and treating on the Seahorse™ assay plates. We have found the growth characteristics on this surface to different from those of standard tissue culture surfaces. Seeding density optimization is performed ahead of beginning the experiments for this assay. The timing is dependent upon the length of the treatment course. The recommended cell number for assessment of trophoblast cell line is between 10 and 80 K cells/well, but this may need to be determined based on the cell type being used. By visualizing under a microscope, verify that the cells are adherent to the cell culture surface. A monolayer of cells must be established to minimize sloughing off cells during the assay. 5. This volume is important to ensure cells adhere to the plate. Only the bottom of the wells is tissue culture treated.

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6. Do not proceed with empty wells because the machine needs a liquid volume in each well to work properly. 7. Verify, via visual inspection, that the water is high enough to keep the sensors submerged. The sensors contain fluorophores associated with O2 and H+ for each well; therefore, they must be hydrated to work properly. 8. To prevent evaporation of the water, verify that the incubator is properly humidified (an open bottle of ddH2O inside the incubator is sufficient). It is important to use a non-CO2 incubator to keep the pH stable. 9. This will help prevent any air bubbles that may be trapped in the overnight incubation and will prevent a falsely elevated OCR. 10. Do not use an aspirator to remove the growth media, as the suction may result in pulling the cells away from the cell culture surface. Add the pre-warmed assay media wash very slowly to the walls of the wells to avoid washing the cells away. 11. There will be a final volume of 525 μL/well at this point. 12. Prepare stock compounds 30 min after placing the culture plate from step 6 above into the non-CO2 incubator (it takes 10–15 min to prepare the stock solutions). 13. Stocks are to be used the day they are reconstituted only. Any remaining stock must be discarded. 14. Pipetting too vigorously may result in loss of stock volume as it will spill over the tube. 15. There are two approaches to loading the injection ports of the sensor cartridge: “Constant Loading Volume/Variable Compound Concentration” or “Constant Compound Concentration/Variable Loading Volume”. This protocol utilized follows the former approach. Optimization of each uncoupler for each cell type ought to be determined by the researcher and must be done prior to knowing which concentration is best to use. For the trophoblast cells used in this protocol, this has been done according to the Seahorse protocol. Figure 2a represents the dilutions used with these trophoblast cells. 16. There is no need to touch the bottom of the ports to add the stocks. Each series of ports must contain the same volume and all wells must be loaded, even those not used in the experiment, otherwise the compounds will not be injected (the same volume of assay media can be loaded into the ports of unused wells). Port D is for acute pre-treatment, which was not used by our research group. 17. Some stock will remain and since it can’t be reused, it must be discarded. When setting up the tubes to create the dilutions for

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the uncouplers (Fig. 2a), the following is recommended: (1) set up 3 × 15 mL tubes in a holder and keep them in order with respect to ports A, B, and C. (2) Use a 5 mL or 10 mL serological pipet to load 2 mL of media into each tube. (3) Use a p1000 pipet to load the difference. 18. Setting up a new plate template: • Creating a new Assay Template or Design on Wave Controller: (a) Navigate to Wave Home then click “Select New”. (b) Select Blank from the list of available Templates. (c) Click the “Design” button to create a New Design tab. (d) Define groups and conditions: The Group Definitions tab is used to define the Injection Strategy, Pre-treatments, Assay Media, and Cell Type to be used in the assay. • The 4 different types of assay conditions are: (a) Injection Strategies. (b) Pre-treatments. (c) Assay Media. (d) Cell Type(s). • Map groups to the plate: The Plate Map is where Wave displays information about each well in the experiment. Name the wells based upon the experimental conditions as per the map used in the cell culture microtiter plate. Indicate which wells are the control wells and which wells are empty. • Define the assay protocol: Click the “Instrument Protocol” tab to define the assay protocol (including measurement and injection cycles) after completing the Groups/Conditions and Plate Map. For further details on setting up the machine and the software, consult the manufacturer’s website. • Review and run the assay. If there is nothing in port 4, this part of the protocol can be removed prior to starting. • Any errors in set-up will prevent the instrument from starting. Ensure to remove Pink hydro boost plate. When ready, press “start” and allow the calibration to begin. • When the calibration is complete, the instrument will indicate that it is ready to have the calibration plate removed and replaced it with the plate containing cells. Ensure the lid is off then press “start”. 19. The machine scans the barcode and can detect the expiration date. The machine will also detect oxygen and pH levels during this calibration period.

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Fig. 3 Illustration and schematic of oxidative phosphorylation and representative OCR tracing. (a) Schematic showing the action of the metabolic drugs used in the extracellular flux assays. (b) Representative mitochondrial stress test. After trophoblast cells were treated with THC for 48 h, OCR was measured at baseline and following consecutive injections of oligomycin (1 μM), FCCP (2 μM), and rotenone/antimycin A (0.5 μM) [7, 8]

20. Total assay time is approximately 90 min. A sample of OCR is depicted in Fig. 3 along with the potential sites of pharmacological inhibition in the ETC. 21. When performing the PBS washes, carefully aim for the walls of the wells to avoid washing away the adherent cells. 22. The plate is placed upside down to prevent residual PBS from diluting the protein. 23. To minimize any residual PBS dripping back into the plate, which would dilute the protein, wipe away any PBS that may have dripped on lid when thawed. 24. No protease nor phosphatase inhibitors required. This protein is only being used to normalize the Seahorse data. 25. The working reagent is stable for several days in a closed container at room temperature. 26. Do this in triplicates if there is enough room on the BCA plate.

Acknowledgments This work was supported by CIHR and NSERC for the Collaborative Health Research Program award grant to S.R. References 1. Natale BV, Gustin KN, Lee K, Holloway AC, Laviolette SR, Natale DRC, Hardy DB (2020) Δ9-tetrahydrocannabinol exposure during rat pregnancy leads to symmetrical fetal growth restriction and labyrinth-specific vascular defects in the placenta. Sci Rep 10:1–15

2. Peng KY, Watt MJ, Rensen S, Willem Greve J, Huynh K, Jayawardana KS, Meikle PJ, Meex RCR (2018) Mitochondrial dysfunction-related lipid changes occur in nonalcoholic fatty liver disease progression. J Lipid Res 59:1977–1986

Oxygen Polarography of Trophoblast Cells 3. Zhao J, Zhang J, Yu M, Xie Y, Huang Y, Wolff DW, Abel PW, Tu Y (2013) Mitochondrial dynamics regulates migration and invasion of breast cancer cells. Oncogene 32:4814–4824 4. Wolff V, Schlagowski A-I, Rouyer O, Charles A-L, Singh F, Auger C, Schini-Kerth V, Marescaux C, Raul J-S, Zoll J, Geny B (2016) Tetrahydrocannabinol induces brain mitochondrial respiratory chain dysfunction and increases oxidative stress: a potential mechanism involved in cannabis-related stroke. Biomed Res Int 28: 101–102 5. Wills L (2017) The use of high-throughput screening techniques to evaluate mitochondrial toxicity. Toxicology 391:34–41 6. Wu M, Neilson A, Swift AL, Moran R, Tamagnine J, Parslow D, Armistead S, Lemire K, Orrell J, Teich J, Chomicz S, Ferrick DA (2007) Multiparameter metabolic analysis

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reveals a close link between attenuated mitochondrial bioenergetic function and enhanced glycolysis dependency in human tumor cells. Am J Physiol Cell Physiol 292:125–136 7. Walker OS, Gurm H, Sharma R, Verma N, May LL, Raha S (2021) Delta-9-tetrahydrocannabinol inhibits invasion of HTR8/SVneo human extravillous trophoblast cells and negatively impacts mitochondrial function. Sci Rep 11:1– 15 8. Walker OS, Ragos R, Gurm H, Lapierre M, May LL, Raha S (2020) Delta-9-tetrahydrocannabinol disrupts mitochondrial function and attenuates syncytialization in human placental BeWo cells. Physiol Rep 8:1–17 9. Krycer J, Fisher-Wellman K, Fazakerley D, Muoio D, James D (2017) Bicarbonate alters cellular responses in respiration assays. Biochem Biophys Res Commun 489:399–403

Chapter 13 Quantifying Trophoblast Function Using the xCELLigence System Rosemary J. Keogh and Padma Murthi Abstract Analyses of trophoblast behavior in vitro are widely used to investigate and understand the trophoblast dysfunction associated with pregnancy pathologies. Key behaviors critical for trophoblast function include adhesion, proliferation, migration, and invasion. Traditionally, these behaviors have been measured using time-consuming single timepoint or endpoint assays. xCELLigence is a breakthrough technology that utilizes gold electrodes embedded in cell culture plates to collect electrical impedance measurements and monitor cell attachment, spreading, movement, and growth. It enables trophoblast functions to be quantitatively monitored in real-time, providing new insights into trophoblast behavior. Key words Adhesion, Extravillous trophoblast, Impedance, Invasion, Migration, Proliferation, xCELLigence

1

Introduction Human placental development is an extremely complex and spatiotemporally orchestrated process. The extravillous trophoblasts play a critical role in this process and ensure successful pregnancy outcomes by remodeling the maternal uterine vasculature to create a high-flow, low-resistance blood supply. These processes ensure that sufficient oxygen and nutrients are delivered to the fetus so that it can grow and develop normally. In the absence of successful vessel remodeling, pregnancy complications such as pre-eclampsia can arise. This has serious health impacts for both mother and baby, and can lead to perinatal death and premature delivery. Understanding the behavior and function of extravillous trophoblasts is central to defining the steps that lead to a successful pregnancy and the serious consequences when the processes are disrupted or defective.

Sandeep Raha (ed.), Trophoblasts: Methods and Protocols, Methods in Molecular Biology, vol. 2728, https://doi.org/10.1007/978-1-0716-3495-0_13, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Cell behaviors critical to extravillous trophoblast function include the ability to proliferate, adhere, migrate, and invade. Traditionally, the only option to monitor these functions in real-time was to use time-lapse video capture. However, this technique involves laborious post-experimental analysis to derive quantitative results. Other methods such as scratch or wound healing assays for investigating cell migration, or trans-well or Boyden chamber assays for cell invasion only provide data at discrete or end time points. Similarly, proliferation assays determine cell number at defined time points. Defining the proliferation and growth characteristics requires a large number of assays at different times, an approach that is both resource- and time-consuming. The xCELLigence system (Agilent Technologies Inc., see Note 1) is a technological breakthrough that enables quantitative monitoring of cell behaviors such as proliferation and migration in realtime. Through the use of specially designed cell culture plates with embedded gold electrodes, cell attachment can be determined by interference to an electrical signal passed through the electrodes. The electrical signal can be repeatedly pulsed at intervals as short as seconds and when combined with measuring the interruption, or impedance, of the electrical signal, effectively enables continuous monitoring of cells in real-time. Importantly, neither the electrodes nor the electron flow through them has any impact on cell function or viability. The impedance measurements are used to derive an arbitrary unit called the cell index (CI). The cell index is calculated from the difference between the impedance measured at any time point (Rn) and the background impedance reading taken at the start of the experiment prior to the addition of cells (Rb); CI = (Rn-Rb)/15. There are a range of xCELLigence analyzers available from a single plate model for investigating proliferation, determining viral titers, and observing cytopathic effects, to a high throughput model which can simultaneously monitor 1526 wells by integrating 4 instruments to undertake studies such as antibody screening (see Note 1). The use of the xCELLigence technology for quantifying trophoblast function is a major technological advance that affords several advantages over traditional assays. Extravillous trophoblast cells such as HTR-8/SVneo, SGH-PL4, TEV-1, or Swan-1 all have different replication and growth characteristics. The ability to monitor cells in real-time with the xCELLigence system enables observation of the lag and log phases of cell attachment and growth. This facilitates experimental design by enabling optimum time points for cell stimulation or harvesting to be determined, which can be used not only for the functional assays described in this chapter but also to inform the design of other types of experiments. Furthermore, there is no need to add labels, remove cells from plates, or manipulate the cells in any other way. The technology is time-saving, easy

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to use, and allows for protocols to be altered during experiments to enable adjustments based on the data that is being collected. Most significantly, the xCELLigence system provides quantitative data in real-time. The methods described in this chapter are for the Agilent xCELLigence real-time cell analysis (RTCA) dual-purpose (DP) system which uses 16-well plates (see Note 2). We present a generic protocol based on our previous work [1, 2] which can be easily adjusted to suit the requirements of many different experimental conditions and cell types.

2 2.1

Materials Equipment

1. Agilent xCELLigence RTCA-DP. 2. Laptop or desktop computer running RTCA software Pro. 3. E-plate 16. 4. CIM-plate 16. 5. CIM-plate 16 assembly tool. 6. Incubator (37 °C, 95% air/5% CO2). 7. Cell culture flasks. 8. 50 mL Falcon centrifuge tubes. 9. Manual or automated cytometer. 10. Cell culture hood.

2.2

Cells

1. The methods described were developed for the HTR-8/SVneo extravillous trophoblast (EVT) cell line (kindly provided by Professor Charles Graham, Queen’s University, Kingston, Canada). 2. These protocols can be used for other EVT cell lines, similar proliferative and invasive cells, or cancer cell lines. 3. For each cell or cell line used, it is essential to carry out initial proliferation (growth curve) experiments by titrating the number of cells per well to determine the optimum number of cells to use [3].

2.3

Reagents

1. Culture media: RPMI (or as appropriate): complete (10% fetal calf serum) and serum free (SF), each containing L-glutamine (2 mmol/L), penicillin (100 IU/mL) and streptomycin (100 μg/mL). 2. Trypsin/EDTA solution (0.05%). 3. Matrigel™ (BD BioSciences, Bedford, MA USA), for invasion assays.

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4. Phosphate buffered saline, pH 7.4. 5. Bromophenol blue (or other inert dye). 6. Diff-Quick staining kit (Thermo Fisher Scientific, Waltham, MA USA).

3

Methods

3.1 Setting Up the Analyzer

1. Prepare the xCELLigence RTCA-DP instrument in a humidified tissue culture incubator at least 24 h prior to commencement of experiments to allow the instrument to equilibrate to the conditions. 2. Ensure that all electrical leads, including those connecting the instrument to the computer being used for data collection, are securely connected and do not compromise the operation of the incubator. 3. Ensure that the water trays in the incubator are kept full at all times because the xCELLigence system electrodes are highly sensitive to changes in humidity. 4. Carry out all procedures in a cell culture hood unless otherwise indicated.

3.2

Proliferation

1. Carry out experiments monitoring proliferation in single chamber, 16-well E-plate 16 (see Note 2). 2. Experiments can be run using only one plate and a single cradle on the analyser. The maximum number of plates that can be run is three. An experiment can be run as a single plate or can be run across two (32 wells) or three (48 wells) plates. 3. Unwrap the required number of E-plate 16 from their packaging. Wear powder-free gloves while handling and do not touch the electrical components. 4. Remove the plate lids and pipette 100 μL SF medium into each well. Replace the lids. 5. Prepare controls and treatments required for the experiment. Treatments should be prepared at the concentration required to account for the final dilution (i.e., 50 μL treatment +100 μL medium in wells +100 μL cell suspension = 250 μL final volume). 6. Pipette 50 μL SF medium with added vehicle (control) or SF medium with added treatments (test) into the wells according to the desired experimental plate layout (see Fig. 1). 7. Replace the plate lids and move the plates to the incubator while preparing cells.

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Fig. 1 Plate layout for a single plate experiment. A layout for a single plate with positive and negative controls in duplicate and four treatments in triplicate. If running one experiment across multiple plates, include controls on each plate 3.3

Cell Preparation

1. Select a flask of cells approximately 80% confluent. Depending on the experiment being performed, cells should have been serum-starved for at least 18 h prior to commencing. 2. Wash cells twice with PBS then add enough Trypsin/EDTA solution to form a thin cover over the surface of the cells. Place the flask in an incubator and monitor regularly for cell detachment. 3. Resuspend the detached cells in complete medium, transfer to a Falcon tube and centrifuge at 200 × g for 10 min to pellet the cells. 4. Resuspend the cells in a small volume of SF medium and count an aliquot to determine the density. Dilute the cell suspension in SF medium to a final concentration of 1 × 104 cells/100 μL.

3.4 Set Up Experimental Protocol

1. Switch on the computer connected to the xCELLigence RTCA-DP and open the RTCA Pro software. 2. Choose the appropriate plate layout in the DP software (one, two, or three cradles). 3. Select the Exp Notes tab and fill in the details for the experiment including title, aim, and any other relevant information that needs to be recorded. 4. Select the Layout tab and enter the cell and treatment details according to the selected plate layout (see Fig. 1). Indicate any wells that are not being used by turning them off while in this tab. 5. Select the Schedule tab and enter the steps and substeps as outlined in Table 1 (see Notes 3 and 4).

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Table 1 RTCA schedule for PROLIFERATION/ADHESION

3.5 Background Readings

Sweeps

Interval

Unit

Comment

Step 1

1

1

min

Background

Step 2-1

20

2

min

Cells attach

Step 2-2

999

1

h

Step 3

100

15

min

Backup step

1. Take the E-plate 16 that has been equilibrating in the incubator and lower it into the first cradle on the RTCA-DP analyzer. Make sure that the plate is sitting flat and then lock it in place by lowering the cradle handle. Repeat for all plates being used in the experiment and note which cradle each plate is placed in. 2. When the cradle handle was locked in place, the analyzer performs an automatic scan. Select the Message tab and “Plate scanned. Connections OK” should be shown. If not, unlock the cradle handle, check the plate is sitting correctly in the cradle and repeat. 3. Start the background reading by selecting the Play button and data recording will commence. The Play button is the forward arrow button that appears in the ribbon at the top left of the experimental program window. A prompt will appear to save the experiment; select “Yes” to record the background impedance measurement. 4. Once the background reading is completed, “Ready for next step” will show in the bottom left of the program window. Select the Message tab to check if there are any issues with the background reading. 5. Lift the cradle handle, remove the plate, and transfer it to the cell culture hood. Repeat for each plate being used in the experiment.

3.6

Addition of Cells

1. Swirl the cell suspension (prepared in Subheading 3.3) and carefully pipette up and down so that the cells are evenly suspended in solution. 2. Remove the lids from the plates and pipette 100 μL of cell suspension into each well according to the desired experimental plate layout. 3. Replace the plate lids and leave in the cell culture hood at room temperature for 30 min without moving to ensure that the cells distribute evenly across the bottom of the wells and do not clump together.

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3.7 Start Experimental Measurements

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1. Take the E-plate 16 that has been equilibrating in the cell culture hood and place it into the same cradle on the RTCADP analyzer that was used when taking the background reading. Make sure that the plate is sitting flat and lock in place by lowering the cradle handle. Repeat for all plates being used in the experiment. 2. The analyzer will perform another automatic scan. Before clicking the Play button to start experimental measurements, make sure that no connection errors are listed in the Message tab (see Subheading 3.5, step 2). If there is an error, unlock the plate by releasing the cradle handle, remove and then replace the plate in the cradle and lower the handle to lock in place again. Check the Message tab to confirm that this issue is resolved. 3. Click the Play button to commence step 2 of the protocol. The analyzer will now commence the measurements as outlined in the schedule. Subsequent steps will start automatically without any need to click the Play button again. 4. Avoid opening and closing the door of the incubator while experiments are in progress. 5. Allow the experiment to run for 18–24 h, or until the experimental effect is observed up to 72 h (see Note 5). 6. Stop the experiment by clicking the Stop button (circle containing an X) that appears in the ribbon at the top left of the experimental program window.

3.8 Graphing the Data

1. Graph the collected data either while the experiment is in progress or after it has finished. It is helpful to graph and display the data while the experiment is running so that progress can be monitored and the experiment aborted if an obvious problem appears (e.g., no response or no signal). 2. Select the Plot tab and graph cell index versus time by selecting Add All which will graph the data for every well. To graph the data for specific wells, highlight the well of interest and select Add>>. Repeat this sequence for each well that you wish to compare. 3. Select the Average and Draw STDev boxes to show the average and standard deviations respectively (wells must be in at least triplicate to calculate SD). 4. Perform other analysis by selecting the required Curve type under Settings for Data Analysis (see Note 6). 5. Copy and paste data into a Microsoft Excel spreadsheet and save for further analysis. 6. Proliferation can be analysed by looking at the cell index at a single time point or the lag and log phases of cell growth can be determined by plotting cell index versus time.

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Adhesion

1. Experiments monitoring adhesion are carried out with the same single chamber, 16-well E-plate 16. Follow the same protocol used for investigating cell proliferation as described in Subheadings 3.1–3.8. Adhesion data can be collected in the same experiments being conducted to measure proliferation by analyzing data collected during the first few hours of the experiment. 2. Adjust the schedule as needed to collect readings at more frequent intervals during the first few hours of the experiment to examine cell adhesion in more detail (e.g., step 2-1 could be increased to 180 sweeps). 3. If desired, perform separate dedicated adhesion experiments over shorter time frames (see Note 7). In this case, the equilibration step described in Subheading 3.6 can be bypassed. 4. Adhesion can be analysed by looking at the cell index at a single time point or cell index versus time can be plotted to compare the speed of attachment to different substrates or in response to different treatments.

3.10

Migration

1. Carry out experiments monitoring cell migration with uncoated, dual chamber, 16-well CIM-plate 16. The protocol is similar to that outlined for proliferation and adhesion with some modifications as outlined below. 2. Unwrap the required number of CIM-plate 16 bottom chambers and place them in the assembly tool, aligning the blue dots on the tool and the plates (see Fig. 2). Wear powder-free gloves while handling the plates and do not touch the electrical contacts.

Fig. 2 Alignment of CIM-plate 16 in plate assembly tool. The correct positioning of the CIM-plate 16 in the assembly tool with blue dots aligned

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Fig. 3 Meniscus above top edge of bottom chamber wells. A clearly defined convex menisus above the top of the well is essential to prevent air bubbles forming when the top and bottom chambers of the CIM-plate 16 are locked together

3. Pipette 165 μL of SF medium with added vehicle (control) or SF medium with added treatments (test) into the wells of the bottom chamber according to the desired experimental plate layout. Note that treatment concentrations need to be calculated based on the 165 uL volume that is contained in the bottom chamber only. 4. Ensure that a distinct convex meniscus is visible after filling the wells (see Note 8 and Fig. 3). 5. Add 100 μL of sterile water containing a small amount of dye (e.g., bromophenol blue) to all the evaporation chambers that surround the wells (see Note 9). There are 29 chambers in total, two of which are not filled. The dye helps to see which chambers have been filled (see Fig. 4a, b). The two larger chambers at either end of the plate should be filled with 200 μL and the two small chambers at either end of the plate can be left unfilled (see Fig. 4c). 6. Lock the two halves of the CIM-plate 16 together by aligning the blue dots on the top and bottom chambers and pushing down firmly. A distinct click-clack sound will be audible if the chambers have been locked together properly. Repeat for each plate being used in the experiment. 7. Pipette 50 μL of SF medium into each well of the top chambers, being careful to prevent air bubbles forming or touching the bottom of the well. 8. Place the plates in the incubator to equilibrate for 1 h while preparing the cells as outlined in Subheading 3.3. For migration experiments, cells must be prepared at a final concentration of 4 × 104 cells/100 μL.

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Fig. 4 Evaporation chambers filled with water coloured with blue dye. (a) and (b) Addition of an inert dye (e.g., bromophenol blue) to the water used to fill evaporation chambers helps to see which chambers have been filled. c) position of larger (filled) and smaller (not filled) evaporation chambers on a CIM-plate 16 Table 2 RTCA schedule for MIGRATION/INVASION Sweeps

Interval

Unit

Comment

Step 1

1

1

min

Background

Step 2-1

240

1

s

Cells attach

Step 2-2

999

1

h

Step 3

100

15

min

Backup step

9. Set up the experimental protocol as outlined in Subheading 3.4 using the schedule outlined in Table 2. 10. Perform the automatic plate scan and background readings as outlined in Subheading 3.5. 11. Pipette cell suspension into the top chambers of the plates as described in Subheading 3.6. 12. Start the data collection as described in Subheading 3.7. 13. Graph the data as described in Subheading 3.8. 14. Migration can be analyzed by looking at the cell index at a single time point or the rate of migration can be determined from the slope of the line when cell index is plotted against time (see Note 10).

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Invasion

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1. Perform invasion experiments using the migration experiment protocol outlined in Subheading 3.10, with the same cell numbers (4 × 104 per well), and additional steps to prepare the CIM-plate 16 with the desired invasion substrate prior to commencing the experiments (see Note 11). 2. To coat the CIM-plate 16 with Matrigel™, place the required number of plates in the plate assembly tool and sit on ice. Ensure the plates are cold. Keep all equipment, including pipette tips and reagents on ice. 3. Thaw the Matrigel™ on ice and dilute 1:10 (v/v) in ice-cold SF medium ensuring there is sufficient for the number of wells to be coated (50 μL required per well). 4. Take the lids off the CIM-plate 16 top chambers and pipette 50 μL of diluted Matrigel™ solution into the required wells. To ensure that the Matrigel™ forms an even coating, immediately remove 30 μL. Work as quickly as possible to coat all required wells. 5. Place the lids back on the top chambers to prevent evaporation and place the plates in a humidified incubator for a minimum of 30 min but no longer than 2 h to allow the Matrigel™ to set (see Note 12). 6. Analyze the invasion data in the same way as migration data with comparison of cell indices at single time points or determining a rate of invasion from the slope of the line when cell index is plotted against time.

3.12 Examination of Cells Post-experiment

1. For both migration and invasion experiments, stain and examine the underside of the CIM-plate 16 top chamber under a microscope to visualize the cells that have passed through the membrane and attached to the lower surface of the chamber. The Diff-Quick staining kit (Fischer Scientific) is suitable for this process. 2. Carefully remove any medium remaining in the wells of the top chamber of the CIM-plate 16 and then separate the two halves of the plate. 3. Keeping the lower surface of the top chamber upright, stain the membrane following the manufacturer’s directions. Make sure that any excess stain is rinsed off. An example of a stained top chamber is shown in Fig. 5.

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Fig. 5 A stained CIM-plate 16 after completion of a migration assay. Cells that have migrated through the membrane of a CIM-plate 16 top chamber can be visualized by staining and examined under the microscope after the experiment is completed

4 Notes 1. xCELLigence analyzers are available from Agilent Technologies Inc. (Santa Clara, CA USA). There are 9 different models, varying in size and application. 2. Proliferation and adhesion analysis can also be performed using the same methodologies with 96-well plates and the xCELLigence RTCA single plate (SP) or multiple plate (MP) systems. Migration and invasion plates are only available in a 16-well format and thus can only be used with the RTCA-DP system. 3. Step 1 is the background calibration and should never be changed. Entering 999 sweeps allows the experiment to continue indefinitely, enabling as many readings as required to be collected. It also allows the step to be terminated if the data shows that the experiment is completed. Step 3 is a backup should additional data collection be required, with the ability to prematurely stop Step 2.2 and advance to Step 3 which is set up with more frequent sweeps. 4. The timings given here are a guide only. The number and duration of steps and substeps, as well as the interval between measurements, can be adjusted according to the characteristics of the cells under investigation and the function being measured to provide more or less detail at particular time points. Several initial optimization experiments may need to be conducted to determine the ideal protocol.

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5. Longer time frames are not recommended as evaporation of medium can occur which will compromise the experimental recordings. 6. Consult the RTCA software Pro manual for detailed instructions on various ways to plot and analyze the data. 7. E-plate 16 can be coated with matrix components such as laminin, fibronectin, or collagen to investigate adhesion to different substrates. Coating the plates does not interfere with their performance. 8. A defined meniscus is critical to prevent air bubbles forming between the two chambers of the plate when they are locked together. Air bubbles will cause interference and prevent the electrodes from working leading to erroneous data being recorded from the well. 9. Do not overfill the evaporation chambers as this will result in siphoning of water onto the lid and into the top chambers. 10. For a migration experiment to be deemed successful, average cell index must be greater than or equal to 0.2. An ideal range for the cell index is 0.5–1.0 or greater. 11. Substrates for invasion assays include Matrigel™, Geltrex™ (Life Technologies Corporation, Carlsbad, CA USA), MaxGel™ (Merck KGaA, Darmstadt, Germany), or fibronectin. In each case, the appropriate concentration to use will need to be determined by titration experiments. 12. Do not leave for more than 2 h or the coating may dry out. Alternatively, covered plates can be left in a cell culture hood at room temperature overnight. The top chamber can be placed loosely onto the empty bottom chamber in order to keep both parts of the plate sterile. References 1. Chau SE, Murthi P, Wong MH et al (2013) Control of extravillous trophoblast function by the eotaxins CCL11, CCL24 and CCL26. Hum Reprod 28(6):1497–1507 2. Yong HE, Murthi P, Wong MH et al (2015) Effects of normal and high circulating concentrations of activin a on vascular endothelial cell functions and vasoactive factor production. Pregnancy Hypertens 5(4):346–353

3. Keogh RJ (2010) New technology for investigating trophoblast function. Placenta 31(4): 347–350

Part III Targeted Therapy

Chapter 14 Liposome-Encapsulated Anti-inflammatory Proteins for Targeted Delivery to the Placenta to Treat Fetal Growth Restriction Padma Murthi and Lynda K. Harris Abstract Fetal growth restriction (FGR), the failure of a fetus to reach its genetically determined growth potential, is a serious complication affecting up to 10% of pregnancies. FGR is a major risk factor for stillbirth and, in the survivors, neurodevelopmental disorders. We have recently identified that the anti-inflammatory and pro-resolving molecule, lipoxin A4 (LXA4) and its soluble receptor, formyl-peptide receptor-2 (FPR-2) are significantly decreased in human placentas from FGR pregnancy. The LXA4 synthetic analog Compound 43 (C43) is considered a safe, anti-inflammatory therapy and is being developed as a treatment for disease conditions with an inflammatory basis, for example, asthma in children. Identification of therapies to treat FGR in utero comes with the need to mitigate their potential side effects and the use of nanoparticlemediated delivery systems could facilitate this. Our current studies are focused on targeting the resolution of inflammation observed in FGR placentas, by synthesizing liposome-encapsulated C43 as a novel therapeutic to improve placental function in FGR. In this chapter, we provide a detailed methodological procedure for the preparation of liposomes and conjugation of the peptide sequences, which selectively bind to the outer placental syncytiotrophoblast layer or the vascular endothelium of the uterine spiral arterioles. Key words Nanoparticles, syncytiotrophoblast, liposome-mediated therapy, fetal growth restriction, placental inflammation

1

Introduction Fetal growth restriction (FGR) is a significant pregnancy complication, where fetuses fail to reach their genetically determined growth potential in utero. FGR affects up to 10% of all pregnancies and contributes significantly to stillbirth and iatrogenic preterm birth. As well as the acute consequences of being born too small, affected offspring are at increased risk of cardiovascular disease, diabetes, and other chronic diseases in later life [1–4]. FGR has a number of different etiologies, but the most common cause is placental insufficiency, which arises from inadequate trophoblast invasion and

Sandeep Raha (ed.), Trophoblasts: Methods and Protocols, Methods in Molecular Biology, vol. 2728, https://doi.org/10.1007/978-1-0716-3495-0_14, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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incomplete maternal uterine spiral artery remodeling. Placental histopathological features from FGR pregnancies include impaired development of the uteroplacental vasculature [5–7], elevated apoptosis [8, 9], and reduced nutrient transport activity [10, 11] in the syncytiotrophoblast. FGR is also associated with utero-placental hypoxia, inflammation, and impaired vascular tone regulation, which are observed later in pregnancy [12]. Recent studies from our laboratory have identified exacerbated placental inflammation as one of the molecular mechanisms that co-exists with the utero-placental insufficiency and abnormal trophoblast function observed in FGR pregnancies. Specifically, our studies have reported that expression of the anti-inflammatory and pro-resolving molecule, lipoxin A4 (LXA4) and its soluble receptor, formyl-peptide receptor-2 (FPR-2) are significantly decreased in human placentas from FGR/SGA pregnancies, and this contributes to abnormal vascular functions, nutrient transport, and endocrine functions [13, 14]. However, the effect of LXA4 as a novel therapy to treat inflammation and improve placental function in FGR is yet to be investigated. Many therapies for pregnancy disorders focus on specifically enhancing uteroplacental perfusion and fetal growth in humans and animal models by systemically administering sildenafil citrate or melatonin, for example; however, detrimental maternal and fetal side effects have been reported [15, 16]. Furthermore, the systemic administration of these compounds can affect normal physiology in other organs, resulting in off-target effects. Rather than directly targeting specific pathways, the use of pro-resolution molecules such as LXA4 and its synthetic analog Compound 43 (C43) as a therapy may resolve the pathological signaling, acting as a protective mechanism to dampen the excessive inflammatory processes that contribute to placental dysfunction in FGR. We have identified the synthetic agonist of FPR-2, C43 as a novel therapeutic for treating placental dysfunction in FGR. LXA4/C43 therapy is considered safe and is being developed as a treatment for other disease conditions with an inflammatory basis such as asthma in children [17]. One of the major concerns in developing and testing therapeutics by pharmaceutical industry for human pregnancies is that maternally administered drugs may have detrimental effects on the developing fetus. To overcome this, Dr. Harris has developed a method of peptide-mediated targeting of payloads to the uterus and placenta using nanoparticle courier systems, with enhanced drug effectiveness and reduced off-target, systemic side effects [18, 19]. Given these promising findings, our current studies are focused on targeting the resolution of inflammation in the human placenta using liposome-encapsulated C43 as a novel therapeutic strategy to improve placental function in FGR.

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Materials 1. 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC). 2. 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] ammonium salt [DSPE-PEG (200)]. 3. 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[maleimide(polyethylene glycol)-2000] ammonium salt DSPE-PEG (2000)-maleimide (Avanti Polar Lipids)]. 4. Methanol and chloroform. 5. Cholesterol. 6. Rotary evaporator and round bottom flasks. 7. Vacuum oven. 8. 1 mL Liposome Mini-Extruder (Avanti Polar Lipids). 9. Slide-a-Lyser dialysis cassettes (MW 10 kDa, and 3.5 kDa cut off). 10. Culture medium: RPMI 1640 containing 10% heat-inactivated FCS (fetal calf serum), 2 mM glutamine, 100 IU/mL penicillin, and 100 μg/mL streptomycin. 11. Culture medium for freshly isolated trophoblasts: Endothelial growth medium (EGM-2) containing EBM™-2 Basal Medium (Catalog No. CC-3156, LONZA) and EGM™-2 SingleQuots™ Supplements, which includes growth factors and antibiotics (Catalog No. CC-4176, LONZA). 12. Culture medium for placental endothelial cells: MTT (3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide) (Catalog No. 11465007001, Sigma-Aldrich), and phosphate-buffered saline (PBS).

3

Methods Peptide-decorated liposomes are prepared using the thin lipid film method and were composed of 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC; 32.5 mM), 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] ammonium salt [DSPE-PEG(200); 1.875 mM], 1,2-distearoylsn-glycero-3-phosphoethanolamine-N-[maleimide(polyethylene glycol)-2000] ammonium salt [DSPE-PEG(2000)-maleimide; 0.625 mM; Avanti Polar Lipids], and 15 mM cholesterol, as previously described [18, 19]. This amount of lipid will produce 1 mL of liposome suspension.

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3.1 Synthesis of the Liposomes and Encapsulation of C43

1. Weigh out all lipids and transfer them directly into a clean and dry glass round-bottomed flask. 2. Fill up the water bath of the rotary evaporator and set to warm to 40 °C. 3. In a fume hood, add approximately 10 mL of methanol:chloroform (9:1 v/v) to the round-bottomed flask and dissolve the lipids by gentle swirling (see Note 1). 4. Connect the flask to the rotary evaporator and run at 270 mbar until all of the solvent has been removed, which takes approximately 5–10 min. A thin lipid film will remain in the bottom of the flask. 5. Dry the lipid film overnight in a vacuum oven (0 mbar, room temperature). 6. To prepare the C43 stock solution for rehydration of the lipid film, add 1 mL of 10 mM HCl to the C43 solution. Transfer 300 μL of this C43/HCl solution to a 7 mL bijou containing 700 μL sterile PBS to produce a final concentration of 320 μmol/L. After mixing, adjust pH of the stock solution to 7.4 (see Note 2). 7. To rehydrate the lipid film, add 1 mL of the desired cargo. This method of passive loading leads to approximately 50% of the cargo being encapsulated inside the liposomes. Thus, if a final concentration of 1 mM cargo is required, rehydrate the lipid film with a 2 mM solution of cargo in PBS. To encapsulate C43, rehydrate the lipid film with the 320 μmol/L stock prepared above (see Note 3). 8. After rehydration, cover the flask tightly with foil to prevent evaporation. Vortex the flask for 5 min to disrupt the lipid film. 9. Transfer the flask to a warming oven set to 45 °C for 15 min, vortex the flask again for 5 min, then return the flask to the oven. 10. Repeat this process for 4 h, vortexing the lipid suspension for 5 min every 30 min, before returning the flask to the oven. This process will produce a suspension of large, multi-lamellar liposomes. 11. Ensure the syringes from a 1 mL Liposome Mini-Extruder (Avanti Polar Lipids) have been thoroughly cleaned with 100% alcohol and dried at room temperature. 12. Pre-warm the glass syringes, heating block and extruder components to 45 °C in the oven for at least 1 h (see Note 4). 13. Pre-warm 15 mL of sterile PBS to 45 °C. Place two 10 mm filter supports and one 19 mm polycarbonate extrusion membrane (200 nm pore size) into a petri dish containing ~10 mL

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of warm, sterile PBS to hydrate and warm for approximately 10 min. 14. Place the filter and filter supports in the extruder, ensuring all components are screwed tightly together. 1 mL of the remaining PBS can be passed through the extruder to check for leaks, but the syringes must be emptied prior to extrusion of the liposome suspension (see Note 5). 15. Fill one syringe with the liposome suspension. Extrude the formulation by passing it from one syringe into the other, through the membrane, 11 times. The last pass should fill the opposite syringe from the one you started with (see Note 6). 16. If the liposomes are to be used in cell or tissue culture, or animal experiments, maintain sterility. Transfer the extruded liposome suspension to a sterile Eppendorf tube and allow to cool to room temperature for 15 min. 17. If more than one liposome suspension needs to be extruded, clean and dry the syringes. Insert a replacement filter and filter supports into the Mini Extruder. All components must be warmed to 45 °C before the next extrusion is performed. 18. The external surface of the liposomes is decorated with the targeting peptide rhodamine-CCGKRK (which selectively binds to the placental syncytiotrophoblast and the uterine spiral arterioles) via a Michael-type addition reaction, whereby the N-terminal cysteine residue is covalently coupled to maleimide groups in the lipid bilayer. 19. Prepare a stock solution of rhodamine-CCGKRK (1.25 mM in sterile PBS) and add 100–900 μL of liposomes, resulting in a final concentration of 1.25 μM. Incubate the liposomes overnight at room temperature to allow peptide conjugation to take place. Wrap the Eppendorf in aluminum foil to protect the rhodamine fluorophore from exposure to light. 20. Wash all Mini Extruder components in warm tap water, followed by dH2O. Wash the syringes with tap water, dH2O, and then acetone to remove any lipid residue and aid drying (see Note 7). 21. Ensure an adequate volume of sterile PBS and a 1 L beaker are autoclaved for dialysis on day 2 of the procedure. 22. After overnight peptide conjugation, remove unconjugated peptide from the liposome suspension by dialysis against sterile PBS (8 × 1 L; 24 h) using Slide-A-Lyzer Dialysis Cassettes with a molecular weight cut off (MWCO) of 10 kDa. The size of the encapsulated cargo will determine the MWCO of choice; the pore size must be large enough to let any unencapsulated cargo exit the dialysis cassette. For empty liposomes, a MWCO of 3.5 kDa is sufficient (see Note 8).

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23. Carefully introduce the liposome suspension into the dialysis cassette using a needle and a 1 mL syringe. Take extreme care not to puncture or tear the dialysis membrane with the needle tip (see Note 9). 24. Fill an autoclaved glass beaker with approximately 1 L of sterile PBS. Place the beaker on a stirring plate and add a magnetic stirrer. Turn on the stirring plate to create a stable vortex of PBS (see Note 10). 25. Cover the beaker with aluminum foil to protect the liposomes from light and allow the cassette to dialyze at 4 °C. 26. Change the PBS every hour for 3–4 h, then leave to dialyze overnight. Aim for 8 changes of PBS within the 24 h dialysis period. 27. Use a clean, sterile needle and syringe to carefully remove the liposomes from the dialysis cassette and transfer them to a sterile Eppendorf tube. Wrap the tube in aluminum foil to protect the liposomes from light and store it at 4 °C. 28. When stored at 4 °C and kept sterile, the liposomes will remain stable for a minimum of 4 weeks. 3.2 Determine the Effect of LiposomeEncapsulated C43 on Placental Trophoblast or Endothelial Cell Viability and Proliferation In Vitro

Principle of the MTT Assay The MTT assay is used to measure cellular metabolic activity as an indicator of cell viability, proliferation, and cytotoxicity. This colorimetric assay is based on the reduction of a yellow tetrazolium salt MTT to purple formazan crystals by metabolically active cells. The viable cells contain NAD(P)Hdependent oxidoreductase enzymes which reduce the MTT to formazan. 1. Seed human placental trophoblasts or endothelial cells into microplates (tissue culture grade, 96 wells, flat bottom).at a concentration of 1 × 105 cells/well in 100 μL culture medium and allow the cells to adhere for 4 h at 37 °C and 5% CO2. 2. Replace culture medium and add fresh medium containing free or liposome-encapsulated C43 [final concentration e.g., 5–50 μM]. 3. Incubate treated cell cultures for 24 h at 37 °C and 5% CO2. 4. After the incubation period, remove the culture medium and add fresh media containing 10 μL of the MTT reagent (final concentration 0.5 mg/mL) to each well. 5. Incubate the microplate for 4 h in a humidified atmosphere at 37 °C and 5% CO2. 6. Add 100 μL of the solubilization solution into each well. 7. Allow the plate to stand overnight in the incubator at 37 °C and 5% CO2.

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8. Check for complete solubilization of the purple formazan crystals and measure the absorbance of the samples using a spectrophotometer/microplate (ELISA) reader. The wavelength to measure absorbance of the formazan product is between 550 and 600 nm according to the filters available for the ELISA reader. The reference wavelength should be more than 650 nm (see Note 11).

4

Notes 1. Ensure that no lipid remains undissolved in the neck of the flask. 2. This is the minimum volume that can be accurately measured by a pH meter; it is easier to prepare a larger volume and freeze the remaining agonist. 3. It is good practice to include a control liposome formulation in any experiment, to assess any effects of the liposome constituents, in the absence of the cargo. Thus, a second lipid film can be prepared in parallel, and hydrated with PBS or the vehicle in which the encapsulated cargo has been dissolved. 4. Do not heat syringes above 80 °C and do not let them undergo rapid temperature changes. 5. Do not twist syringes once they are in place within the extruder, as they come unscrewed and will leak. 6. The extrusion process will produce a sterile, unilamellar liposome suspension with an average diameter of 200 nm, free from any remnants or contaminants >200 nm found in the original suspension. 7. Do not wash the extruder components with detergent (e.g., Decon) or acetone as they damage the washers. Do not wash the syringes with detergent. 8. Dialysis cassettes should be wetted by immersion in PBS for 10 min prior to use. 9. Do not introduce any excess air; this can be removed again with the empty needle and syringe. Also, be aware of the maximum volume capacity of the cassette; two or more cassettes may be needed to dialyze larger volumes of liposomes. 10. Add the cassette to the beaker; the cassette should float, but not hit sides or bottom of the beaker. 11. The darker the solution, the greater the number of viable, metabolically active cells.

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References 1. Malhotra A et al (2019) Neonatal morbidities of fetal growth restriction: pathophysiology and impact. Front Endocrinol (Lausanne) 10: 55 2. Thornburg KL et al (2008) The role of growth in heart development. Nestle Nutr Workshop Ser Pediatr Program 61:39–51 3. Barker DJ (2006) Adult consequences of fetal growth restriction. Clin Obstet Gynecol 49(2): 270–283 4. Baschat AA, Odibo AO (2011) Timing of delivery in fetal growth restriction and childhood development: some uncertainties remain. Am J Obstet Gynecol 204(1):2–3 5. Macara L et al (1996) Structural analysis of placental terminal villi from growth-restricted pregnancies with abnormal umbilical artery Doppler waveforms. Placenta 17(1):37–48 6. Ptacek I et al (2016) Quantitative assessment of placental morphology may identify specific causes of stillbirth. BMC Clin Pathol 16:1 7. Junaid TO et al (2014) Fetoplacental vascular alterations associated with fetal growth restriction. Placenta 35(10):808–815 8. Levy R et al (2002) Trophoblast apoptosis from pregnancies complicated by fetal growth restriction is associated with enhanced p53 expression. Am J Obstet Gynecol 186(5): 1056–1061 9. Heazell AE et al (2011) Intra-uterine growth restriction is associated with increased apoptosis and altered expression of proteins in the p53 pathway in villous trophoblast. Apoptosis 16(2):135–144 10. Glazier JD et al (1997) Association between the activity of the system A amino acid transporter in the microvillous plasma membrane of the human placenta and severity of fetal compromise in intrauterine growth restriction. Pediatr Res 42(4):514–519

11. Johansson M et al (2002) Activity and protein expression of the Na+/H+ exchanger is reduced in syncytiotrophoblast microvillous plasma membranes isolated from preterm intrauterine growth restriction pregnancies. J Clin Endocrinol Metab 87(12):5686–5694 12. Baker BC et al (2021) Hypoxia and oxidative stress induce sterile placental inflammation in vitro. Sci Rep 11(1):7281 13. Lappas M et al (2018) Formyl peptide receptor-2 is decreased in foetal growth restriction and contributes to placental dysfunction. Mol Hum Reprod 24(2):94–109 14. Murthi P et al (2020) Decreased placental FPR2 in early pregnancies that later developed small-for-gestation age: a potential role of FPR2 in the regulation of epithelialmesenchymal transition. Cells (4):9, 921 15. Inocencio IM et al (2019) Effects of maternal sildenafil treatment on vascular function in growth-restricted fetal sheep. Arterioscler Thromb Vasc Biol 39(4):731–740 16. Gonzalez-Candia A et al (2016) Potential adverse effects of antenatal melatonin as a treatment for intrauterine growth restriction: findings in pregnant sheep. Am J Obstet Gynecol 215(2):245 e241–245 e247 17. Kong X et al (2017) Pilot application of lipoxin A4 analog and lipoxin A4 receptor agonist in asthmatic children with acute episodes. Exp Ther Med 14(3):2284–2290 18. Harris LK (2016) Could peptide-decorated nanoparticles provide an improved approach for treating pregnancy complications? Nanomedicine (Lond) 11(17):2235–2238 19. King A et al (2016) Tumor-homing peptides as tools for targeted delivery of payloads to the placenta. Sci Adv 2(5):e1600349

Chapter 15 Trophoblast-Targeted Liposomes for Placenta-Specific Drug Delivery Baozhen Zhang, Xiujun Fan, and Nihar R. Nayak Abstract A major challenge in developing potential treatments for pregnancy complications is minimizing adverse effects to the fetus and mother. Placenta-targeted drug delivery could reduce the risks of drug treatments in pregnancy by targeting tissue where most pregnancy complications originate and decreasing dosages. We previously developed a tool for the targeted delivery of drug-carrying nanoparticles to the placenta using a synthetic placental chondroitin sulfate A-binding peptide (plCSA-BP) derived from the malarial protein VAR2CSA, which binds a distinct type of chondroitin sulfate A (CSA) exclusively expressed by placental trophoblasts. Liposomes are a type of nanoparticle already approved for use in humans by the Food and Drug Administration (FDA) and used successfully for the treatment of a wide range of diseases. Here, we present a detailed method to create plCSA-BP-decorated liposomes that can be used to deliver drugs specifically to placental trophoblasts. Liposomes are first generated by the standard film method and then conjugated to plCSA-BPs using the 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride/Nhydroxysulfosuccinimide (EDC/NHS) bioconjugate technique. This protocol may facilitate bench-tobedside translation of drug discovery for the treatment of pregnancy disorders by reducing risks of side effects, and enabling rapid and scalable production. Key words Trophoblast, Placental chondroitin sulfate A, Placental chondroitin sulfate A binding peptide, Liposome, EDC/NHS

1

Introduction Pregnancy disorders are common and can have life-long consequences for both the mother and fetus but treatments that address the underlying pathophysiology of these disorders are lacking. One of the many barriers impeding the development of drug therapies for these disorders is the risk of side effects for the fetus during critical developmental stages. The most common and severe pregnancy disorders, such as pre-eclampsia, are thought to originate in the placenta, and strategies that interrupt pathogenetic mechanisms arising in the placenta have the potential to reduce serious pregnancy risks. Drug delivery methods that specifically target the

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placenta could minimize adverse effects of drugs on the fetus and mother by reducing systemic drug loads and exposure to other potentially sensitive tissues. In the last decade, investigators have developed two general strategies for targeting drug delivery to the placenta: one strategy directly targets the placenta using molecules that selectively bind to molecules enriched on placental membranes, and one strategy targets the uterus via uterus-enriched molecules and then relies on controlled release from nanoparticles to indirectly deliver drugs to the placenta. Molecules that have been used for direct targeting to the placenta include the following: GKRK/CCGKRK and iRGD, originally identified as tumor-homing peptides and subsequently shown to selectively target the placenta [1, 2]; anti-epidermal growth factor receptor (EGFR) antibodies [3]; and peptides selected in phage-display screens for binding to the placental vasculature [4]. EGFR levels are low in most tissues, but very high in trophoblasts. Both the tumor-homing peptides later shown to target the placenta and the placental vasculature-targeted peptides found through molecular screens have been shown to bind selectively to the endothelium of spiral arteries and remodel these arteries through targeted drug delivery. Nanoparticles targeting the uterus have been generated by decoration with oxytocin receptor or oxytocin antagonists, as oxytocin is highly expressed in the uterus during pregnancy [5]. We developed a method that is based on a naturally occurring placental targeting process associated with human infection of the malarial parasite, P. falciparum. P. falciparum-infected erythrocytes bind to placental chondroitin sulfate A (plCSA), which is exclusively present on trophoblasts, through the malarial VAR2CSA protein [6, 7]. In our early work, we demonstrated that a synthetic plCSA-binding peptide (plCSA-BP), which is derived from the malarial VAR2CSA sequence, specifically binds to human syncytiotrophoblasts and mouse placental labyrinth [8]. We synthesized plCSA-NPs (pICSA-nanoparticles) by conjugating plCSA-BP with lipid-polymer nanoparticles. The resulting plCSA-NPs exhibited good monodispersity, high drug-loading capability, sustained drug release, and excellent in vivo stability. More importantly, plCSA-NPs efficiently delivered drugs to trophoblasts, did not detectably cross the placenta from mother to fetus, and did not cause any apparent fetal toxicity [8, 9]. Liposomes are composed of natural phospholipids and cholesterol and have an internal aqueous core [10] that makes them suitable for the delivery of both hydrophilic and hydrophobic drugs [11, 12]. Liposomes are fully biodegradable, highly biocompatible, and safe. Thus, liposomes are the first nanotherapeutics to be approved by the Food and Drug Administration (FDA) and used for treatment of a wide range of diseases [13, 14]. In this protocol, we have focused on the preparation of plCSA-NPs. First,

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Fig. 1 The schematic depicting synthesis of plCSA-liposomes. Liposomes are synthesized by the original film method and then plCSA-liposomes are generated using the EDC/NHS bioconjugation technique

drug-loaded liposomes are synthesized by the film method. Then, plCSA-BP is conjugated to nanoparticles using a common bioconjugate technique (Fig. 1) that employs 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC), which is a popular crosslinking agent for conjugating peptides containing amines and carboxylates [15]. The water-soluble EDC reacts with carboxylates to form an active ester intermediate. The ester intermediate is highly reactive with amine molecules and forms stable amido bonds. N-hydroxysulfosuccinimide (NHS) is the most common conjugation reagent in surface and nanoparticle conjugation reactions [16, 17]. NHS can reduce the number of side reactions, and enhance the stability and yield of ester intermediates [18]. Several excellent books on bioconjugate techniques have been published over the last decade [19–21]. Readers are encouraged to consult these works to obtain more information on other methods of conjugating peptides to nanoparticles that may be most appropriate for their studies.

2

Materials 1. plCSA binding peptide (plCSA-BP, EDVKDINFDTKEK FLAGCLIVSFHEGKC) is commercially available from several peptide synthesis companies and dissolved in 10–20% acetonitrile. plCSA-BP contains a cysteine (Cys) residue to allow its covalent coupling to the liposomal surface (see Note 1). 2. Phospholipids and cholesterol: Distearoylphosphatidylcholine (DSPC), cholesterol and 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-maleimide (polyethyleneglycol 2000) carboxylic acid (DSPE-PEG-COOH) are available from Avanti

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Polar Lipids. All lipids are dissolved in suitable chloroformmethanol (2:1, v/v) and are stable for several months at 20 °C. 3. Hydration buffer: Store deionized water in a glass apparatus. Phosphate-buffered saline (PBS) and Tris-buffered saline are available commercially and are stored at 4 °C. 4. Organic solvents: Acetonitrile, chloroform, and methanol should be analytical grade and are available commercially. Since chloroform can deteriorate on storage over more than 1–3 months, it should be redistilled before use or used fresh. 5. Conjugated reagents: 2-(morpholino) ethanesulfonic acid (MES), EDC, and NHS are available from Sigma. Prepare a 0.1 M MES stock solution by dissolving 2.17 g of MES in 100 mL of deionized water, and store at 4 °C. 6. Glass beads used for distillation are commercially available. Round-bottom flasks, Stirred-cell ultrafiltration units, and extrusion devices are available from T&T Scientific Corporation (Knoxville, TN, USA) (see Notes 2 and 3). 7. plCSA-liposomes characterization devices: The Malvern Zetasizer Nano ZS is available from Malvern Panalytical. 8. Bicinchoninic acid (BCA) protein assay kit and plate reader. 9. 20× PBS (provide the concentration of the constituent components for 20×).

3

Methods

3.1 Film Method to Synthesize Liposomes

1. This method has been previously reported [21, 22]. Prepare the desired molar ratio of DSPC cholesterol and DSPE-PEGCOOH (2:1:3) in 10 mL of chloroform-methanol (2:1, v/v), and add the mixture to a round-bottom flask. 2. Remove the solvent by rotary evaporation at approximately 40 °C in a water bath under reduced pressure (below 0.1 Pa) so that a thin film of lipids is deposited on the inner wall of the flask. 3. Add 100 mL of 0.5 mm glass beads to the flask containing the dried lipid mixture, then add an appropriate volume of hydration buffer (such as deionized water, PBS, or Tris-buffered saline) to the flask so that the total lipid per milliliter of buffer does not exceed 2% by weight (see Note 4). 4. Vortex mix the flask for 15 min or longer so that the lipid film is dispersed into the hydration buffer.

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5. Add the hydrated lipids and drugs into an extrusion device with a desired of diameter filter 21 times to obtain drug-loaded liposomes. (see Note 5). 6. Purify the liposomes by washing in hydration buffer using stirred-cell ultrafiltration and lyophilize. 3.2 Conjugation of Peptides to Liposomes

1. Add the desired molar ratio of liposomes to 0.1 M MES buffer, pH 6.0 (see Note 6).

3.2.1

Ester Activation

2. Add a ten-fold molar excess of EDC and NHS (final concentration ratio 2:5) to the above buffer and react for 0.5–1 h at room temperature (see Note 7).

3.2.2

Amine Reaction

1. Increase the buffer pH to 7.2–7.5 by using 20× PBS, pH 7.4. 2. Add plCSA-BPs (10× molar ratio of conjugated carboxyl) to the activated liposomes. Place the reaction components on a shaker at 4 °C, and allow the reaction to proceed overnight (see Note 8). 3. Purify the plCSA-liposomes as described in Subheading 3.1, step 6 and lyophilize.

3.3 Characterization of the plCSAliposomes

Some of the important parameters for characterizing the plCSAliposomes are as follows: 1. The size of the plCSA-liposomes. 2. Encapsulation efficiency and loading efficiency. 3. Conjugation efficiency. Each is discussed in detail below.

3.3.1 Determination of the Size of plCSAliposomes

1. Dilute plCSA-liposomes with deionized water. Load the desired volume of the liposomes into a cuvette.

3.3.2 Determination of the Encapsulation Efficiency and Loading Efficiency

1. The encapsulation efficiency and drug loading efficiency are determined by reference to the standard curve with ultraviolet (UV) spectroscopy using a known amount of drug. Prepare and measure all solutions in triplicate to minimize error.

2. The size, polydispersity index (PDI), and zeta potential can be determined using a Malvern Zetasizer Nano ZS with dynamic light scattering (see Note 9).

2. Calculate the encapsulation efficiency using the following equation: Encapsulation efficiency = ððAmount of drugs in liposomesÞ=ðAmount of added drugÞÞ × 100%

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Calculate the loading efficiency using the following equation: Loading efficiency = ððAmount of drugs in liposomesÞ=ðTotal weight of materialsÞÞ × 100% 3.3.3 Determination of the Conjugation Efficiency

Measurement of the plCSA-BP conjugation efficiency using the BCA assay [23, 24] 1. Measure the absorbance at 562 nm on a plate reader. Use the generated standard curve to calculate the amount of plCSA-BP. Prepare and measure all solutions in triplicate to minimize errors. 2. Calculate the plCSA-BP conjugation efficiency using the following equation: Conjugation efficiency = ððAmount of peptide in liposomesÞ=ðAmount of added peptideÞÞ × 100%

4

Notes 1. If plCSA-BP has low solubility, sonication in a bath sonicator may be helpful. 2. Stirred-cell ultrafiltration is used to synthesize large-volume (>50 mL) liposomes and the membrane diameter or molecular weight cut-off limit (NMWL) should be larger than those of the drugs. If synthesizing small-volume liposomes, an ultrafiltration centrifuge tube is preferred. 3. In this protocol, the extrusion device is used to synthesize large-volume liposomes. If synthesizing small-volume liposomes, the Avanti mini extruder is a suitable alternative. 4. When a buffer is chosen as the medium for dispersal of dried lipids, facts about subsequent experiments should be borne in mind. For example, if some experiments involve calcium, then PBS is to be scrupulously avoided. Calcium phosphate precipitation would interfere with the absorbance at 330 nm. 5. The diameter of filters for determining the sizes of liposomes. Smaller liposomes are known to have long-circulating character, lower polydispersity, and more stability, all of which consequently decrease the drug loading efficiency [25]. For the extrusion method to synthesize the liposomes, liposome diameter should be larger than the filter diameter. For example, a filter with a pore size of 100 nm allows the formation of

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liposomes with diameters of 110–120 nm. If smaller liposomes are desired, continued filtering through 80 nm pores is needed. 6. The pH of the buffer is important. The optimal pH of ester activation is 5.0–6.0 and the amine reaction is most efficient at pH 7.2–7.5. For best results, the reaction should be performed in two steps. First, ester activation in MES (or other non-carboxylate buffer) at pH 5.0–6.0, then raise the pH to 7.2–7.5 with PBS (or other non-amine buffer). 7. For easy addition of the correct quantity of EDC and NHS, a stock solution may be prepared and used immediately. 8. Considering that the half-life of NHS esters is 4–5 h at pH 7.0, the duration of the amine reaction exceeds 10 h at 4 °C. Raising the temperature increases the reaction rate but also increases non-covalent conjugation. The reaction can be performed for 2 h at room temperature. 9. PDI is a measure of the heterogeneity of a sample based on size. International standards organizations have established that PDI values 0.7 are common to polydisperse distributions of particles [26]. Zeta potential is the nature of the electrostatic potential near the surface of a particle; if it is less than -15 mV, it usually represents the onset of agglomeration. The higher the absolute zeta potential, the more stable the particle. Zeta potentials of >30 mV are considered optimal for good stabilization of nanodispersion [27].

Acknowledgments The work was supported by grants from the National Natural Science Foundation of China (82302374), March of Dimes Birth Defects Foundation and the Preeclampsia Foundation to NRN at Stanford University and WSU. NRN is supported by the NICHD grant #R01HD088549 at WSU and UMKC (the content is solely the responsibility of the authors and does not necessarily represent the official views of the US National Institutes of Health). References 1. King A, Ndifon C, Lui S et al (2016) Tumorhoming peptides as tools for targeted delivery of payloads to the placenta. Sci Adv 2: e1600349 2. Beards F, Jones LE, Charnock J et al (2017) Placental homing peptide-microRNA inhibitor conjugates for targeted enhancement of intrinsic placental growth signaling. Theranostics 7: 2940–2955

3. Kaitu’u-Lino TJ, Pattison S, Ye L et al (2013) Targeted nanoparticle delivery of doxorubicin into placental tissues to treat ectopic pregnancies. Endocrinology 154:911–919 4. Cureton N, Korotkova I, Baker B et al (2017) Selective targeting of a novel vasodilator to the uterine vasculature to treat impaired Uteroplacental perfusion in pregnancy. Theranostics 7: 3715–3731

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5. Wathes DC, Borwick SC, Timmons PM et al (1999) Oxytocin receptor expression in human term and preterm gestational tissues prior to and following the onset of labour. J Endocrinol 161:143–151 6. Salanti A, Staalsoe T, Lavstsen T et al (2003) Selective upregulation of a single distinctly structured var gene in chondroitin sulphate A-adhering Plasmodium falciparum involved in pregnancy-associated malaria. Mol Microbiol 49:179–191 7. Salanti A, Clausen TM, Agerbæk M et al (2015) Targeting human cancer by a glycosaminoglycan binding malaria protein. Cancer Cell 28:500–514 8. Zhang B, Tan L, Yu Y et al (2018) Placentaspecific drug delivery by trophoblast-targeted nanoparticles in mice. Theranostics 8:2765– 2781 9. Zhang B, Cheng G, Zheng M et al (2018) Targeted delivery of doxorubicin by CSA-binding nanoparticles for choriocarcinoma treatment. Drug Deliv 25:461–471 10. Bozzuto G, Molinari A (2015) Liposomes as nanomedical devices. Int J Nanomedicine 10: 975–999 11. Alavi M, Karimi N, Safaei M (2017) Application of various types of liposomes in drug delivery systems. Adv Pharm Bull 7:3–9 12. Sercombe L, Veerati T, Moheimani F et al (2015) Advances and challenges of liposome assisted drug delivery. Front Pharmacol 6:286 13. Gebre MS, Brito LA, Tostanoski LH et al (2021) Novel approaches for vaccine development. Cell 184:1589–1603 14. Allen TM, Cullis PR (2013) Liposomal drug delivery systems: from concept to clinical applications. Adv Drug Deliv Rev 65:36–48 15. Totaro KA, Liao X, Bhattacharya K et al (2016) Systematic investigation of EDC/sNHSmediated bioconjugation reactions for carboxylated peptide substrates. Bioconjug Chem 27:994–1004 16. Nair M, Johal RK, Hamaia SW et al (2020) Tunable bioactivity and mechanics of collagen-based tissue engineering constructs: a comparison of EDC-NHS, genipin and TG2 crosslinkers. Biomaterials 254:120109

17. Surendran SP, Thomas RG, Moon MJ et al (2020) A bilirubin-conjugated chitosan nanotheranostics system as a platform for reactive oxygen species stimuli-responsive hepatic fibrosis therapy. Acta Biomater 116:356–367 18. Liu EY, Jung S, Yi H (2016) Improved protein conjugation with uniform, macroporous poly (acrylamide-co-acrylic acid) hydrogel microspheres via EDC/NHS chemistry. Langmuir 32:11043–11054 19. Hermanson GT (2013) Bioconjugate techniques. Academic press 20. Pabst G, Kucˇerka N, Nieh M-P et al (2014) Liposomes, lipid bilayers and model membranes: from basic research to application. CRC Press 21. Basu SC, Basu M (2002) Liposome methods and protocols. Springer Science & Business Media 22. Patil YP, Jadhav SJC, Lipids PO (2014) Novel methods for liposome preparation. Chem Phys Lipids 177:8–18 23. Altintas I, Heukers R, Van Der Meel R et al (2013) Nanobody-albumin nanoparticles (NANAPs) for the delivery of a multikinase inhibitor 17864 to EGFR overexpressing tumor cells. J Control Release 165:110–118 24. Chen H, Gao J, Lu Y et al (2008) Preparation and characterization of PE38KDEL-loaded anti-HER2 nanoparticles for targeted cancer therapy. J Control Release 128:209–216 25. Zhang L, Chan JM, Gu FX et al (2008) Selfassembled lipid- polymer hybrid nanoparticles: a robust drug delivery platform. ACS Nano 2:1696–1702 26. Rane SS, Choi P (2005) Polydispersity index: how accurately does it measure the breadth of the molecular weight distribution? Chem Mater 17:926–926 27. Samimi S, Maghsoudnia N, Eftekhari RB et al (2019) Chapter 3 – Lipid-based nanoparticles for drug delivery systems. In: Mohapatra SS, Ranjan S, Dasgupta N, Mishra RK, Thomas S (eds) Characterization and biology of nanomaterials for drug delivery. Elsevier, Amsterdam, pp 47–76

Part IV Advanced 3D Models

Chapter 16 Methods for Co-culture of Primary Human Extravillous Trophoblast Cells and Uterine Natural Killer Cells Xiaowen Gan, Fen Ning, and Gendie E. Lash Abstract During early pregnancy, fetal-derived extravillous trophoblast cells (EVT) from the placenta invade the maternal decidua and inner third of the uterus where they establish fetal tolerance and remodel the uterine spiral arteries, which ensures establishment of a successful pregnancy. Aberrant EVT invasion and spiral artery remodeling is associated with a number of pregnancy complications including miscarriage, preeclampsia, fetal growth restriction, and placenta accrete. During invasion of the maternal tissues, the EVT interact with a number of different cell types including the decidual leukocytes. EVT express HLA-C, HLA-G, HLA-E, and HLA-F and interact with uterine natural killer (uNK) cells through a series of different receptors. Epidemiological evidence suggests that different combinations of HLA-C and killer cell Ig-like receptor (KIR) haplotypes impact pregnancy success. Therefore, there is much interest in the functional consequence of interactions between EVT and uNK cells, and several different methodologies have been used to isolate these different cell types and their co-culture. Key words Placenta, Extravillous trophoblast cells, Uterine natural killer cells, Cell isolation, Coculture

1 Introduction Establishment of a successful pregnancy requires the coordinated invasion of extravillous trophoblast cells (EVT) through the maternal decidua and into the inner third of the myometrium. This invasion process anchors the placenta to the uterine wall, facilitates spiral artery remodeling, and establishes fetal tolerance. Failure in this process is associated with a number of obstetric complications including preeclampsia, fetal growth restriction, miscarriage, preterm birth, and stillbirth [1–5]. EVT are unique in their expression of human leukocyte antigen (HLA), which is restricted to the class I molecules HLA-C, HLA-G, and HLA-E. These HLA species

Xiaowen Gan and Fen Ning contributed equally with all other contributors. Sandeep Raha (ed.), Trophoblasts: Methods and Protocols, Methods in Molecular Biology, vol. 2728, https://doi.org/10.1007/978-1-0716-3495-0_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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interact with a range of receptors including members of the killer cell Ig-like receptor (KIR), leukocyte Ig-like receptor (LILR), and C-type lectin-like receptor families on uterine natural killer (uNK) cells, decidual macrophages and T cells to aid in establishing fetal tolerance [6, 7]. There is epidemiological evidence that different combinations of HLA-C and KIR haplotypes can lead to adverse pregnancy outcomes such as preeclampsia, fetal growth restriction, or miscarriage [8–11]. Due to these unique interactions between EVT and uNK cells, many researchers have been interested in the functional impact of their interactions [12–14]. There is a paucity of animal models to fully study these interactions, and in vitro culture models have been established. While there are some trophoblast-like cell lines available, such as the immortalized first trimester cell line HTR-8/ SVneo and the choriocarcinoma cell lines BeWo, JAR, and JEG-3 cells, they should be used with caution as they show large discrepancies in gene expression profiles when compared to primary trophoblast cells [15, 16]. Therefore, it is necessary to develop appropriate strategies to isolate and culture primary EVT. In addition, uNK cells are phenotypically distinct from peripheral blood NK cells, and recent single-cell sequencing studies have shown there are three distinct subsets of uNK cells [17, 18]. Here we describe the common methods for isolation of both EVT and uNK cells from the same pregnancies to allow for co-culture studies. Methods for EVT isolation include villous explant outgrowth and enzymatic digestion of first trimester villous or term chorion basal plate. For explant outgrowth, placental tissues are cleaned and dissected before plating on Matrigel in a culture dish and observing outgrowth from the explant [19–22]. For enzymatic digestion of tissues from the first or third trimester, minced tissues are subjected to trypsinization and Percoll gradient centrifugation [23–26]. After isolation, the purity should be assessed using trophoblast markers. Isolation of uNK cells requires enzymatic digestion of the decidua and then either CD56-conjugated magnetic bead column separation or flow cytometry cell sorting [27, 28].

2 2.1

Materials Equipment

Ensure all instruments and solutions are sterile. 1. Container for transfer of placental tissue from the operating room to the laboratory. 2. Scrapers. 3. Flasks. 4. Glass funnel.

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5. Pasteur pipettes. 6. Falcon Centrifuge tube: 15 mL, 50 mL. 7. 150 mm Petri dishes. 8. Large plastic tray. 9. Large metal sieve (100 μm) with waste collection tray, autoclaved to sterilize. 10. Gauze (butter muslin cut to 200-mm squares and then autoclaved to sterilize), Double thickness sterile gauze. 11. 70- and 40-μm cell filters. 12. 20 × 100 mm-style polystyrene cell culture dish. 13. 10 mL syringe. 14. Small sterile surgical scissors and forceps. 15. Centrifuge. 16. Pipettes. 17. Cell counter machine. 18. Cell counter slides. 19. 100 μm cell strainer. 20. 30 μm cell strainer. 21. Flow cytometry tubes. 22. 1.5 mL centrifuge tube. 23. Vortex. 2.2 Solutions and Media

1. 0.9% sodium chloride: 9 g NaCl in 1000 mL deionized water. 2. 10× Hank’s Balanced Salt Solution (HBSS) without phenol red: NaCl 8 g, KCl 400 mg, CaCl2 140 mg, MgSO47H2O 100 mg, MgCl2-6H2O 100 mg, Na2HPO4-2H2O 60 mg, KH2PO4 60 mg, D-Glucose 1 g, NaHCO3 350 mg, 100 mL deionized water. To make 1× HBSS, dilute the 10× HBSS with deionized water. 3. Digestion medium I (for EVT isolation from first trimester villous tissue). 0.125% Trypsin

1 mL 2.5% Trypsin

DNase Ia

200 μL

HBSS

20 mL a

To prepare DNase I, reconstitute 15,000 units in 2.56 mL PBS, 200 μL aliquots and freeze in -20 °C 4. Digestion medium II (for EVT isolation from term villous tissue).

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0.125% Trypsin

125 mg trypsin

Collagenase

100 mg

DMEM

100 mL

5. Newborn calf serum (NCS) and fetal calf serum (FCS). 6. DMEM-F12 complete medium: DMEM-F12 completed with 2 mM L-glutamine, 10% heat-inactivated FCS, 1 IU/mL penicillin, 1 μg/mL streptomycin and 1.5 μg/mL amphotericin-B. 7. Percoll gradient preparation: Density centrifugation medium: 100% Percoll should be diluted to 90% Percoll with 10× HBSS. Preferably use a 50 mL round bottom glass tube for preparing the gradient. Draw a line on the tube between 30% and 35% and another between 50% and 55%. From bottom to top, apply the Percoll gradient in the following sequence: 70%, 60%, 55%, 50%, 45%, 40%, 35%, 30%, 20%, 10% respectively. 2 mL of solution for each concentration. Apply the layers with Pasteur pipettes gently. Percoll gradient (%)

90% Percoll (mL)

1× HBSS (mL)

70

21

6

60

18

9

55

16.5

10.5

50

15

12

45

13.5

13.5

40

12

15

35

10.5

16.5

30

9

18

20

6

21

10

3

24

8. Growth factor reduced Matrigel: Dilute Matrigel to desired concentration using serum-free medium. 9. MACS○R system (Miltenyi Biotech) (see Note 1), gentle MACS C tubes (Miltenyi Biotech), gentle MACS dissociator (Miltenyi Biotech). 10. Class II microbiological safety cabinet. 11. RPMI 1640 medium: culture medium consisting of RPMI 1640+ Glutamax plus 10% FBS and antibiotics. 12. Phosphate buffered saline (PBS) 1× at room temperature. 13. Type IV collagenase.

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14. DNase I. 15. Ficoll lymphocyte isolation solution 1.077 ± 0.001 g/mL) (TBDTM).

(relative

density

16. Erythrocyte lysate. 17. MACS Buffer: Phosphate-buffered saline (PBS), 0.5% bovine serum albumin (BSA) and 2 mM EDTA. Adjust pH to 7.2, filter and remove bacteria with 0.22 μm pin filter, store at 4 °C. 18. Trypsin/EDTA. 19. CD56 antibody-coated magnetic Microbeads. 20. FACS flow cytometer. 21. BSA. 22. PBS solution. 23. 1% Fixing Cells with Paraformaldehyde (PFA). 24. 0.4% Trypan Blue Solution. 25. FITC-conjugated anti-CD56 monoclonal antibody. PerCPconjugated anti-CD16. 26. FACs buffer: PBS supplemented with 2% BSA. Store at 4 °C. Keep sterile. 27. Blocking buffer: Add at 2 μg/mL anti-FcγRII/FcγRIII monoclonal antibody (clone 2.4G2) to PBS with 2% FBS. Prepare fresh each time. Store at 4 °C. 28. Flow Cytometry Staining Buffer. 29. Fixation/permeabilization buffer. 30. Erythrocyte lysate. 31. MACS Buffer: Phosphate-buffered saline (PBS), 0.5% bovine serum albumin (BSA) and 2 mM EDTA. Adjust pH to 7.2, filter and remove bacteria with 0.22 μm pin filter, store at 4. 32. Lymphoprep© (see Note 4).

3

Methods

3.1 Tissue Preparation

Before obtaining tissues, ensure that all appropriate ethical approvals and informed consents have been obtained. Collect first trimester products of conception after surgical election termination of an apparently normal pregnancy. Obtain gestational age using ultrasound measurement of crown rump length and last menstrual period dating. Note that drugs such as mifepristone, misoprostol, and progesterone may influence the endocrine environment of the uterus. Tissue samples should be transferred to the laboratory as quickly as possible, washed extensively in 0.9% saline solution and

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placenta and decidua separated based on gross morphological appearance for EVT and uNK cell isolation respectively. 3.2 First Trimester Villous Explant Outgrowth

1. Pre-warm the DMEM-F12 complete medium to 37 °C. 2. Thaw growth factor reduced Matrigel before use on ice. Using cooled pipettes, mix the Matrigel to homogeneity. Add diluted Matrigel to Petri dishes and incubate for 1 h at room temperature. Remove excess Matrigel and leave a thin coat on each dish. 3. Collect the villous tissue and wash the tissue in sterile 0.9% sodium chloride solution 2–3 times to remove any blood clots and mucus. 4. Carefully examine villous tissue and ensure the absence of decidua and fetal membranes. Use scissors and forceps to cut the tissues into small sections of approximately 2–3 mm3. 5. Resuspend the tissues in DMEM-F12 complete medium and seed them in Matrigel-coated Petri dishes. Culture in a cell culture incubator at 37 °C with 5% CO2. 6. Change the culture medium on a daily basis. After 12–14 days, sufficient EVT will have grown out from the placental explant for further experimentation. 7. To assess the purity of EVT, perform immunofluorescence staining for HLA-G and cytokeratin 7.

3.3 First Trimester Villous Enzymatic Digestion

1. Prewarm the digestion medium I, DMEM-F12 complete medium, and sufficient NCS to 37 °C. Thaw the Matrigel on ice. 2. Obtain first trimester placental tissues and wash them in sterile 0.9% sodium chloride solution. 3. Dissect the tips of the villi, avoiding villi stems and vascular material. Wash dissected tissue 3–4 times in 1× HBSS to remove blood cells. 4. Add the tissue to a 50 mL Falcon tube containing digestion medium I. For approximately 5 g of tissue use 20 mL digestion medium I. Digest tissue for 30 min at 37 °C without shaking. 5. After tissue sediment, gently remove the supernatant and filter through gauze (40 μm pore). 6. To stop the digestion, add NCS to the filtrated supernatant to a final concentration of 5%. 7. Collect the remaining tissue and repeat steps 4–6 above. 8. Combine the two filtered supernatants and centrifuge at 300 × g for 10 min at room temperature. 9. Resuspend the cell pellet in 1 mL DMEM-F12 complete medium.

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10. Prepare the Percoll gradient while tissue is digesting. Layer the resuspended pellet on a Percoll gradient carefully without disrupting the layers. 11. Centrifuge at 1200 × g for 30 min at room temperature. 12. While centrifuging, prepare Matrigel-coated plates or inserts dependent on experimental protocol. For example, use 20 μL for each well of a 24-well plate, 10 μL for inserts. Add diluted Matrigel to plate or insert. Incubate for 1 h at room temperature. Remove excess Matrigel and leave a thin coat on each vessel. 13. Collect the trophoblast cells between the 35% and 50% layer. Dilute with approximately 30 mL DMEM-F12 medium without FCS. 14. Centrifuge at 300 × g for 10 min at room temperature to remove excess Percoll solution. 15. Resuspend the cell pellet in 5 mL DMEM-F12 complete medium. 16. Count cells on a hemocytometer. Plate cells at desired concentration. 3.4 Isolate EVT from Term Placenta with Enzymatic Digestion

1. Carefully dissect the villous tissue and the amniochorion membrane to obtain the term basal plate, which is a 3–6 mm thick membrane. Wash the tissue in sterile 0.9% sodium chloride solution (see Note 1). 2. Mince the tissue into fragments with scissors and forceps. 3. Incubate the fragments in 50 mL digestion medium II for 20 min at 37 °C, shaking with a speed of 150 rpm (see Note 7). 4. Filter the supernatant and collect in a tube pre-filled with NCS. 5. Repeat steps 3 and 4. 6. Combine the two supernatants and centrifuge at 300 × g for 10 min and resuspend the cell pellet in 2 mL DMEM-F12 complete medium. 7. Separate the EVT with 5%, 20%, 30%, 40%, 50%, and 60% Percoll gradient. Centrifuge at approximately 300 × g for 20 min at room temperature. The EVT can be seen in between 35% and 50% (see Note 8). 8. Collect cells located in the layers of interest with a pipette and transfer them to a 50 mL Falcon tube. Make up the volume to 50 mL with DMEM-F12 medium. Centrifuge for 10 min at 300 × g at room temperature. 9. Resuspend the cells pellet and count cells on a hemocytometer. Seed them at preferred density.

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3.5 uNK Cell Isolation Using Magnetic Sorting

Obtain decidual tissue as described above (see Notes 9 and 10) 1. Wash the decidual tissue in PBS or sterile 0.9% saline solution and physically remove any large blood clots (see Note 1). 2. Mince the clean decidual pieces using sterile disposable scalpels into approximately 1 mm2 fragments. 3. Collect the tissue fragments and transfer them to a 50 mL centrifuge tube. 4. Add 15 mL RPMI 1640 medium containing 1.5 mg type I DNase and 24 mg type IV collagenase to the tube containing the tissue fragments. 5. Seal the tube tightly and digest the tissue for 1 h at 37 °C with gentle agitation. 6. Add 20 mL RPMI 1640 medium to the mixture and let it stand for 5 min to allow undigested tissue to sediment. 7. Aspirate the supernatant and filter through the 100-μm and then the 40-μm cell filters to remove debris. 8. Centrifuge the filtrate for 5 min at 300 × g at room temperature, and then resuspend the resultant cell pellet in 40 mL PBS. 9. Centrifuge the cells for 5 min at 300 × g at room temperature, and then resuspend the resultant cell pellet in 4 mL Ficoll lymphocyte isolation solution. 10. Centrifuge the cells for 5 min at 300 × g at room temperature and draw the white film from the liquid with a 1 mL syringe to a 1.5 mL centrifuge tube carefully to obtain lymphocyte cells. 11. For each 1 × 107 lymphocyte cells, add 80 μL MACS Buffer and 20 μL anti-human CD56 binding micromagnetic beads. Incubate on ice for 15 min in the dark. 12. Add 1 mL MACS buffer to every 1 × 107 cells and centrifuge for 5 min at 300 × g at room temperature. Resuspend the resultant cell pellet in 300 μL MACS buffer to remove the unbound antibody magnetic beads (see Note 2). 13. Prepare the MACS○R system. Place the gentle MACS C tubes in a gentle MACS dissociator with a “waste” box beneath. Wash the gentle MACS C tubes with 3 mL MACS buffer and then add the cell suspensions to the gentle MACS C tubes (see Note 3). 14. Wash the tube with 3 mL MACS Buffer 3 times to remove the cells without CD56 beads. 15. Remove the gentle MACS C tubes from gentle MACS dissociator and place it in a collection tube. 16. Add 2 mL of MACS buffer and immediately flush out the labelled cells by applying the plunger.

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17. Centrifuge the filtrate for 5 min at 300 × g at room temperature, and resuspend the uNK cell pellet in 10 mL RPMI 1640 medium for subsequent use. 3.6 uNK Isolation by Flow Cytometry

There are two main types of NK cells: CD56dim + CD16+ and CD56bright + CD16-, uNK cells being of the latter phenotype. Further fractionation can be performed to obtain uNK1, uNK2 or uNK3 subsets (see Note 5) [18]. 1. Refer to the above section to obtain lymphocyte cells. 2. Resuspend the cells with FACs buffer to a total of 100 μL in 1.5 mL centrifuge tube. 3. To Block non-specific Fc-mediated interactions, add 100 μL blocking buffer and incubate on ice for 10–20 min before staining. 4. Add 1 mL FACs buffer to the filtrate. 5. Centrifuge the filtrate for 5 min at 300 × g at 4 °C to collect the cells in the bottom of the 1.5 mL centrifuge tube. 6. Aliquot 50 μL of cell suspension (from 105 to 108 cells) to each tube. 7. Combine the recommended quantity of each primary antibody (CD56, CD16, CD3) according to the instructions in an appropriate volume of Flow Cytometry Staining Buffer such that the final staining volume is 100 μL and add to cells. 8. Pulse vortex gently to mix. 9. Incubate for at least 30 min at 2–8 °C or on ice. Protect from light. 10. Add at least 1 mL FACs buffer to the lymphocyte cells incubated with antibodies. 11. Centrifuge the filtrate for 5 min at 300 × g at 4 °C to pellet the cells. 12. Discard the supernatant and obtain the cell pellet. 13. Resuspend the cell pellet in an appropriate volume of Flow Cytometry Staining Buffer. 14. Run the cells in the Flow cytometry machine FACS flow sorter, collecting the CD56brightCD16- population (see Note 7).

3.7 EVT and uNK Cell Co-culture

Isolate EVT and uNK cells as described above. For co-culture experiments, it is best to obtain EVT and uNK cells from the same products of conception to ensure that the two cell types are semi-allogeneic and remove the confounding variable of genetic differences (see Note 11) [29].

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Direct Co-culture

1. Coat the appropriate number of wells of a 24-well tissue culture plate with Matrigel. 2. Plate 2 × 105 EVT in complete DMEM-F12 medium. 3. Allow to adhere and wash gently prior to addition of 2 × 105 uNK cells in complete DMEM-F12 medium. 4. Culture for desired length of time in a standard cell culture incubator at 37 °C and 5% CO2 in air.

3.9 Indirect Coculture

1. Coat the appropriate number of wells of a 24-well tissue culture plate with Matrigel. 2. Plate 2 × 105 EVT in complete DMEM-F12 medium. 3. Allow to adhere and wash gently. Replace the medium and insert a 0.4 μm cell culture transwell insert to the well. 4. Plate 2 × 105 uNK cells in complete DMEM-F12 medium into the transwell insert. 5. Culture for desired length of time in a standard cell culture incubator at 37 °C and 5% CO2 in air.

4

Notes 1. Ensure the tissue is well washed and free of any obvious blood clots. 2. Always work with ice-cold MACs buffer. 3. There are non-trophoblast cells such as lymphocytes, leukocytes, macrophages, endothelial cells, fibroblasts, and syncytial fragments in the cell harvests. Immunomagnetic microspheres coated with antibodies against specific antigens on cell surface are used to deplete the contaminated cells [15]. 4. Lymphoprep©, a commercially available solution, is used for purification of peripheral blood mononuclear cells by density gradient centrifugation [30]. Although the purification significantly eliminates most of the non-trophoblastic cells, the extra cost for this approach and the reduction in cytotrophoblast yield make this step neglected for most purposes [15]. 5. To assess the quality and content of cell isolation, identification of the amount and the types of trophoblasts, as well as major groups of contaminating cells, are preferred. 6. Confirm the purity of the trophoblast cells by flow cytometry, immunofluorescence staining, or immunocytochemical staining. For identification of contaminating mesenchyme-derived cells, vimentin is considered to be a good marker [16]. The purity of all forms of trophoblast cells is determined with FITC-conjugated monoclonal antibodies against

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cytokeratin-7. HLA-G is a specific marker of the EVT. For EVT subpopulations, those with proliferative features express α6 integrin and can be identified by CD49f VLA6 antibody while an invasive phenotype expresses α1 integrin and can be identified by anti-CD49a alpha1 [15, 31]. After approximately 4 h, rinse the cells twice with warm culture medium to remove unattached cells and syncytial fragments. 7. There are significant variations in the quality of enzymes and batch-to-batch variation in enzyme for trophoblast isolation. Time may need to be adjusted depending on trypsin strength. Some commercially available trypsin preparations do not work at all and several different manufacturers may need to be trialed. 8. When centrifuge in the Percoll gradient solution, the brake should be off in order to obtain better separation. 9. Trophoblast cells may be cryopreserved with reasonable viability. Perform standard procedure to freeze and thaw the trophoblasts. 10. Depending on the outcome measures single cell control cultures should also be established. 11. For cell tracking experiments both cell types respond well to cell tracker dyes such as CFSE, FarRed, etc. (Molecular Probes). References 1. Knofler M et al (2019) Human placenta and trophoblast development: key molecular mechanisms and model systems. Cell Mol Life Sci 76(18):3479–3496 2. O’Tierney-Ginn PF, Lash GE (2014) Beyond pregnancy: modulation of trophoblast invasion and its consequences for fetal growth and longterm children’s health. J Reprod Immunol 104-105:37–42 3. Khong TY et al (1986) Inadequate maternal vascular response to placentation in pregnancies complicated by pre-eclampsia and by smallfor-gestational age infants. Br J Obstet Gynaecol 93(10):1049–1059 4. Pijnenborg R et al (1991) Placental bed spiral arteries in the hypertensive disorders of pregnancy. Br J Obstet Gynaecol 98(7):648–655 5. Ball E et al (2006) Late sporadic miscarriage is associated with abnormalities in spiral artery transformation and trophoblast invasion. J Pathol 208(4):535–542 6. Raulet DH, Vance RE (2006) Self-tolerance of natural killer cells. Nat Rev Immunol 6(7): 520–531

7. Lanier LL (2005) NK cell recognition. Annu Rev Immunol 23:225–274 8. Hiby SE et al (2004) Combinations of maternal KIR and fetal HLA-C genes influence the risk of preeclampsia and reproductive success. J Exp Med 200(8):957–965 9. Hiby SE et al (2008) Association of maternal killer-cell immunoglobulin-like receptors and parental HLA-C genotypes with recurrent miscarriage. Hum Reprod 23(4):972–976 10. Hiby SE et al (2010) Maternal activating KIRs protect against human reproductive failure mediated by fetal HLA-C2. J Clin Invest 120(11):4102–4110 11. Hiby SE et al (2014) Maternal KIR in combination with paternal HLA-C2 regulate human birth weight. J Immunol 192(11):5069–5073 12. Sharkey AM et al (2015) Tissue-specific education of decidual NK cells. J Immunol 195(7): 3026–3032 13. Tilburgs T et al (2015) Human HLA-G+ extravillous trophoblasts: immune-activating cells that interact with decidual leukocytes. Proc Natl Acad Sci U S A 112(23):7219–7224

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14. Lash GE et al (2011) Interaction between uterine natural killer cells and extravillous trophoblast cells: effect on cytokine and angiogenic growth factor production. Hum Reprod 26(9): 2289–2295 15. Cervar-Zivkovic M, Stern C (2011) Trophoblast isolation and culture. In: The Placenta. Wiley, pp 153–162 16. Lash GE et al (2009) IFPA meeting 2008 workshops report. Placenta 30(Suppl A):S4– S14 17. Bulmer JN, Lash GE (2005) Human uterine natural killer cells: a reappraisal. Mol Immunol 42(4):511–521 18. Vento-Tormo R et al (2018) Single-cell reconstruction of the early maternal-fetal interface in humans. Nature 563(7731):347–353 19. Olga Genbacev SAS, Miller RK (1992) Villous culture of first trimester human placentamodel to study extravillous trophoblast (EVT) differentiation. Placenta 13:22 20. Vicovac L, Jones CJ, Aplin JD (1995) Trophoblast differentiation during formation of anchoring villi in a model of the early human placenta in vitro. Placenta 16(1):41–56 21. Caniggia I et al (1997) Endoglin regulates trophoblast differentiation along the invasive pathway in human placental villous explants. Endocrinology 138(11):4977–4988 22. James JL, Stone PR, Chamley LW (2005) Cytotrophoblast differentiation in the first trimester of pregnancy: evidence for separate progenitors of extravillous trophoblasts and syncytiotrophoblast. Reproduction 130(1): 95–103 23. Tarrade A et al (2001) Characterization of human villous and extravillous trophoblasts

isolated from first trimester placenta. Lab Investig 81(9):1199–1211 24. Lash GE et al (2010) Secretion of angiogenic growth factors by villous cytotrophoblast and extravillous trophoblast in early human pregnancy. Placenta 31(6):545–548 25. Biadasiewicz K et al (2014) Extravillous trophoblast-associated ADAM12 exerts pro-invasive properties, including induction of integrin beta 1-mediated cellular spreading. Biol Reprod 90(5):101 26. Borbely AU et al (2014) The term basal plate of the human placenta as a source of functional extravillous trophoblast cells. Reprod Biol Endocrinol 12:7 27. Male V et al (2010) Natural killer cells in human pregnancy. Methods Mol Biol 612: 447–463 28. Male V, Gardner L, Moffett A (2012) Isolation of cells from the feto-maternal interface. Curr Protoc Immunol Chapter 7:Unit 7 40 1–11. https://doi.org/10.1002/0471142735. im0740s97 29. Fonseca BM et al (2020) Decidual NK cellderived conditioned medium from miscarriages affects endometrial stromal cell decidualisation: endocannabinoid anandamide and tumour necrosis factor-alpha crosstalk. Hum Reprod 35(2):265–274 30. Trundley A et al (2006) Methods for isolation of cells from the human fetal-maternal interface. Methods Mol Med 122:109–122 31. Blaschitz A et al (2000) Antibody reaction patterns in first trimester placenta: implications for trophoblast isolation and purity screening. Placenta 21(7):733–741

Chapter 17 Trophoblast Organoids as a Novel Tool to Study Human Placental Development and Function Sandra Haider, Martin Kno¨fler, and Paulina A. Latos Abstract The human placenta provides the site of exchange between the maternal and fetal bloodstreams, acts as an endocrine organ, and has immunological functions. The majority of pregnancy disorders including preeclampsia and fetal growth restriction have their roots in pathological placentation. Yet, the underlying molecular causes of these complications remain largely unknown, not least due to the lack of reliable in vitro models. Recent establishment of 2D human trophoblast stem cells and 3D trophoblast organoids has been a major advancement that opened new avenues for trophoblast research. Here we provide a protocol detailing isolation of cytotrophoblast from the first trimester human placenta, establishment of trophoblast organoids, their culture and differentiation conditions. Overall, we describe an in vitro system that offers an excellent model to study the molecular basis of placental development and disease. Key words Trophoblast organoids, Human placenta, Syncytiotrophoblast, Cytotrophoblast Extravillous trophoblast

1

Introduction The placenta, a defining organ of placental mammals, is essential to sustain embryonic development during pregnancy. Not only does it facilitate exchange of nutrients, gases, and metabolites between the mother and the embryo, but it also has important immunological and endocrine functions crucial for pregnancy adaptations. The placenta is a composite organ comprising the trophectoderm-derived trophoblast component and the mesoderm-derived vasculature. Despite serving analogous functions across the species, placentas exhibit breathtaking morphological and anatomical diversity in terms of cell type specialization and depth of trophoblast invasion [1]. Consequently, animal placentas provide limited models of the human organ.

Sandeep Raha (ed.), Trophoblasts: Methods and Protocols, Methods in Molecular Biology, vol. 2728, https://doi.org/10.1007/978-1-0716-3495-0_17, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Fig. 1 Illustration of a first trimester placenta villous tree and trophoblast organoids (TB-ORG). In floating villi, the villous core (VC) surrounds the fetal capillaries (FC) and is covered by the cytotrophoblast (CT) that fuses into the multinucleated syncytiotrophoblast (ST). Anchoring villi attach to the maternal decidua. CT gives rise to the cell column trophoblast (CCT) that further differentiates into invasive extravillous trophoblast (EVT). EVTs specialize into interstitial EVTs (iEVT). A subset of iEVTs invade spiral arteries (mSA) to replace maternal endothelial cells as endovascular EVT (eEVT)

Following implantation of the human blastocyst, the trophectoderm layer gives rise to the mononuclear cytotrophoblast (CT) and primitive syncytium (ST). Subsequently, the CT proliferates and migrates through the primary syncytium, and eventually forms the primary villi. The extra-embryonic mesoderm then gives rise to the villous mesenchymal compartment, including fetal capillaries. The villi continue to rapidly branch, expand, and form the villous trees [1–3]. The basic unit of the first trimester placenta, the villous tree, consists of a varying number of anchoring villi attached to the maternal uterus through cell columns (CCT) and the floating villi bathed in maternal blood (Fig. 1). While the base of the column consists of rows of proliferative progenitor cells, the distal tip of the column gives rise to the extravillous trophoblast (EVT), invading the inner lining of the maternal uterus. Deciduaembedded interstitial EVTs (iEVTs) modulate and attenuate the maternal immune responses, and ensure the acceptance of the fetal semi-allograft. A subpopulation (endovascular EVTs, eEVTs)

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colonizes the maternal spiral arteries and enlarges the arterial lumina in order to generate vessels of low resistance [1–3]. In contrast, the floating villi consist of the proliferative cytotrophoblast layer covered by the multinucleated syncytiotrophoblast (ST) layer. The ST arises from an asymmetric cell division, differentiation, and fusion of the underlying CT, and provides the site of exchange between the maternal and fetal bloodstreams. As such, the ST is densely packed with a plethora of transporters, including those for amino acids and glucose, and contains numerous microvilli that dramatically increase the exchange surface. Importantly, the ST also serves as an endocrine organ, which produces and secretes vital pregnancy hormones including human chorionic gonadotropin (hCG), placental lactogen, and pregnancy-specific glycoproteins (PSGs). Defects in specification and morphogenesis of these highly specialized trophoblast cell types result in a range of placental disorders and pregnancy complications, such as pre-eclampsia, fetal growth restriction, and preterm birth. Until recently, the studies of molecular mechanisms driving human placental development and disease were hampered by the lack of reliable, physiologically relevant in vitro models. Commonly used systems included choriocarcinoma-derived cell lines (JEG3, JAR, BeWo, ACH3P), transformed cell lines (HTR-8/SVneo, SGHPL-4-5), primary cell cultures, and human embryonic stem cell-derived cultures [1, 2]. While these models recapitulate certain aspects of trophoblast development and/or differentiation, they have numerous caveats including material availability, reproducibility, unclear molecular identity, and cancer-like physiology. A breakthrough came in 2018 when Okae and colleagues established human trophoblast stem cells (hTSCs) derived from both primary CT and human blastocysts [4]. Human TSCs self-renew and, when induced, differentiate into both ST and EVTs that mirror the molecular identity signatures of their in vivo counterparts [4]. Subsequently, we and others derived human trophoblast organoids from primary CT [5, 6]. In the past decade, 3D organoids revolutionized the modeling of human development and disease [7]. The key hallmark of 3D organoids is self-organization by spontaneous developmental patterning and morphogenesis that enables them to recapitulate aspects of tissue biology and architecture. Self-organizing organoids can be derived either from tissue stem cells/progenitors or from pluripotent stem cells [7, 8]. Formation of epithelial structures of most human organs (gut, liver, stomach, lung, retina, brain, kidney, etc.) was successfully recapitulated in vitro using remarkably similar organoid culture conditions [9]. These conditions include embedding in Matrigel containing a host of extracellular matrix proteins, such as laminins, which are important for establishing a basolateral-apical epithelial polarity [10].

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Common organoid medium (OM) components include stimulation of EGF-MAPK and canonical WNT signaling (by Rspo-1 or CHIR99021) that drive mitogenic proliferation and stem cell selfrenewal, respectively. Moreover, inhibition of TGF-beta/BMP signaling (by Noggin, LDN, and A83-01) is vital to quench epithelial differentiation [7, 8]. Finally, auxiliary components, such as PGE2 and Nicotinamide, further stimulate proliferation and survival of multiple types of epithelia within self-organizing organoid systems. Based on the rationale that CT exhibits epithelial properties, we successfully applied these culture conditions and were the first to report the derivation of human trophoblast organoids [5]. Since then, both hTSCs and organoid models have been widely used, providing novel insights into the molecular mechanisms underlying trophoblast identity [10–12, 15–21].

2

Materials 1. First trimester placenta (gestational week 6–7). Usage of human primary tissues must comply with the national and institutional regulations. Consult appropriate local institutions for the code of conduct. Handle tissues as potentially hazardous for group 3 blood-borne pathogens and use appropriate personal protective equipment. 2. Advanced Dulbecco’s Modified Eagle Medium: Nutrient Mixture F12 (DMEM/F12) medium. 3. 50× B27 supplement. 4. 100× Insulin-Transferrin-Selenium-Ethanolamine supplement.

(ITS-X)

5. 100× L-glutamine. 6. 1 M HEPES. 7. 10 mg/mL gentamicin. 8. Recombinant human epidermal growth factor (EGF) protein. 9. CHIR 99021. 10. A83-01. 11. Y-2637. 12. Recombinant human transforming growth factor beta-1 (TGFβj1). 13. Matrigel® Matrix, growth factor reduced. 14. Cell Recovery Solution (CRS). 15. TrypLE. 16. 10× phosphate buffered saline (PBS). 17. Cellbanker™ 2.

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18. Fetal bovine serum (FBS). 19. Percoll™. 20. 2.5% Trypsin. 21. DNAse I. 22. 10× Hanks’ Balanced Salt Solution (HBSS). 23. Histosec Pastilles. 24. Anti-phycoerythrin (PE) MicroBeads. 25. Anti-human leukocyte antigen G (HLA-G) PE. 26. autoMACS Rinsing Solution. 27. Formaldehyde (FA). 28. Xylene. 29. Alcian blue solution. 2.1 Tissue Culture Plastic

1. MS columns for the positive selection of cells (Miltenyi Biotec, cat. no. 130-042-201). 2. 40 μm, 70 μm and 100 μm cell strainers. 3. 4-well / 24-well culture dishes. 4. Plastic pipettes, sterile. 5. 50 mL Tubes. 6. 15 mL Tubes. 7. Tissue homogenizing no. 91-PCS-CKM).

3

kit,

CKMix

(Peqlab/VWR,

cat.

Methods

3.1 General Instructions

1. Perform all experiments under sterile conditions in a biosafety cabinet. 2. Clean everything using 70% EtOH before it is placed under the biosafety cabinet. 3. Use an aspirator to remove supernatants/solutions as tissues, cells, and organoids are not firmly attached at the bottom of the tubes due to gentle centrifugation steps. 4. Keep all buffers, solutions, and media sterile and at 4 °C unless otherwise stated. It is recommended to use ice buckets with crushed ice.

3.1.1 Preparation of Solutions

1. 1× PBS: Dilute 10× PBS with sterile, deionized water. 2. 1× Mg2+/Ca2+-free HBSS: Dilute 10× HBSS with sterile, deionized water. 3. FBS: Prepare 25 mL aliquots and store them at -20 °C.

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Table 1 Dilution series for the Percoll gradient Percoll (%)

90% Percoll (mL)

HBSS (mL)

70

7

2

60

6

3

55

5.5

3.5

50

5

4

45

4.5

4.5

40

4

5

35

3.5

5.5

30

3

6

20

2

7

10

1

8

4. 90% Percoll: Mix 40.5 mL Percoll with 4.5 mL 10× HBSS. Always prepare fresh. Prepare a dilution series from 70% to 10% (Table 1). The dilutions can be stored at 4 °C for up to 4 weeks. 5. Erythrocyte lysis buffer: 155 mM NH4Cl, 10 mM KHCO3, 0.1 mM EDTA, pH 7.3. Sterile filter or autoclave the buffer. Store at room temperature. Note that proper erythrocyte lysis requires the exact pH of 7.3. 6. Matrigel: Thaw Matrigel according to the manufacturer’s instructions (in a box with crushed ice at 4 °C overnight). Aliquot 0.5 mL into 1.5 mL tubes under sterile conditions. Store the aliquots at -20 °C. Once thawed, keep the tubes at 4 °C for up to 2 weeks. Handle Matrigel on ice all the time as it will quickly solidify at room temperature. Pipette Matrigel carefully to avoid any air bubbles. 7. Placenta digestion solution: 1× HBSS, 0.25% Trypsin, 0.625 mg/mL DNAse I. 8. 2.5% Trypsin: Prepare 5 mL aliquots in 15 mL conical tubes. Store the aliquots at -20 °C. 9. 0.1 g/mL DNAse I: Dissolve the DNAse I in sterile 1× HBSS and 25 mM HEPES on a roll mixer. Prepare 1 mL aliquots of the DNAse I and store them at -80 °C. 10. Organoid medium (OM) for organoid formation from isolated villous CTs (vCTs): Advanced DMEM/F12, 1× B27, 1× ITS-X, 1× glutamate, 0.01 M HEPES, 0.1 mg/mL gentamicin, 100 ng/mL EGF, 3 μM CHIR99021, 1 μM A83-01, 5 μM Y-2637.

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11. OM for organoid propagation: Advanced DMEM/F12, 1× B27, 1× ITS-X, 1× glutamate, 0.01 M HEPES, 0.1 mg/mL gentamicin, 100 ng/mL EGF, 3 μM CHIR99021, 1 μM A83-01. 12. EVT differentiation medium (EVT-DM): Advanced DMEM/ F12, 1× B27, 1× ITS-X, 1× glutamate, 0.01 M HEPES, 0.1 mg/mL gentamicin, 100 ng/mL EGF, 1 μM A83-01. 13. EVT differentiation medium-2 (EVT-DM-2): Advanced DMEM/F12, 1x B27, 1x ITS-X, 1x glutamate, 0.01 M HEPES, 0.1 mg/ml gentamicin, 5 ng/ml TGFβj1. 14. Organoid washing media (wash-OM): Advanced DMEM/ F12, 0.1 mg/mL gentamicin. 15. Organoid thawing media (thaw-OM): Advanced DMEM/ F12, 10% FBS. 3.2

3.2.1

Isolation of vCT

Tissue Preparation

Isolation of vCT is performed as described previously [13, 14] with some modifications. The general procedure is illustrated in Fig. 2. The vCT isolation protocol includes tissue preparation (Subheading 3.2.1), tissue digestion (Subheading 3.2.2), Percoll gradientdependent vCT enrichment (Subheading 3.2.3), and erythrocyte removal (Subheading 3.2.4). This process is summarized in Fig. 2a 1. Prepare cold 1× PBS and 1× HBSS. 2. After surgery, cool the placental tissue to 4 °C and process it within 2 h. 3. Prepare two 10 cm cell culture dishes with 20 mL of cold PBS each. Use sterile plastic forceps to gently wash the placental tissue in the cold PBS of the first culture dish in order to remove blood clots and non-placental tissue (e.g., decidua). Directly pluck any sticky blood clots from the placental tissue with sterile forceps. Transfer the placental tissue into the second PBS-containing dish to further rinse the tissue. 4. Pipette 20 mL cold 1× HBSS into a 10 cm cell culture dish. Transfer the washed placenta into the dish and cut the villi from the chorionic membrane using sterile plastic forceps and scalpels. 5. Further mince the placental villi into small pieces (2–4 mm). 6. Transfer the placental villi in 1× HBSS into 50 mL conical tubes and pellet tissue fragments using a centrifuge (at 1.000 rpm (~100 × g), 1 min, 4 °C). Carefully remove the supernatant and keep the tissue on ice. Proceed with tissue digestion.

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Fig. 2 Isolation of cytotrophoblast (CT) from human first trimester placenta and derivation of TB-ORG. (a) Placental tissue is washed to remove blood clots, cut from the chorionic membranes, and minced. (b) Villous fragments are enzymatically digested and loaded on a Percoll gradient. CT is enriched and collected. (c) CT is embedded in a 60:40 Matrigel:medium ratio, plated as domes, and covered with organoid medium 3.2.2 Tissue Digestion

This protocol is designed as a 3–4 step sequential digestion (Fig. 2b), to carefully remove placental cells layer by layer. The tissue texture of early placentas and the used enzymes enable the enrichment of trophoblast populations with a minimal stromal cell contamination (see Note 1).

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Estimate the size of the tissue pellet using the conical tube scale after (see point 6 of the tissue preparation) (see Note 2). Herein, the digestion for 2–5 mL tissue pellet is described: 1. Prepare four 50 mL conical tubes: Pipette 18 mL of 1× HBSS into three tubes (digestion 1–3) and 9 mL of 1× HBSS into one tube (digestion 4). Pre-warm the tubes in a 37 °C water bath. 2. Thaw 7 mL of trypsin and 0.45 mL of DNAse I. Keep both on ice. 3. Prepare four 50 mL conical tubes with 5 mL FBS each. Keep on ice. 4. Add 2 mL trypsin and 0.125 mL DNAse I into 18 mL pre-warmed HBSS. 5. Pipette the solution onto the tissue pellet and invert the tube 4–6 times. Incubate it for 8–10 min at 37 °C in a water bath. Invert the tube 3–5 every 2 min (first digestion). 6. Remove the tube from the water bath and let the tissue settle for 2 min. 7. Put a sterile 100 μm cell strainer onto one of the FBS-containing tubes from step 3. 8. Pipette the first digestion solution from step 6 onto the cell strainer to separate undigested tissue fragments. 9. Mix the FBS with the first digestion solution by inverting the tube 3–5 times. Pipette a 5 μL sample onto a glass slide and examine the content under a phase-contrast microscope. Discard the first digestion as it predominantly contains ST fragments and CCT. 10. Transfer the undigested tissue from the cell strainer into a fresh 1× HBSS-containing tube from step 1 using sterile forceps. Store the cell strainer for reuse in a sterile 10 cm culture dish in a biosafety cabinet. 11. Add 2 mL of trypsin and 0.125 mL of DNAse I. Invert the tube 4–6 times and incubate it for 15 min at 37 °C in a water bath. Invert the tube 3–5 times every 2 min (second digest). 12. Remove the tube from the water bath and let the tissue settle down for 2 min. 13. Place the used 100 μm cell strainer onto a fresh FBS-containing tube from step 3. 14. Filter the second digestion solution through the cell strainer. Mix the FBS with the digestion solution by inverting the tube 3–5 times. Pipette a 5 μL sample onto a glass slide and examine the content under a phase-contrast microscope. The second digestion solution should already contain single cells and beads on string-like cell groups of vCT. Fill the tube up to 50 mL with 1× HBSS and store the second digestion on ice.

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15. Place the undigested tissue from the cell strainer into a fresh 1× HBSS-containing tube from step 1 using sterile forceps. Store the cell strainer in a sterile 10 cm culture dish in the biosafety cabinet. 16. Add 2 mL of trypsin and 0.125 mL of DNAse I. 17. Invert the tube 4–6 times and incubate it for 15 min at 37 °C in a water bath. Invert the tube 3–5 times every 2 min (third digest). 18. Remove the tube from the water bath and let the tissue settle down for 2 min. 19. Place the used 100 μm cell strainer onto a fresh FBS-containing tube from step 3. 20. Filter the third digestion solution through the cell strainer. Mix the FBS with the digestion solution by inverting the tube 3–5 times. Pipette a 5 μL sample onto a glass slide and examine the content under a phase-contrast microscope. The third digestion also contains single cells and beads on string-like cell groups of vCT. Fill the tube up to 50 mL with 1× HBSS and store the third digestion on ice. 21. Transfer the undigested tissue from the cell strainer into the last fresh 1× HBSS-containing (9 mL) tube from step 1 using sterile forceps. Store the cell strainer in a sterile 10 cm culture dish in the biosafety cabinet. 22. Add 1 mL of trypsin and 0.0625 mL DNAse I. 23. Invert the tube 4–6 times and incubate it for 10 min at 37 °C in a water bath. Invert the tube every 2 min. 24. Remove the tube from the water bath and let the tissue settle down for 2 min. 25. Place the used 100 μm cell strainer onto a fresh FBS-containing tube from step 3. 26. Filter the fourth digestion solution through the cell strainer. Mix the FBS with the digestion solution by inverting the tube 3–5 times. Pipette a 5 μL sample onto a glass slide and examine the content under a phase-contrast microscope. The fourth digestion may still contain single cells and beads on stringlike cell groups of vCT. If so, fill the tube up to 50 mL with HBSS and store the fourth digestion on ice. If no cells are visible, discard the fourth digest. 27. Centrifuge the digestion solutions 2, 3, and potentially 4 at 1.500 rpm (~160 × g) for 5 min at 4 °C. Discard the supernatants, resuspend the pellets in 5 mL 1× HBSS, and pool all the samples. Perform another centrifugation at 1.500 rpm (~160 × g) for 5 min at 4 °C. Discard the supernatant and keep the pellet on ice.

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The Percoll gradient will enrich vCT populations and remove cell debris, stromal, and blood cells. The vCT will predominantly collect between 35% and 55% of Percoll concentrations. Perform one Percoll gradient for up to 5 mL of starting tissue (Fig. 2b). 1. Prepare the Percoll gradient fresh (e.g., during one of the digestion steps). Mark a 50 mL conical tube at the 9 mL and 20 mL scale. 2. Use a tube holder to position the tube at a 30–40° angle. Fix the tube with adhesive tape. 3. Set the suction/expulsion speed on the pipette controller to a minimum level and insert a 5 mL pipette. 4. Invert the 70% Percoll dilution 3 times, and pipette slowly 3 mL along the tube wall into the tilted tube. Do not change the pipette. 5. Invert the 60% Percoll dilution 3 times, and carefully pipette 3 mL of it along the tube wall on top of the first layer. 6. Continue with the same procedure for all remaining Percoll dilutions. 7. Turn the gradient-containing tube in an upright position. The gradient may be stored at 4 °C until use (see Note 3). 8. Place a 70 μm cell strainer on top of a 50 mL conical tube. Resuspend the cell pellet from Subheading 3.2.2, step 27 in 5 mL 1× HBSS and filter the cell solution. 9. Tilt the Percoll gradient and carefully pipette the filtered cell solution on top of the gradient. 10. Perform a centrifugation step at 2.350 rpm (~400 × g) for 24 min at 4 °C. Make sure that the acceleration and deceleration of the centrifuge are reduced to 1 and 0, respectively (see Note 4). After the centrifugation, cloudy rings of the assembled cell population will be visible. 11. Use sterile plastic pipettes to carefully remove and discard the Percoll layers until the 20 mL mark is reached (see Note 5). 12. Use a fresh pipette to collect the CT-containing Percoll layers (35–55% Percoll concentration) until the 9 mL mark in a 50 mL conical tube. Discard the remaining gradient. 13. Invert the tube 4–6 times and divide the solution into four 50 mL conical tubes. The volume should not exceed 5 mL per tube since too high Percoll concentrations decrease cell pelleting efficiency. 14. Fill the tubes up with 1× HBSS. 15. Centrifuge the samples at 1.500 rpm (~160 × g) for 5 min at 4 °C. Small cell pellets should be visible.

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16. Discard the supernatants. Resuspend and pool the pellets in 1× HBSS in one 50 mL conical tube. Fill the tube up with HBSS and perform an additional centrifugation step (1.500 rpm (~160 × g), 5 min, at 4 °C). Discard the supernatant and store the pellet on ice. 3.2.4 Removal of Erythrocytes

If the cell pellet appears reddish, start with step 1. If the cell pellet is completely white, proceed with step 4. 1. Resuspend the cell pellet in 5 mL of erythrocyte lysis buffer. Keep the cells in the 50 mL conical tube. Incubate the cells on a roller mixer at room temperature for 5 min. 2. Fill the tube up with 1× HBSS (it will turn pink). Centrifuge the cells at 1.500 rpm (~160 × g) for 5 min at 4 °C. 3. If the pellet still appears reddish, repeat steps 1 and 2. If the pellet is white, proceed with step 4. 4. Resuspend the cell pellet in 10 mL 1× HBSS and count the cells. 5. Decide on the number of domes for organoid derivation (see Note 6). Transfer the required number of cells into a new tube and store it on ice for organoid derivation (see Subheading 3.3). The remaining cells can be pelleted and frozen. It is recommended to freeze 0.5 – 5 × 106 cells using 0.5 – 1 mL Cellbanker 2 freezing solution. Centrifuge the cells at 1.500 rpm (~160 × g) for 5 min at 4 °C and resuspend in Cellbanker 2 solution and freeze aliquots at -80 °C overnight followed by liquid nitrogen long-term storage. This material can be used for later organoid derivation, and subsequent analyses of the starting cell population.

3.3 Formation of Trophoblast Organoid (TB-ORG) Domes

For TB-ORG formation, isolated vCT is first resuspended in an organoid medium (vCT-OM) (Fig. 2c). Matrigel is added to reach a final ratio of 40:60 vCT-OM:Matrigel. The following procedure describes the formation of four TBORG domes, each consisting of 40 μL vCT-OM/Matrigel. Upscale as needed. 1. Place a small ice bucket with crushed ice into the biosafety cabinet. 2. Place the tubes with 4 × 105 vCT cells (from Subheading 3.2.3), the Matrigel, the wash-OM, and vCT-OM on ice (see Note 7). 3. Place a 4-well culture dish in the incubator. 4. Centrifuge the cells at 1.500 rpm (~160 × g) for 5 min at 4 °C. 5. Resuspend the cells in 0.5 mL wash-OM. 6. Centrifuge the cells at 1.500 rpm (~160 × g) for 5 min at 4 °C.

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7. Resuspend the cells in vCT-OM to reach a final volume of 64 μL (be precise, see Note 8). 8. Pipette 96 μL of Matrigel to the vCT-OM/cell mixture and mix carefully by pipetting up and down 3–5 times. Avoid making air bubbles (see Note 9). 9. Place the pre-warmed 4-well plate into the biosafety cabinet (see Note 10). 10. Pipette 40 μL of the vCT-OM/cell/Matrigel mixture per well into the center of each well. 11. Carefully transfer the plate into the incubator. 12. Incubate the plate for 1–2 min at 37 °C. 13. Turn the plate upside down in one swift movement and incubate the plate for 15 min in the incubator. The cells will evenly distribute in the solidifying vCT-OM/Matrigel. 14. Pre-warm 2 mL of vCT-OM to 37 °C. 15. After 15 min, turn the plate again and add 0.5 mL of vCT-OM per well. Ensure that the dome is covered. 16. Monitor the cultures on a regular basis and exchange the vCT-OM every 3–4 days (Fig. 3). Visible growth of TB-ORG starts after 2–3 days. After 5–8 days, TB-ORG need to be split for the first time.

Fig. 3 Illustration (upper panel) and bright field images (lower panel) of growing and fully mature TB-ORG. After 7 days of culture, small TB-ORG are visible, and after 2–3 weeks, mature TB-ORG have formed. TB-ORG consist of undifferentiated cytotrophoblast (CT) encapsulating differentiated syncytiotrophoblast (ST), as indicated on the TB-ORG section stained with Periodic Acid-Schiff (PAS)

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3.4 Splitting of TBORG

Depending on the organoid size and culture density, use one of the following two procedures: see Subheading 3.4.1 for domes containing a high density of small TB-ORG and see Subheading 3.4.2 for domes containing large TB-ORG.

3.4.1 Splitting of Four TB-ORG Domes with High Density of Organoids

Note that the usual splitting ratio is 1:4. 1. Place a small ice bucket with crushed ice into the biosafety cabinet. 2. Place a sterile 1.5 mL tube, a sterile 15 mL conical tube, Matrigel, wash-OM, and OM on ice (see Note 7). 3. Place a 24-well plate (or four 4-well plates) into the incubator. 4. Remove and discard the organoid medium. Add 0.5 mL CRS per well. 5. Place the plate into a sterile bag and transfer it to the fridge for 35–45 min in order to dissolve the Matrigel. 6. Gently pipette the TB-ORG/CRS mixture up and down 4–6 times and collect the solution in the pre-cooled 15 mL tube. 7. Pellet the TB-ORG at 1.500 rpm (~160 × g) for 5 min at 4 °C (see Note 11). Remove and discard the CRS. 8. Resuspend the TB-ORG pellet in 0.5 mL wash-OM and transfer the solution into the pre-cooled 1.5 mL vial. 9. Centrifuge the TB-ORG at 1.500 rpm (~160 × g) for 5 min at 4 °C. 10. Resuspend the TB-ORG in OM to reach a final volume of 256 μL (be precise, see Note 8). Pipette the solution up and down several times until it appears homogeneous. 11. Carefully mix the TB-ORG/OM with 384 μL Matrigel. Avoid air bubbles (see Note 9). 12. Place the pre-warmed 24-well plate into the biosafety cabinet (see Note 10). 13. Place 40 μL of the TB-ORG/OM/Matrigel mixture into the center of each of the 16 wells of the 24-well dish. Gently re-mix the solution occasionally (after 4 - 6 domes) to ensure equal organoid load per dome. 14. Carefully transfer the plate into the incubator for 1 min. 15. With one swift movement, turn the plate upside-down and incubate it for 15 min at 37 °C. 16. Pre-warm 8 mL OM. 17. After 15 min, pipette 0.5 mL OM per TB-ORG well. Make sure the domes are covered with medium (see Note 12). 18. Monitor the growth of TB-ORG regularly. 19. Change the medium every 2–4 days.

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Note: Unlike isolated cells, TB-ORG have a higher density and tend to sink to the bottom of the well. This risk is further increased when numerous domes are created at once. As a consequence, the TB-ORG attach to the bottom of the well instead of being embedded in Matrigel (see Note 13). 3.4.2 Splitting of Two Domes Containing Large TB-ORG

Note that large TB-ORG are completely dissolved to single cell cultures containing vCT stem cells for starting TB-ORG anew. Successful splitting and culturing of large organoids requires the removal of differentiated STB parts as they will not form TB-ORG. Depending on the TB-ORG size and quantity, it is recommended to split the cultures at a 1:2 or 1:3 ratio. 1. Place a small ice bucket with crushed ice into the biosafety cabinet. 2. Place a sterile 1.5 mL vial, a sterile 15 mL conical tube, Matrigel, PBS, wash-OM, and OM on ice (see Note 7). 3. Place a new 4-well culture plate into the incubator. 4. Carefully aspirate and discard the vCT-OM and add 0.5 mL CRS per well. 5. Place the plate into a sterile plastic bag (to avoid contamination) and transfer the plate into the fridge for 35–45 min in order to dissolve the Matrigel. 6. Gently pipette the TB-ORG/CRS mixture up and down 4–6 times and collect the solution in the pre-cooled 15 mL tube. 7. Pellet the TB-ORG at 1.500 rpm (~160 × g) for 5 min at 4 °C. 8. A TB-ORG pellet should be clearly visible. Remove and discard the supernatant. 9. Add 1 mL TrypLE to the TB-ORG. Mix and incubate at 37 °C for 10 min in a water bath. 10. Mix by inverting every 3 min. 11. Place a 40 μm cell strainer on top of a 50 mL conical tube. 12. Remove the 15 mL tube from the water bath and pipette the TB-ORG up and down for 10–15 times until the solution appears almost homogeneous. 13. Filter the TB-ORG/TrypLE mixture through the cell strainer. 14. Add cold 1× PBS to a total volume of 5 mL. Invert the tube 3–5 times. 15. Pellet the cells at 1.500 rpm (~160 × g) for 5 min at 4 °C. 16. Resuspend the pellet with 0.5 mL wash-OM and pipette the solution into the pre-cooled 1.5 mL tube. 17. Pellet the cells at 1.500 rpm (~160 × g) for 5 min at 4 °C.

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18. Resuspend the TB-ORG with OM to reach a final volume of 64 μL (be precise, see Note 8). Pipette the solution up and down several times to get a homogeneous solution. 19. Mix the TB-ORG/OM with 96 μL Matrigel. Be careful to avoid any air bubbles (see Note 9). 20. Place the pre-warmed 4-well dish into the biosafety cabinet (see Note 10). 21. Place 40 μL of the TB-ORG/OM/Matrigel mixture into the center of each well of the 4-well dish. 22. Carefully transfer the plate into the incubator. Incubate the plate for 1–2 min at 37 °C. 23. With one swift movement, turn the plate upside down and incubate the plate for 15 min at 37 °C. 24. Pre-warm 2 mL OM. 25. After 15 min, pipette 0.5 mL OM per TB-ORG well. Ensure that all the domes are covered. 26. Monitor the TB-ORG growth and change the OM every 3–4 days. TB-ORG should emerge after 6–8 days. 3.5 Freezing and Thawing of TB-ORG

3.5.1 ORG

Freezing of TB-

Viable TB-ORG can be frozen for long-term storage. It is recommended to pool TB-ORG from at least two densely packed TB-ORG domes per cryovial. After thawing, it is recommended to plate the TB-ORG in four domes (1:2 split). 1. Label and place 1.5 mL cryovials (ideally with a screw cap) on ice. 1. Follow steps 1–11 of Subheading 3.4.1. 2. Gently resuspend the TB-ORG in 0.5 mL cold Cellbanker 2 per sample. 3. Pipette the solution into the 1.5 mL cryovials. 4. Place the cryovials into a -80 °C freezer. 5. Transfer the cryovials into liquid nitrogen storage within 3 weeks (see Note 14).

3.5.2

Thawing of TB-ORG

1. Pre-warm 10 mL of thaw-OM in a 15 mL conical tube to 37 °C in a water bath. 2. Place a small ice bucket with crushed ice into the biosafety safety cabinet (see Note 7). 3. Place a sterile 1.5 mL tube, Matrigel, wash-OM, and OM on ice. 4. Place a 4-well plate into the incubator. 5. Place the TB-ORG/Cellbanker 2-containing cryovial at 37 °C in a water bath to thaw.

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6. Transfer the TB-ORG/Cellbanker 2 solution into the pre-warmed thaw-OM in the biosafety cabinet. 7. Invert the tube 4–6 times. 8. Pellet the TB-ORG at 1.500 rpm (~160 × g) for 5 min at 4 °C. 9. Resuspend the TB-ORG pellet in 0.5 mL of wash-OM and transfer it to the pre-cooled 1.5 mL vial. 10. Centrifuge the TB-ORG at 1.500 rpm (~160 × g) for 5 min at 4 °C. 11. Resuspend the TB-ORG in OM to reach a final volume of 64 μL (be precise, see Note 8). Pipette the solution up and down several times to get a homogeneous solution. 12. Mix the TB-ORG-OM with 96 μL Matrigel. Be careful to avoid any air bubbles (see Note 9). 13. Place the pre-warmed 4-well plate in the biosafety cabinet. 14. Place 40 μL of the TB-ORG/OM/Matrigel mixture into the center of each well of the 4-well dish. 15. Carefully transfer the plate into the TC incubator, and incubate it for 1–2 min at 37 °C. 16. Turn the plate upside down in one swift movement and incubate it for 15 min at 37 °C. 17. Pre-warm 2 mL of OM. 18. After 15 min, pipette 0.5 mL OM into each TB-ORG well. Ensure that the TB-ORG domes are covered. 3.6 TB-ORG Differentiation

TB-ORG – ST formation: TB-ORG consist of vCT that fuses toward the center and form hormone-producing, multinucleated ST-like structures (Fig. 4) [5, 6]. Compared to the architecture of placental villi, where ST overlays the CT, TB-ORG display an inside-out structure. This arrangement likely results from Matrigel providing basement membrane-like components and determining polarity of the CT and in turn ST formation. Thus, ST differentiates spontaneously in the inner part of the TB-ORG and recapitulates the in vivo process. TB-ORG differentiation – EVT induction: EVT differentiation is initiated by ceasing WNT activation (removal of CHIR99021 from the OM) while TGFβj signaling is still inhibited. As a consequence, HLA-Gpos EVTs arise from TB-ORG (Fig. 5). After 4–5 days, however, in order to obtain fully mature, secretory EVTs, TGFβj signaling needs to be allowed (removal of A83-01 from the EVT-DM) and activated (addition of TGFβj1, EVT-DM2) for an additional 4–5 days [15]. The EVT differentiation can be initiated either a) directly after splitting, or b) in embedded TB-ORG. Both protocols should be tested.

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Fig. 4 TB-ORG consist of the cytotrophoblast (CT) and the syncytiotrophoblast (ST). Immunofluorescence analyses of TB-ORG (upper panel) and first trimester placental tissue (lower panel). TEAD4 and p63 mark CT, ENDOU stains ST, and DAPI indicates nuclei. In placental tissue, the ST form the outer layer covering the CT, while TB-ORG display an inside-out structure with the inner ST and outer CT

Fig. 5 TB-ORG can be used as a model for EVT differentiation. Brightfield and immunofluorescence images demonstrate that modification of the organoid medium (lower panel) elicits the outgrowth of EVT from TB-ORG, as indicated by expression of the EVT marker HLA-G. DAPI marks nuclei 3.6.1 EVT Induction Directly After Splitting

EVT induction using two confluent TB-ORG domes as starting material is described below. 1. Place a small ice bucket with crushed ice into the biosafety cabinet. 2. Place two sterile 1.5 mL tubes, Matrigel, wash-OM, and EVT-DM on ice (see Note 6).

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3. Place two fresh 4-well plates into the incubator. 4. Remove and discard the OM. Add 0.5 mL CRS to two TBORG wells each. 5. Put the plate into a sterile plastic bag and transfer the plate into the fridge for 35–45 min in order to dissolve the Matrigel. 6. Gently pipette the TB-ORG/CRS mixture up and down 4–6 times and collect it in the pre-cooled 1.5 mL tube. 7. Pellet the TB-ORG at 1.500 rpm (~160 × g) for 5 min at 4 °C. 8. Remove and discard the supernatant. 9. Resuspend the TB-ORG pellets in 0.5 mL wash-OM. 10. Centrifuge the TB-ORG at 1.500 rpm (~160 × g) for 5 min at 4 °C. Discard the supernatant. 11. Resuspend the TB-ORG pellets with EVT-DM to reach a final volume of 128 μL (be precise, see Note 7). 12. Carefully mix the TB-ORG/EVT-DM with 192 μL Matrigel. Avoid air bubbles (see Note 8). Keep on ice. 13. Place the pre-warmed 24-well plates into the biosafety cabinet (see Note 10). 14. Pipette 40 μL of the TB-ORG/EVT-OM/Matrigel mixture into the center of the wells of the 4-well plates. 15. Carefully transfer the plates into the incubator and incubate the plate for 1 min. 16. With a single swift movement, turn the plate upside down and incubate the plate for 15 min at 37 °C. 17. Pre-warm 4 mL of EVT-OM at 37 °C in the water bath. 18. After 15 min, add 0.5 mL EVT-OM per well. Ensure that the domes are covered with medium. 19. Monitor the culture regularly. 20. Change the medium every 2–4 days. 21. After 2–4 days, an EVT outgrowth will be visible. 22. After 4–5 days, prewarm wash-OM and EVT-DM-2. 23. Carefully aspirate and discard the supernatant from the organoid wells. 24. Add 0.5 ml wash-OM to each organoid well and incubate the organoids in the incubator for 1 h at 37 °C. 25. Aspirate and discard the wash-OM. 26. Add 0.5 mL EVT-DM-2 to each organoid well. 27. Change the medium every 2–4 days. 28. Culture the organoids in EVT-DM-2 for another 4–5 days.

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3.6.2 EVT Induction of TB-ORG

Starting material: 4-well plate of TB-ORG. 1. Pre-warm 2 mL wash-OM and 2 ml EVT-DM. 2. Place the TB-ORG 4-well plate into the biosafety cabinet. 3. Remove and discard the OM from the cultures. 4. Add 0.5 mL wash-OM per TB-ORG well and put the plate back into the incubator. 5. Incubate the plate for 1 h at 37 °C (see Note 15). 6. Remove and discard the wash-OM. 7. Add 0.5 mL of EVT-DM per well. 8. Monitor the culture regularly. 9. Change the medium every 2–4 days. 10. After 2–4 days, an EVT outgrowth should be visible. Culture for up to 10 days. 11. After 4–5 days, prewarm wash-OM and EVT-DM-2. 12. Carefully aspirate and discard the supernatant from the organoid wells. 13. Add 0.5 mL wash-OM to each organoid well and incubate the organoids in the incubator for 1 h at 37 °C. 14. Aspirate and discard the wash-OM. 15. Add 0.5 ml EVT-DM-2 to each organoid well. 16. Change the medium every 2–4 days. 17. Culture the organoids in EVT-DM-2 for another 4–5 days.

3.6.3 Isolation of Trophoblast Subtypes from TB-ORG

Since TB-ORG consist of various trophoblast subpopulations (vCT, ST, and EVTs after differentiation), it may be useful to enrich respective populations for subsequent analyses. Below, a protocol to enrich HLA-Gpos EVTs after differentiation experiments is described. Start with at least 12 wells (three 4-well plates) of TBORG-EVT cultures differentiated for 10 days. 1. Place a small ice bucket with crushed ice into the biosafety cabinet. 2. Place sterile 1.5 mL, conical 15 mL, and conical 50 mL tubes, as well as PBS, FBS, and MACS buffer on ice. 3. Remove and discard the EVT-DM. Add 0.5 mL CRS per well. 4. Place the plate into a sterile plastic bag and transfer the plate to a fridge for 35–45 min in order to dissolve the Matrigel. 5. Gently pipette the TB-ORG/CRS mixture up and down 4–6 times and collect the solution in the pre-cooled 1.5 mL tube. 6. Pellet the TB-ORG at 1.500 rpm (~160 × g) for 5 min at 4 °C. 7. Remove and discard the CRS.

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8. Add 4 mL TrypLE to the TB-ORG. Vortex the tube briefly and incubate it at 37 °C for 10 min in a water bath. 9. Briefly vortex the tube every 2 min. 10. Place a 70 μm cell strainer on top of a conical tube. 11. Remove the 15 mL tube from the water bath and pipette the TB-ORG up and down 10–15 times until the solution appears almost homogeneous. 12. Filter the TB-ORG/TrypLE mixture through the cell strainer. 13. Add cold PBS to a total volume of 10 mL. Invert the tube 3–5 times. 14. Pellet the cells at 1.500 rpm (~160 × g) for 5 min at 4 °C. 15. Resuspend the cell pellet with 1 mL cold MACS buffer. Transfer the solution into the pre-cooled 1.5 mL tube. 16. Take a small amount and count the cells. 17. Pellet the cells at 1.500 rpm (~160 × g) for 5 min at 4 °C. The following protocol describes the purification of HLA-Gpos cells from 2 × 106 cells using magnetic separation and MS columns according to the manufacturer’s instructions (see Note 16). 18. Resuspend the pellet with 90 μL cold MACS buffer. 19. Add 10 μL PE-conjugated HLA-G antibodies and incubate the tube on a roller mixer for 30 min at 4 °C. 20. Add 1 mL MACS buffer. Invert the tube 3–5 times. Pellet the cells (1.500 rpm (~160 × g), 5 min, 4 °C). 21. Resuspend the cells with 80 μL of cold MACS buffer. Add 20 μL of PE-labeled magnetic beads. Incubate the tube for 20 min on a roll mixer at 4 °C. 22. Add 1 mL cold MACS buffer. Invert the tube 3–5 times. Pellet the cells (1.500 rpm (~160 × g), 5 min, 4 °C). 23. Place a 70 μm cell strainer on top of a 50 mL conical tube. 24. Resuspend the cell pellets with 0.5 mL MACS buffer and filter the solution through the cell strainer. 25. Place one MACS separation column into the magnetic holder. 26. Equilibrate the column with 1 mL cold MACS buffer. Avoid air bubbles. 27. Wait until the column reservoir is empty. 28. Place 15 mL conical tubes underneath the column to collect the flow-through. 29. Add 0.5 mL filtered cell solution into the column reservoir. HLA-Gpos cells will be kept in the column while HLA-Gneg cells will flow into the 15 mL tube. 30. Wait until the column reservoir is empty.

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31. Store the tube with HLA-Gneg cells for later analyses. 32. Wash the column 3 times with 0.5 mL cold MACS buffer. Avoid air bubbles. Wait until the column reservoir is empty every time. 33. Remove the column from the magnetic holder and put it on top of a fresh 15 mL conical tube. 34. Add 2 mL of MACS buffer onto the column reservoir and press the liquid through the column. The solution will contain the HLA-Gpos EVTs. 35. Count the number HLA-Gpos EVTs.

of

both

HLA-Gneg

cells

and

36. Proceed with downstream analyses, or pellet and freeze the cells for later analyses. 3.7 Molecular and Histological Analysis of TB-ORG

TB-ORG can either be lysed to obtain RNA, DNA, and protein preparations, or fixed and embedded in paraffin for spatial localization of proteins. Additionally, the supernatant can be stored and investigated for secreted proteins.

3.7.1 TB-ORG Lysis for RNA and Protein Preparation

It is recommended to remove the Matrigel prior to lysis since this ensures that the proteins present in the Matrigel do not hamper the analysis interpretation (see Note 17). The collection of two TB-ORG domes after Matrigel removal is described. 1. Plate a small ice bucket with crushed ice into or nearby the biosafety cabinet. 2. Place 1× PBS and several 1.5 mL tubes on ice. 3. Collection and storage of the organoid supernatant: Remove and collect the medium in one 1.5 mL tube. Centrifuge the supernatant at 2.000 rpm (~280 × g) for 10 min at 4 °C to remove cell debris. Pipette the supernatant into a fresh 1.5 mL vial and store it at -80 °C. 4. Add 0.5 mL CRS per TB-ORG well. 5. Place the plate into a sterile bag and transfer the plate into the fridge for 35–45 min in order to dissolve the Matrigel. 6. Gently pipette the TB-ORG/CRS mixture up and down 4–6 times and collect the solution in the pre-cooled 1.5 mL vial. 7. Pellet the TB-ORG at 1.500 rpm (~160 × g) for 5 min at 4 °C. 8. A TB-ORG pellet should be clearly visible. Remove and discard the CRS. 9. Add an appropriate RNA or protein lysis buffer to the tubes. It is recommended to use TriFast (PeqLab) for RNA and subsequent DNA and protein isolations according to the manufacturer’s instructions.

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Fig. 6 Fixation and embedding of TB-ORG into paraffin. Matrigel domes containing TB-ORG are fixed in formaldehyde (FA), washed with 70% EtOH, and incubated with alcian blue-containing 70% EtOH to ensure their visibility. Following dehydration and xylene incubation, the TB-ORG are transferred into small metal molds containing 65 °C liquid paraffin and incubated on a hot plate. After cooling, the paraffin blocks with embedded TB-ORG (light blue) are ready to be sectioned for the downstream immunodetection analysis

10. OPTIONAL: The TB-ORG lysis might be supported by mechanical force. It is recommended to use the Precellys tissue homogenizing CKMix tubes. Pipette the TB-ORG/lysis buffer mixture into the CKMix tubes and mix twice for 20 s each using the Precellys 24 (see Note 18). 11. Proceed with the RNA and protein lysis protocols according to the respective manufacturer’s instructions. 12. Alternatively, the TB-ORG pellets (see step 9) can be frozen and stored at -80 °C. 3.7.2 Preparation of Paraffin-embedded TBORG Tissue Blocks

The main difficulty in embedding the trophoblast organoids is their small size (Fig. 6)) (see Note 19). 1. Dilute 7.5% FA with 1× PBS to 4%. Add 1 mL into a 1.5 mL tube. 2. Prepare 70%, 96%, and 100% EtOH, Xylene, and liquid paraffin. 3. Take a spatula and transfer the whole TB-ORG-containing dome into the 4% FA solution.

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4. Fix the TB-ORG for at least for 1 h (up to 24 h) at 4 °C (see Note 20). 5. Quickly spin the tube down in a microcentrifuge and carefully remove the 4% FA solution. 6. Wash the TB-ORG with 1× PBS: Add 1 mL 1× PBS. Invert the tube 3–5 times. Short-spin the TB-ORG down and remove the PBS. Repeat 3 times. Ensure that the FA is completely removed (see Note 21). 7. Add 1 mL 70% EtOH. Invert the tube 3–5 times. Short-spin the TB-ORG down and remove the EtOH. 8. Mix 10 μL of alcian blue with 1 mL of fresh 70% EtOH. 9. Add the 70% EtOH/alcian blue mixture to the TB-ORG. Invert the tube 3–5 times and incubate the TB-ORG for 30 min at room temperature (see Note 22). 10. Short-spin the tube and replace the 70% EtOH with 96% EtOH. Invert the tube 3–5 times. Incubate the TB-ORG for 10 min at room temperature. Repeat the step once with fresh 96% EtOH. 11. Short-spin the tube and replace the 96% EtOH with 100% EtOH. Invert the tube 3–5 times. Incubate the TB-ORG for 10 min at room temperature. Repeat the step once with fresh 100% EtOH. 12. Short-spin the tube and replace the 100% EtOH with xylene. Invert the tube 3–5 times. Incubate the TB-ORG for 30 min at room temperature. Repeat the step once with fresh xylene. 13. Short-spin the tube and remove as much xylene as possible. 14. Pour hot liquid paraffin wax at 65 °C into small metal base molds. 15. Using heat-safe forceps, transfer the TB-ORG from the tubes into the molds, and incubate them for 30 min at 65 °C on the appropriate heating plate. 16. Carefully remove the paraffin wax and pour fresh, hot wax onto the TB-ORG-containing molds. Perform another incubation step for 30 min at 65 °C. 17. Ensure that the TB-ORG are gathered in the center of the metal molds. Carefully move the mold to the cold plate for 5–10 sec. This fixes the TB-ORG in the position. 18. Place plastic histo-cassettes on top and fill the whole metal mold with hot paraffin. 19. Transfer the device/metal mold to a cold plate to allow solidification. 20. After 30 min, remove the solid TB-ORG-paraffin block from the metal mold. 21. Store the paraffin samples at room temperature.

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Notes 1. In placental tissue of later gestational weeks (>8-week placenta), stromal cell contamination may occur more frequently. Hence, we recommend a stromal cell removal step at the end of the vCT isolation procedure, precisely after erythrocyte removal in Subheading 3.2.4, step 3. (a) Resuspend the cells in 10 mL pre-warmed advanced DMEM/F12 medium containing 10% FBS. (b) Plate the cell suspension on a 10 cm culture dish. Incubate for 45 min at 37 °C. (c) Collect the non-adherent cells (i.e., vCT) and proceed with erythrocyte removal in Subheading 3.2.4, step 4. 2. In our experience, a 1–2 mL tissue pellet requires 10 mL of digestion solution, while a 2–5 mL tissue pellet requires 20 mL of digestion solution per digestion step. 3. Avoid rapid movements and vibrations when handling the gradient. 4. During the centrifugation step, rapid acceleration/deceleration must be reduced to a minimum. Please note, that despite the brake being set to 0, some centrifuges decelerate rather abruptly which could reduce the cell separation efficiency. This should be checked in advance. 5. Move the plastic pipette in slow circles along the tube wall while carefully moving downwards and aspirating the liquid. 6. We recommend preparing eight domes at 40 μL cell/vCTOM/Matrigel mixture because (i) eight domes allow for larger subsequent experimental scope as well as back-up freezing of established cultures and (ii) larger volumes of medium and Matrigel are easier to pipette. 7. Matrigel quickly starts to solidify at temperatures above 4 °C which makes pipetting more difficult and might result in partial solidifying of Matrigel in the pipette tip. Hence, we strongly recommend cooling all vials and solutions on ice prior to use. 8. The ratio of 40:60 cell/OM:Matrigel needs to be accurately maintained. An imbalance (i.e., excess cell/OM in relation to Matrigel) reduces the dome stiffness, which results in its loss. 9. Once air bubbles occur, they can hardly be removed. Therefore, we recommend avoiding dispensing the total volume out of the pipette tip to reduce the risk of air bubble formation when mixing the cells/OM with Matrigel. 10. A TB-ORG pellet should be clearly visible. If not, pipette the whole solution up and down, and make sure to rinse the walls of the tube. Repeat the centrifugation and remove the CRS.

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11. Ensure that the domes do not dry out, not even partially. 12. In order to avoid rapid cooling of the pre-warmed culture dish, place it on top of a plastic tube rack in the biosafety cabinet. 13. Reducing the initial incubation time at 37 °C in the upright position from 1–2 min to 10–20 s would improve the outcome. Alternatively, generation of small “beds” of Matrigel is beneficial. To generate small “beds” of Matrigel, do the following: (a) Place a small ice bucket with crushed ice into the biosafety cabinet. (b) Place a sterile 1.5 mL tube, Matrigel, and OM on ice. (c) Prepare 120 μL of a 40:60 OM:Matrigel solution. Mix 48 μL OM with 72 μL Matrigel. (d) Pipette 7 μL of the mixture onto the center of one well. Carefully distribute the drop with the pipette tip to the approximate diameter of the later TB-ORG dome. (e) Perform the same procedure for the remaining 15 wells. (f) Return the plate to the incubator for at least 15 min prior seeding the new TB-ORG cultures. 14. Several manufacturers leave the decision to the researcher if long-term storage will be done at -80 °C or in liquid nitrogen. However, we observed a higher viability when cells and TB-ORG are transferred to liquid nitrogen no later than 2–3 weeks. 15. The 1 h incubation will ensure the complete removal of CHIR99021, which will support EVT differentiation. 16. The reagent amount and columns required for PE-beaddependent cell separation need to be adapted depending on the cell number. We recommend following the manufacturer’s protocols. 17. The whole TB-ORG-containing domes, including the Matrigel, can be lysed for subsequent RNA and protein preparation. However, the obtained protein mixture will originate both from TB-ORG and Matrigel. 18. There are different tube variants available. We recommend the CKM for a volume of up to 2 mL. However, if the volume does not exceed 0.3 mL, alternative tubes can be used. 19. In routinely used tissue-embedding cassettes, even when using filter strips, hardly any TB-ORG would remain to be embedded in paraffin. Therefore, they need to be processed manually. 20. Some antigens are sensitive to prolonged fixation time. Thus, the fixation time needs to be adjusted appropriately.

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21. Incomplete removal of FA will result in the formation of small precipitates in the subsequent 70% EtOH/alcian blue step. These precipitates are hardly distinguishable from TB-ORG and complicate the paraffin-embedding procedure. 22. The alcian blue will give the TB-ORG a light blue color helping to enhance the visibility of the small TB-ORG structures.

Acknowledgments This work was supported by the Austrian Science Fund (grant P-34588-B awarded to SH and grant P-31738-B26 awarded to PL). We are grateful to Gudrun Meinhardt and Henrieta Papuchova for valuable comments. References 1. Hemberger M, Hanna CW, Dean W (2020) Mechanisms of early placental development in mouse and humans. Nat Rev Genet 21:27–43. https://doi.org/10.1038/s41576-0190169-4 2. Turco MY, Moffett A (2019) Development of the human placenta. Development 146: dev163428. https://doi.org/10.1242/dev. 163428 3. Kno¨fler M, Haider S, Saleh L et al (2019) Human placenta and trophoblast development: key molecular mechanisms and model systems. Cell Mol Life Sci 76:3479–3496. https://doi.org/10.1007/s00018-01903104-6 4. Okae H, Toh H, Sato T et al (2018) Derivation of human trophoblast stem cells. Cell Stem Cell 22:50–63.e6. https://doi.org/10.1016/ j.stem.2017.11.004 5. Haider S, Meinhardt G, Saleh L et al (2018) Self-renewing trophoblast organoids recapitulate the developmental program of the early human placenta. Stem Cell Rep. https://doi. org/10.1016/j.stemcr.2018.07.004 6. Turco MY, Gardner L, Kay RG et al (2018) Trophoblast organoids as a model for maternal-fetal interactions during human placentation. Nature 564:263–267. https://doi. org/10.1038/s41586-018-0753-3 7. Lancaster MA, Huch M (2019) Disease modelling in human organoids. Dis Model Mech 12. https://doi.org/10.1242/dmm.039347 8. Schutgens F, Clevers H (2020) Human organoids: tools for understanding biology and treating diseases. Annu Rev Pathol 15:211– 234. https://doi.org/10.1146/annurev-path mechdis-012419-032611

9. Kim J, Koo B-K, Knoblich JA (2020) Human organoids: model systems for human biology and medicine. Nat Rev Mol Cell Biol 21:571– 584. https://doi.org/10.1038/s41580-0200259-3 10. Saha B, Ganguly A, Home P et al (2020) TEAD4 ensures postimplantation development by promoting trophoblast self-renewal: an implication in early human pregnancy loss. Proc Natl Acad Sci U S A 117:17864–17875. https://doi.org/10.1073/pnas.2002449117 11. Meinhardt G, Haider S, Kunihs V et al (2020) Pivotal role of the transcriptional co-activator YAP in trophoblast stemness of the developing human placenta. Proc Natl Acad Sci U S A 117: 13562–13570. https://doi.org/10.1073/ pnas.2002630117 12. Hornbachner R, Lackner A, Haider S et al (2021) MSX2 safeguards syncytiotrophoblast fate of human trophoblast stem cells. Proc Natl Acad Sci U S A 118(37):e2105130118 13. Kliman HJ, Nestler JE, Sermasi E et al (1986) Purification, characterization, and in vitro differentiation of cytotrophoblasts from human term placentae. Endocrinology 118:1567– 1582. https://doi.org/10.1210/endo-1184-1567 14. Tarrade A, Lai Kuen R, Malassine´ A et al (2001) Characterization of human villous and extravillous trophoblasts isolated from first trimester placenta. Lab Investig 81:1199–1211. https://doi.org/10.1038/labinvest.3780334 15. Sandra, Haider Andreas Ian, Lackner Bianca et al (2022) Transforming growth factor-β signaling governs the differentiation program of extravillous trophoblasts in the developing human placenta Proceedings of the National

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Academy of Sciences 119(28). https://doi. org/10.1073/pnas.2120667119 16. Kaela M, Varberg Esteban M, Dominguez Boryana et al (2023) Extravillous trophoblast cell lineage development is associated with active remodeling of the chromatin landscape Abstract Nature Communications 14(1). https://doi.org/10.1038/s41467023-40424-5 17. Chen, Dong Shuhua, Fu Rowan M et al (2022) A genome-wide CRISPR-Cas9 knockout screen identifies essential and growthrestricting genes in human trophoblast stem cells Abstract Nature Communications 13(1). https://doi.org/10.1038/s41467022-30207-9 18. Mariyan J, Jeyarajah Gargi, Jaju Bhattad Rachel D et al (2022) The multifaceted role of GCM1 during trophoblast differentiation in the human placenta Proceedings of the National

Academy of Sciences 119(49). https://doi. org/10.1073/pnas.2203071119 19. Kaela M, Varberg Khursheed, Iqbal Masanaga et al (2021) ASCL2 reciprocally controls key trophoblast lineage decisions during hemochorial placenta development Significance Proceedings of the National Academy of Sciences 118(10). https://doi.org/10.1073/pnas. 2016517118 20. Liheng, Yang Eleanor C, Semmes Cristian et al (2022) Innate immune signaling in trophoblast and decidua organoids defines differential antiviral defenses at the maternal-fetal interface eLife. https://doi.org/1110.7554/ eLife.79794 21. (2024) Trophoblast organoids with physiological polarity model placental structure and function ABSTRACT Journal of Cell Science 137(5). https://doi.org/10.1242/jcs.261528

Chapter 18 A Three-Dimensional Trophoblast Invasion Microfluidic Platform for Toxicological Screening Yong Pu and Almudena Veiga-Lopez Abstract To improve our understanding of human placental function and placental cell responses to pregnancy stressors, the development of in vitro models that better recapitulate the in vivo placental microenvironment is needed. Here, we describe a three-dimensional (3D) silicone polymer polydimethylsiloxane (PDMS) microfluidic platform for modeling human trophoblast invasion recreating a placental heterocellular microenvironment. This platform allows the formation of a cellular barrier establishing a chemical gradient and real-time evaluation of trophoblast cell invasion and heterocellular cell-to-cell interactions. Key words Three-dimensional, Microfluidics, Trophoblast, Invasion

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Introduction The human placenta is unique structurally and functionally [1] and thus no animal model (except for some non-human primates) is deemed to be a representative model for human placentation. The ethical concerns and the difficulty of accessing human pregnancies, as well as the structural complexity associated with the placenta, have hampered our understanding of human placental structure and biology and the development of suitable in vitro models to evaluate trophoblast function. The placenta harbors different types of trophoblast cells, one of which are extravillous trophoblasts (EVTs). EVTs are invasive trophoblasts that proliferate and migrate toward the maternal endometrium invading into the decidual layer. This process is required for proper conceptus attachment and placenta development. Throughout pregnancy, EVTs continue to invade and provide support to uterine spiral artery remodeling, a key event in human placental development. Defects of EVT invasiveness result in deficient spiral artery remodeling, which is associated with abnormal placentation and pregnancy outcomes [2–4].

Sandeep Raha (ed.), Trophoblasts: Methods and Protocols, Methods in Molecular Biology, vol. 2728, https://doi.org/10.1007/978-1-0716-3495-0_18, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Classic in vitro models used to test EVT migration and invasion include the scratch [5] or the transwell assay [6]. More recent, three-dimensional (3D) invasion models, such as the spheroid gel invasion assay, spheroid confrontation assay model, spheroid/ monodispersed cell invasion assay model, vertical gel invasion assay model, and 3D cell tracking assay model have been developed [7–9]. Abbas et al. [10] incorporated the use of continuous flow and shear stress to the study of trophoblast cell invasion by using microfluidics, a state-of-the-art technology that allows for media flow to bathe the cell culture device. In the trophoblast invasion model presented here, we have recapitulated a heterocellular placental microenvironment using a 3D microfluidic platform. This system allowed us to (1) establish an endothelial barrier to allow for an active diffusion gradient, (2) test trophoblast cell invasion into an endothelial cellular compartment, and (3) test trophoblastendothelial cell interactions.

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Materials Prepare all solutions with ultrapure sterile water. Work under sterile conditions (see Note 1).

2.1

Instruments

1. Microfluidic chip (Synvivo 102008; Fig. 1). 2. Vacuum desiccator. 3. Syringe pump (Harvard Apparatus 70-3007). 4. Slide clamps (SynVivo 202001). 5. Tygon tubing (internal diameter 0.08636 cm, outer diameter 0.13716 cm). 6. 1 mL syringe with Luer-Lok tip. 7. 24-gauge blunt-tipped needles. 8. Disposable Petri dish (100 mm). 9. Metal tweezers. 10. Metal scissors. 11. Inverted fluorescence microscope. 12. CO2 cell culture incubator. 13. 15 mL tubes. 14. 0.22 μm filters.

2.2

Reagents

1. 1× DPBS without Ca2+ and Mg2+. 2. Fibronectin (ThermoFisher 33016015). Dissolved in 1× DPBS to a final concentration of 200 μg/mL (store at -20 °C). 3. Human umbilical vein endothelial cells (Lonza CC-2935).

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Fig. 1 Basic instruments. (a) Silicone polymer polydimethylsiloxane (PDMS) 3D microfluidic chip. (b) 3D microfluidic chip scheme depicting the following components: (1) a central compartment (red) with a central feeder line supplied by two inlet ports (i1, i2) connected to two outlet ports (o1, o2) and (2) two outer channels (blue) with two outer feeder lines supplied by two inlet ports (i3, i4) connected to two outlet ports (o3, o4). The barrier between the central compartment and the outer channels is filled with pillars (pillar spacing (PS): 3 μm; total width: 50 μm). (c) Utensils needed: Scissors, forceps, 1 mL syringe, blunted needle hub, clamp (blue), tubing cuttings, and microfluidic chip. (d) Syringe pump

4. First-trimester human placenta HTR8/SVneo trophoblast cells (ATCC CRL-3271). 5. Endothelial cells complete medium. • EGM-2 basal media (Lonza CC-3156). • EGM-2 supplement kit (Lonza CC-4176). 6. HTR8/SVneo growth medium. • DMEM/F12. • 10% fetal bovine serum. • 10 mM HEPES. • 1% penicillin-streptomycin. 7. Trypsin-EDTA. 8. 10% formalin.

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Methods Chip Priming

1. Place the microfluidic chip in a Petri dish inside the vacuum desiccator (Fig. 2). The vacuum desiccator should be connected to a vacuum line. 2. Immerse the chip in sterile 1× DPBS (see Note 2). 3. Turn on the vacuum pump and allow the vacuum to run overnight. 4. Turn off the vacuum pump, release the vacuum, and remove the Petri dish from the desiccator while keeping the chip immersed in DPBS. 5. Cut 5-cm-long pieces of Tygon tubing (Fig. 1c) (see Note 3). 6. Insert the Tygon tubing pieces into one inlet pore (i1) of the central compartment, and all four outlet pores (o1, o2, o3, o4) (Fig. 1b for pore details and Fig. 2d for final configuration). 7. Ensure that there are no air bubbles in any of the compartments of the chip using a microscope (4× magnification) (Fig. 3c, d) (see Note 4). 8. The device is now perfused with DPBS and ready for fibronectin coating (Fig. 2d) (see Note 5).

3.2 Fibronectin Coating

1. Thaw fibronectin (see Note 6). 2. Dissolve fibronectin in sterile 1× DPBS to a final concentration of 200 μg/mL and pass through a 0.22 μm filter. 3. Connect a needle to a 1 mL syringe and draw up the fibronectin solution (see Note 7). 4. Connect an 20.3-cm-long tubing to the syringe needle. 5. To coat the central compartment, place a drop of 1× DPBS on top of central compartment inlet port (i2; Figs. 1b and 4a). Make sure that a drop of fibronectin comes out of the tubing (asterisk, Fig. 4b) before pushing the tubing into the port of the chip. Connect as shown in Fig. 4c. Once connected, slowly push the fibronectin solution from the syringe into the tubing to fill the chip. The fibronectin solution will come out of the inlet port (i1) first (asterisk, Fig. 5a). Clamp the tubing placed on inlet port (i1) one-third into the tubing (Fig. 5b, arrow). 6. Continue to fill the chip with the fibronectin solution, which will now come out through the outlet port(s) (o1 and o2; asterisks; Fig. 5b). Clamp the tubing from the outlet(s) ports (o1 and o2) one-third into the tubing (Fig. 5c, arrow) (see Note 8).

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Fig. 2 Microfluidic chip priming. (a) Microfluidic chip onto Petri dish before priming. (b) Desiccator containing microfluidic chip bathed in DPBS in a Petri dish. (c) Desiccator connected to vacuum line. (d) Microfluidic chip onto Petri dish ready to be coated with fibronectin

Fig. 3 Checking cell seeding integrity under the microscope. (a) Microfluidic chip after cell seeding. Note that only the central compartment and the bottom outer channel have been seeded with cells. (b) Microfluidic chip after cell attachment. The central compartment and both outer channels have been seeded with cells. (c, d) Microfluidic chip with various air bubbles (white arrows)

Fig. 4 Fibronectin coating (part 1). (a) Microfluidic chip with a DPBS drop over i1 port for fibronectin coating. (b, c) Connecting of the fibronectin solution tubing with i1 port. (d) Cut the tubing connected to i1 port. (e) Connecting of the fibronectin solution tubing with i4 port. (f) Connecting of the fibronectin solution tubing with i3 port. Asterisk denotes drop at the end of tubing to be connected

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Fig. 5 Fibronectin coating (part 2). (a) Fibronectin solution from i2 port comes out from i1 port. (b) i1 port is then clamped and fibronectin solution comes out through o1 and o2 ports. (c) o1, o2, and i2 ports are then clamped and fibronectin solution from i4 port comes out o4 port. (d) After clamping o4 and i4 port, the fibronectin solution from i3 port comes out of port o3. (e, f) Outlook of the chip after clamping of the remainder of the ports

7. Cut tubing of the inlet port i2 to ~2-inch of length (Fig. 4d) and clamp the inlet port (i2) tubing on one-third into the tubing (Fig. 5c). 8. To coat the outer channels, connect the Tygon tubing to inlet port i4 (Figs. 1b and 4e), flush the fibronectin into the chip, until a drop comes out of o4 (Fig. 5c). Then, clamp and cut the excess tubing. Repeat this same process for inlet port i3 (Figs. 4f and 5d). 9. All ports should now be clamped and should be kept clamped (Fig. 5e, f) unless otherwise stated. 10. Keep chip on Petri dish and place inside the vacuum desiccator overnight at 4 °C. 3.3

Cell Seeding

1. At least 2 days before chip cell seeding, seed HUVECs and HTR8/SVneo cells into separate 100 mm dishes (see Note 9). 2. Remove the chip from the vacuum desiccator and place at room temperature. Carefully, wipe the excess humidity off the chip with a Kimwipe. 3. Let the chip warm up to room temperature for 30 min. 4. Trypsin digest and harvest the cultured HUVECs and HTR8/ SVneo cells and adjust each cell density to 30 million cells per mL. 5. To seed the cells into the central compartment, draw up the HUVECs single-cell suspension using a 1 mL syringe

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connected to a needle. Connect the needle to a 4-inch-long tubing. Push the syringe until a drop of the single-cell suspension appears on the tip of the tubing. 6. Place a drop of 1× DPBS on top of the inlet port (i2) (see Note 10). 7. Pull i2 tubing out and unclamp outlet ports (o1 and o2). 8. Connect syringe tubing containing cell suspension with inlet port i2 and gently push the cell suspension into the chip allowing 1–2 drops out of the outlet ports o1 and o2 (see Note 11). 9. Clamp tubing of ports o1 and o2 followed by i2. Cut tubing of inlet port i2 to ~2-inch of length. 10. To seed the cells into the outer channel (i4, o4), draw up the HTR8/SVneo single-cell suspension using a 1 mL syringe connected to a needle. Connect the needle to a 15-cm-long tubing. Push the syringe until a drop of the single-cell suspension appears on the tip of the tubing. 11. Place a drop of 1× DPBS on top of the inlet port (i4) (see Note 12). 12. Pull i4 tubing out and unclamp outlet port (o4). 13. Connect syringe tubing containing cell suspension with inlet port i4 and gently push the cell suspension into the chip allowing 1–2 drops out of the outlet port o4 (see Note 13). 14. Clamp tubing of port o4 followed by i4. Cut tubing of inlet port i4 to ~2-inch of length. 15. Repeat Steps 10–14 for outer channel i3, o3. 16. Use a Kimwipe to remove the excess 1× DPBS off the chip. 17. Place the chip into the CO2 incubator to allow for cell attachment (~4–6 h). 3.4

Media Flow

1. After a 4–6 h incubation, check under the microscope to confirm that cells have been attached (Fig. 3b). 2. Prepare three 1 mL syringes filled with endothelial cell complete medium (1 syringe) and HTR8/SVneo growth medium (2 syringes) (see Note 14). 3. Connect each syringe to a ~50.8-cm tubing (see Note 15). 4. To connect media syringe to the central compartment, place a drop of 1× DPBS on top of the inlet port (i2), and unclamp the outlet pore tubing (o1 and o2). 5. Pull out of the inlet pore tubing (i2). Connect syringe tubing containing cell media with inlet port i2 and gently push the medium into the chip allowing 1–2 drops out of the outlet

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Fig. 6 Assembled microfluidic platform. (a) Pump loaded with three cell medium syringes connected to the chip (1 central compartment and 2 outer channels) and outflow of media connected to collecting 0.75 mL tubes. (b) A magnified view of the detail of the chip and collecting tubes. (c) Microfluidic chip under fluorescence microscope

ports o1 and o2 to ensure all cells are bathed with the medium (see Note 16). 6. To capture the medium effluent, insert the outlet pore tubing into a 0.75 mL perforated tube (Fig. 6). The lid of the tube can be perforated using a fine pair of scissors. 7. To connect media syringe to the outer channels, place a drop of 1× DPBS on top of the inlet port (i3), and unclamp the outlet pore tubing (o3). 8. Connect syringe tubing containing cell media with inlet port i3 and gently push the medium into the chip allowing 1–2 drops out of the outlet ports o3 to ensure all cells are bathed with the medium. 9. Repeat steps 7 and 8 for channel i4, o4. 10. Place the chip into the CO2 incubator and place all three syringes onto the syringe pump (Fig. 6). 11. To avoid any tubing stretches, use a piece of tape to secure the tubing onto the side of the CO2 incubator. 12. Program the syringe pump as follows (see Note 17): • Infuse mode: constant rate. • Syringe: BD 1 mL (diameter to 4.699 mm).

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• Flow rate: 0.01 μL/min. • Infusion time: 72:00:00 (72 h). 13. Start the syringe pump program. 14. Incubate the chip overnight at 37 °C and 5% CO2. 15. Visualize cells under the microscope every 24 h. 3.5 Real-time Imaging

1. After 24 h incubation (with medium flow), pause the syringe pump program and take the chip out of the incubator, near the syringe pump. 2. Carefully take the syringe off the syringe pump and put the chip under the fluorescence microscope for imaging (Fig. 6c) (see Note 18). 3. Use a 4× objective to focus and then image at higher magnification as needed (see Note 19). 4. After imaging, place the chip back into the CO2 incubator and place the syringes onto the syringe pump holder and resume the syringe pump program.

3.6 Antibody Staining and Imaging

1. Connect the needle with a 1 mL syringe, draw up 10% formalin into a 1 mL syringe, then connect the needle to a 20.3 cm long tubing (see Note 20). 2. Place a drop of 1× DPBS on top of the tubing for the inlet port (i2). 3. Connect syringe tubing containing 10% formalin with inlet port i2 and gently flush the medium into the chip allowing 1–2 drops out of the outlet ports o1 and o2. 4. Repeat steps 2 and 3 for both outer channels. 5. Incubate the chip at room temperature for 20 min. 6. Wash all channels of the chip with a 1 mL syringe filled with 1× DPBS as in step 3. 7. Follow your immunofluorescence staining protocol (primary antibody, blocking reagent, secondary antibody) to complete the staining. Then clamp all tubing and proceed to imaging. 8. For long-term storage, the chip can be kept inside a humid chamber at 4 °C. A humid chamber can be created by placing DPBS-soaked Kimwipes inside the Petri dish and sealing the dish with parafilm (see Note 21).

3.7

Cell Harvesting

1. To harvest cells, pause the syringe pump program, take the chip out of the incubator, and place it near the syringe pump. Then, cut all the tubing off from their corresponding syringes. The cut should be ~5 cm from the ports.

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2. To connect the trypsin syringe to the central compartment, place a drop of 1× DPBS on top of the inlet port (i2). 3. Connect syringe tubing containing trypsin with inlet port i2 and gently flush the trypsin into the chip allowing 1–2 drops out of the outlet ports o1 and o2. Then, cut the tubing ~5 cm from the ports. 4. Repeat steps 2 and 3 for channels i3, o3 and i4, o4. 5. Place the chip into the CO2 incubator for 3–5 min. 6. Connect a 1 mL syringe with a needle and draw up pre-warmed DMEM/F12 (with 10% serum) into the syringe. 7. Place a drop of 1× DPBS on top of the inlet port (i2). 8. Pull out of the inlet pore tubing (i2) and connect syringe tubing containing growth medium with inlet port (i2) and gently flush the medium into the chip and harvest media coming off the outlet ports o1 and o2 using a 0.75 mL tube. The trypsinized cells should be washed with the medium (see Note 22). 9. Clamp all inlet and outlet tubing of the central compartment (i1, i2, o1, o2). 10. To harvest cells from the outer channels, repeat steps 2–9 with channels i3, o3 and i4, o4. 11. Centrifuge the 0.75 mL tubes (300 × g 5 min). Cells can be used for further culture or stored as frozen cell pellet for further use.

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Notes 1. Any manipulation of the chip that includes opening and closing ports should be done under sterile conditions in a biosafety cabinet. 2. The chip should be fully immersed in DPBS. Failing to do so will result in incomplete priming. 3. The cut of the Tygon tubing should be as flat (90° cut, not angled) as possible. This will help a clean fit of the tubing into the chip. 4. Air bubbles will result in poor extracellular matrix coating and cell attachment. If air bubbles are present, repeat the process of chip priming (step 1 of 3.1, Chip Priming) until no air bubbles are present in the chip (Fig. 3c, d). 5. The primed chip needs to be coated on the same day. A delayed coating may result in air bubbles inside the chip. 6. Avoid shaking thawed fibronectin, as this can cause clumps and misalignment of fibers.

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7. Make sure no air bubbles are present in the syringe. Air bubbles in the syringe may lead to air bubbles inside the chip. 8. Before clamping the tubing from the outlet ports o1 and o2, place the chip under the microscope to check whether any air bubbles have been formed (Fig. 3c, d). If yes, keep flushing with fibronectin solution until no air bubbles are present. 9. Cell culture details can be found in our previous study [11]. Do not allow cells reach 100% confluency, as cells would have better attachment and proliferation ability during the logarithmic phase. 10. This step is critical as failing to add the droplet may lead to air bubbles inside the chip. 11. Cell density in the central compartment can now be checked under the microscope. If cells are not well distributed across the compartment, continue flushing the cell suspension until a uniform distribution is achieved (Fig. 3a). 12. This step is critical as failing to add the droplet may lead to air bubbles inside the chip (Fig. 3c, d). 13. Similar to the central compartment, cell density should be checked for each of the outer channels (Fig. 3a). 14. There should be no air bubbles in the syringes. This is especially critical in the connection area of the syringe and the needle hub. 15. Suggested tube length from the chip to the pump will depend on the distance at which the pump is located from the CO2 incubator. If the pump is placed above the incubator, each tubing section that goes from the chip to the syringe is ~50.8 cm. The chip requires 3 long tubing sections, one for the central compartment inlet pore (i2) and two for the outer channel inlet pores (i3 and i4). 16. The tubes do not need to be clamped in this step as media needs to be able to flow out of the chip. 17. It is important to select the correct syringe type. Otherwise, the flow rate will not be accurate. Infusion conditions (flow rate, time) should be changed as per the experimental needs. 18. This step should be done very carefully. Even slight movements can disrupt the cells within the chip. 19. A 4× objective lens is needed to image the entire chip (all channels). 20. Before staining, it is advisable to capture phase contrast images of the cells to record cellular morphology and density prior to staining.

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21. The PDMS must be kept hydrated or evaporation will lead to air bubbles forming inside the chip or cells drying out. Unclamping the tubing and introducing additional liquid into the chip will hydrate the channels and chambers. 22. In our experience, 200 μL is enough to remove all cells from the chip.

Funding Source This publication was supported by the National Institute of Environmental Health Sciences of the National Institute of Health (R01ES027863 to A.V-L and P30ES027792). References 1. Schmidt A, Morales-Prieto DM, Pastuschek J, Frohlich K, Markert UR (2015) Only humans have human placentas: molecular differences between mice and humans. J Reprod Immunol 108:65–71 2. Lim KH, Zhou Y, Janatpour M, McMaster M, Bass K, Chun SH, Fisher SJ (1997) Human cytotrophoblast differentiation/invasion is abnormal in pre-eclampsia. Am J Pathol 151: 1809–1818 3. Barrientos G, Pussetto M, Rose M, Staff, A. C, Blois SM, Toblli JE (2017) Defective trophoblast invasion underlies fetal growth restriction and preeclampsia-like symptoms in the strokeprone spontaneously hypertensive rat. Mol Hum Reprod 23:509–519 4. Sebire NJ, Fox H, Backos M, Rai R, Paterson C, Regan L (2002) Defective endovascular trophoblast invasion in primary antiphospholipid antibody syndrome-associated early pregnancy failure. Hum Reprod 17: 1067–1071 5. Liang CC, Park AY, Guan JL (2007) In vitro scratch assay: a convenient and inexpensive method for analysis of cell migration in vitro. Nat Protoc 2:329–333 6. Albini A, Iwamoto Y, Kleinman HK, Martin GR, Aaronson SA, Kozlowski JM, McEwan

RN (1987) A rapid in vitro assay for quantitating the invasive potential of tumor cells. Cancer Res 47:3239–3245 7. Zambuto SG, Clancy KBH, Harley BAC (2019) A gelatin hydrogel to study endometrial angiogenesis and trophoblast invasion. Interface Focus 9:20190016 8. Wong MK, Wahed M, Shawky SA, DvorkinGheva A, Raha S (2019) Transcriptomic and functional analyses of 3D placental extravillous trophoblast spheroids. Sci Rep 9:12607 9. Kramer N, Walzl A, Unger C, Rosner M, Krupitza G, Hengstschlager M, Dolznig H (2013) In vitro cell migration and invasion assays. Mutat Res 752:10–24 10. Abbas Y, Oefner CM, Polacheck WJ, Gardner L, Farrell L, Sharkey A, Kamm R, Moffett A, Oyen ML (2017) A microfluidics assay to study invasion of human placental trophoblast cells. J R Soc Interface 14. https:// doi.org/10.1098/rsif.2017.0131 11. Pu Y, Gingrich J, Veiga-Lopez A (2021) A 3-dimensional microfluidic platform for modeling human extravillous trophoblast invasion and toxicological screening. Lab Chip 21(3): 546–557

Chapter 19 Three-Dimensional In Vitro Human Placental Organoids from Mononuclear Villous Trophoblasts or Trophoblast Stem Cells to Understand Trophoblast Dysfunction in Fetal Growth Restriction Cherry Sun, Joanna L. James, and Padma Murthi Abstract The placenta plays a critical role in the efficient delivery of nutrients and oxygen from mother to fetus to maintain normal fetal growth. Human placental development and function is a highly orchestrated process, which is spatially and temporally controlled by hormones and growth factors. Specialized epithelial cells called trophoblasts play key roles in placental exchange capacity, and their abnormal function and development contribute to many pregnancy complications, including fetal growth restriction (FGR), a condition in which the fetus does not reach its full growth potential in utero. Great variation in the anatomy and development of the placenta in animal model systems (in vivo) and 2D culture model systems of trophoblasts (in vitro) limits our ability to understand pregnancy disorders. Generating in vitro models that recapitulate the unique features of the human placenta has been challenging. Here, we describe detailed methods to isolate mononuclear villous trophoblasts (containing cytotrophoblasts and trophoblast stem cells) from first trimester placentae, and use both these and trophoblast stem cell populations that can be grown long term in a three-dimensional (3D) placental organoid culture system. Key words Human placental organoids, Villous cytotrophoblasts, Trophoblast stem cells, Trophoblast dysfunction, Fetal growth restriction

1

Introduction Fetal growth restriction (FGR) is defined as a birth weight of less than the third growth percentile for gestational age and is characterized by a deceleration or stagnation in fetal growth trajectory, placing the fetus at high risk of stillbirth or iatrogenic preterm birth [1, 2]. In addition to the immediate consequences of being born too small (increased perinatal morbidity and mortality), affected

These authors “Joanna L. James” and “Padma Murthi” contributed equally to this work and are joint senior authors. Sandeep Raha (ed.), Trophoblasts: Methods and Protocols, Methods in Molecular Biology, vol. 2728, https://doi.org/10.1007/978-1-0716-3495-0_19, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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offspring are at increased risk of cardiovascular disease, diabetes, and other chronic diseases in later life [3]. Prediction, detection, and treatment of pregnancies at high risk of FGR remain major challenges, with early delivery the only option to prevent stillbirth [4]. This is largely due to a lack of detailed understanding of the molecular etiology of FGR and the crucial role that placental dysfunction plays in this disorder [5]. Trophoblasts play crucial roles in fetal growth by facilitating adequate exchange of nutrients and oxygen between the maternal and fetal circulations. All mature trophoblasts differentiate from a population of trophoblast stem cells (TSCs). These rapidly proliferating cytotrophoblasts either fuse to form the outer multinucleated syncytiotrophoblast or, in the tips of anchoring villi, give rise to a different subset of cytotrophoblasts that differentiate into extravillous trophoblasts (EVTs), which grow out from the placenta and invade the decidua [6]. Here, EVTs play important roles in remodeling the spiral arteries into wide non-vasoactive vessels that can appropriately deliver large volumes of maternal blood to the placenta, which ensures an adequate supply of nutrients and oxygen to the fetus [7]. Important cross-talk also exists between trophoblasts and components of the mesenchymal core of villi, including mesenchymal stromal cells and vascular cells, that can impact placental morphogenesis [8]. Trophoblast differentiation and function from early pregnancy is thus crucial to pregnancy success and is dysfunctional in FGR pregnancies where placentae exhibit a thickened syncytiotrophoblast layer with decreased transporter expression, inadequate EVT invasion, and insufficient spiral artery remodeling [5]. Despite knowledge of its importance in pregnancy, the placenta remains one of the least studied organs in the human body. Human placental development and function is a highly orchestrated process, which is tightly controlled spatially and temporally. Establishing models that accurately describe this process is important to understand the developmental origins of FGR. Current placental research relevant to FGR mostly involves in vivo animal models and in vitro model systems using trophoblast-derived cells [6]. Both methods have significant limitations. In particular, neither adequately captures the in vivo structural and functional cellular microenvironment of the human placenta, thereby restricting insights into the molecular mechanisms leading to placental dysfunction, FGR, and stillbirth. Recently, the placental research field has been advanced through the development of placental organoids that can be derived from primary human placental cells. Organoids selfaggregate to form “mini organs” in vitro that mimic in vivo placental structure and function. Trophoblast organoid models were developed in 2018 from “cytotrophoblasts” that contain a mix of true cytotrophoblasts and human trophoblast stem cells (TSCs)

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[9, 10]. Indeed, organoids are considered stem cell models as they replicate the in vivo stem cell niche to maintain their differentiation capacity. Trophoblast organoids have also been generated directly from cultured TSCs, creating greater potential to genetically manipulate the input material and model developmental timelines of human placental development across gestation and in placental pathologies such as FGR. Here, we describe methods to isolate mononuclear villous trophoblasts (containing cytotrophoblasts and TSC) from first trimester placentae, and use both these and TSC populations to generate and assess trophoblast organoids.

2

Materials

2.1 Isolation of Mononuclear Villous Trophoblasts

1. 1× phosphate buffered saline solution (PBS). 2. DNase I stock (10 mg/mL): Weigh 1 g of lyophilized DNase I into a 100 mL glass bottle and add 100 mL of sterile PBS. Leave at 4 °C for 2 h, sterile filter, and freeze in 3 mL aliquots at 20 °C. 3. Trypsin stock (2.5%): Weigh 2.5 g of Trypsin into a 100 mL glass bottle and add 100 mL of sterile PBS. Leave at 4 °C for 2 h, sterile filter, and freeze in 2 mL aliquots at 20 °C.

2.2 Culture of Sidepopulation Trophoblasts and Okae Human TSC

1. Collagen-IV-coated 6-well plates: Add 5 μL of Collagen-IV stock (1 mg/mL) to 995 μL of PBS for each well to be coated. Incubate 1 mL of Collagen-IV solution in each well for 2 h at 37 °C, then remove and wash once with PBS. Wells can be stored in PBS for up to 1 month at 4 °C or used immediately. 2. Human TSC medium: DMEM/F12 medium, 1% ITS-X supplement, 0.3% Bovine Serum Albumin (BSA), 0.5% Penicillin Streptomycin, 0.1 mM 2-mercaptoethanol, 0.8 mM Valproic acid (Abcam, USA), 5 μM Y-27632 (Sigma-Aldrich, USA), 2 μM CHIR99021 (Sigma-Aldrich, USA), 1 μM SB431542 (Abcam, USA), 0.5 μM A83-01 (Sigma-Aldrich, USA), 1.5 μg/mL L-ascorbic acid, 50 ng/mL EGF.

2.3 Trophoblast Organoid Formation

1. Wash Medium: DMEM/F12 medium, 5% Fetal Bovine Serum (FBS), 1% Penicillin Streptomycin. 2. Trophoblast organoid medium (TOM): Advanced DMEM/ F12, 1:100 B27 Supplement, 1:100 N2 supplement, 1:100 L-glutamine, 10 mM HEPES, 0.5% Penicillin Streptomycin (all Gibco, USA), 3 μM CHIR99021 (Sigma-Aldrich, USA), 1 μM A83-01 (Sigma-Aldrich, USA), 100 ng/mL EGF (see Note 1).

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3. Trypan Blue (0.3%): Weigh 1.5 g of Trypan Blue powder into a 15 mL Falcon tube and add 5 mL of PBS. Sterile filter and store as 1 mL aliquots. 4. Cultrex® Reduced Growth Factor Basement Membrane Extract (BME), Type R1 (R&D Systems). 5. Cell Recovery Solution (Corning, USA). 2.4 Preparation of Organoids for Histological Analysis

1. Histogel (Epredia™ HG-4000-012). 2. 5% and 10% formalin. 3. Tissue-Tek cryomold. 4. P1000 pipette and pre-warmed pipette tips. 5. Water bath maintained at 60 °C. 6. PBS as wash buffer.

2.5 Fixation and Immunohistochemical Staining of Organoids

1. 10% formalin. 2. 0.1% saponin. 3. Blocking buffer containing 5% BSA/0.1% saponin. 4. Primary antibodies (see Table 1). 5. PBS as wash buffer. 6. Secondary antibodies conjugated with fluorophores. 7. Hoechst 33342 (1 mg/mL) or 4′,6-diamidino-2-phenylindole (DAPI).

3

Methods Carry out all procedures in a Class II tissue culture hood.

3.1 Generation of First Trimester Trophoblast Organoids: Isolation of First Trimester Mononuclear Villous Trophoblasts

1. Add 10 mL of PBS and all placental tissue fragments into a 10 cm plastic Petri dish. Use forceps to identify placental membranes with villous tissue attached and transfer to a separate Petri dish containing 5 mL of PBS. Fold the membrane so that villous trees protrude from the folded edge and roll the scalpel along this edge to remove villous fragments. Transfer all villous tissue to a 25 mL universal tube and record the final wet weight (a minimum of 1 g of villous tissue is recommended to proceed) (see Note 2). 2. Prepare the Enzyme Mixture by combining 2 mL of Trypsin stock (final concentration of 0.25%), 3 mL of DNase I stock (final concentration of 1.5 mg/mL), and 15 mL of PBS. Mix by inversion (see Note 3).

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Table 1 Protein markers that can be used to distinguish cytotrophoblast, syncytiotrophoblast, and extravillous trophoblast Protein marker

Notes

Reference

α6β4integrin

Unique cytotrophoblast marker. Specific detection of cytotrophoblasts in the placenta can be achieved using antibodies reactive with β4 integrin.

[13, 14]

CDX2 Commonly used “cytotrophoblast” markers that are also expressed by TSCs E-cadherin TEAD4 TP63

[9, 10]

Syncytin-1 Syncytiotrophoblast marker (also upregulated by some villous cytotrophoblast [15] during initiation of fusion) hCG ENDOU

Syncytiotrophoblast markers

[9, 13]

HLA-G

Extravillous trophoblast marker

[9, 10]

3. First enzymatic digest: Add 10 mL of the Enzyme Mixture to the villous tissue, gently invert to mix. Place the tube in a 37 °C water bath for 10 min, inverting to mix every 2 min. 4. Add PBS to fill the tube and let the villous tissue settle under gravity. Remove and discard all supernatant (containing syncytiotrophoblast fragments and EVTs), repeating this wash step 10 times until the supernatant is clear. 5. Second enzymatic digest: Add the remaining 10 mL of Enzyme Mix, gently invert to mix, and leave the tube on its side overnight at 4 °C. 6. The following morning, fill the tube with PBS, invert to mix, and let villous tissue settle. Set up a 50 mL Falcon tube containing 3 mL of FBS with a 70 μM mesh filter and collect supernatant into this tube. Repeat washes until four Falcon tubes are full. 7. Centrifuge the combined supernatants at 450 × g for 8 min. Resuspend cell pellets in 15 mL of Wash Medium and pipette the cell suspension into a 10 cm plastic Petri dish. Place into a 37 °C incubator for 10 min to deplete contaminating mesenchymal cells by plastic adhesion (see Note 4). 8. Collect the cell suspension into a 25 mL tube. Gently wash the Petri dish twice with 5 mL of Wash Medium to collect remaining trophoblasts and add this to the cell suspension. Centrifuge at 450 × g for 8 min (see Note 5).

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3.2 Side-population Trophoblast or Okae TSC Culture

1. Following fluorescence-activated cell sorting (FACS), or thawing from liquid nitrogen, seed side-population trophoblasts into the wells of 6-well plates pre-coated with Collagen IV (see Subheading 2.2) at a density of 10,000–15,000 cells/ well. Alternatively, TSCs generated using the methods published by Okae et al. can be obtained from primary first trimester placental tissue as described [11], or purchased commercially from Riken Cell Bank, Japan and seeded onto Collagen IV at 50,000 cells/well. 2. Culture cells in human TSC medium [11] until 80% confluent, with a full medium change every 2–3 days. 3. Detach cells by incubating each well with 1 mL of TrypLE for 5 min. Add an equal volume of Wash Medium to deactivate enzymes and centrifuge at 450 × g for 8 min. 4. Discard the supernatant and follow Subheading 3.3 to produce organoids.

3.3 Trophoblast Organoid Formation

1. Resuspend cell pellet in 0.5 mL of ice-cold TOM (see Note 1) and transfer the cell suspension to a 1.5 mL Eppendorf tube. 2. Centrifuge cells at 100 × g for 3 min in a benchtop centrifuge and resuspend in 1 mL of ice-cold TOM. Keep cells on ice. 3. Perform a cell count by adding 45 μL of Trypan Blue stock and 5 μL of well resuspended cells to a clean 1.5 mL Eppendorf tube. Mix well and count cells with a hemocytometer to calculate the volume of cell suspension required to produce the desired number of organoid domes (see Note 6). 4. Mix the 1 mL cell suspension thoroughly by gently pipetting up-and-down to avoid introducing bubbles and transfer the volume of required cell suspension calculated in step 6 to a new 1.5 mL Eppendorf tube (see Note 7). 5. Gently pipette the Cultrex® BME solution up-and-down a few times without introducing bubbles and remove the required volume. Add BME to the master mix cell suspension by gently pipetting up-and-down 5–10 times to ensure thorough mixing (see Notes 8 and 9). 6. Working quickly, place a 40 μL drop of master mix in the center of each well of a 12-well plate and transfer the plate to a 37 °C incubator for 2 min (see Note 10). 7. Turn the plate upside down to ensure even spreading of cells in the solidifying domes and incubate for a further 15 min at 37 °C. 8. Turn the plate back to normal orientation and remove from the incubator. Carefully overlay 500 μL of warm TOM over each dome.

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Fig. 1 Phase contrast images of organoids after 14 days of culture produced with (a) Mononuclear villous trophoblasts (cytotrophoblasts and side-population trophoblasts/TSCs), (b) Side-population trophoblasts at passage 7, isolated from a 7.5-week gestation placenta, and (c) Human TSCs (CT27, Okae et al. [11]). Scale bars represent 200 μm

9. Return the plate to a 37 °C incubator and allow organoids to form in a humidified ambient oxygen atmosphere containing 5% CO2. Perform a full medium change every 2–3 days. Small organoids (50–100 μm) are visible 4–5 days after seeding, and organoids are large enough to passage or manipulate for paraffin embedding (200–500 μm) by day 20–30 in culture (Fig. 1). 3.4 Organoid Passaging, Freezing, or Stimulation of Extravillous Trophoblast Outgrowth

1. To harvest organoids remove medium from wells and perform a PBS wash, then incubate organoid domes with 500 μL of Cell Recovery Solution for 1 h at 4 °C (see Note 13). 2. Use an FBS-coated pipette tip to resuspend the organoid dome in each well and collect released organoids (small white dots should be visible) in a 15 mL Falcon tube. 3. Rinse each well with ice-cold PBS to maximize organoid retention. 4. Centrifuge for 3 min at 100 × g to pellet organoids. Discard supernatant. From here it is possible to freeze or passage organoids or set up cultures from which extravillous trophoblast outgrowth can be stimulated as below. 5. To freeze organoids for later use: Resuspend in 10% Dimethylsulfoxide (DMSO) in FBS and freeze at -80 °C prior to transfer to liquid nitrogen. 6. To passage organoids: Organoids can be passaged for further culture by dissociating large organoids with repeated pipetting through a P200 tip. These fragments can be resuspended in TOM and Cultrex® BME as per Subheading 3.3. 7. To stimulate extravillous trophoblast outgrowth: Follow Subheading 3.3 to reintroduce organoids to Cultrex® BME domes, however, use TOM without CHIR99021 to promote extravillous trophoblast outgrowth formation.

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3.5 Histological Assessment of Trophoblast Organoids

1. Fix organoids in Cultrex® BME with 500 μL of 5% formalin per well overnight.

3.5.1 Fixation and Embedding of Trophoblast Organoids for Immunohistochemical Analysis

3. Dissociate Cultrex® BME mechanically with a P1000 pipette by slowly pipetting 10 times up and down (see Note 14).

2. Remove formalin and rinse in 500 μL PBS.

4. Transfer organoid-containing solution to a tube and centrifuge at 400 × g for 3 min (see Note 15). 5. Warm a 1 mL aliquot of Histogel in a 60 °C water bath (see Note 16). 6. Add 150 μL of Histogel and quickly resuspend the organoids in the tube. 7. Transfer the entire volume quickly into a Tissue-Tek cryomould, making a single drop in the center of the mold. Wait for 3–5 min for solidification. 8. Once the sample is set, remove the cryomould and transfer to a histology cassette. 9. Store submerged in 10% formalin until processing into paraffin wax blocks. 10. Paraffin wax block formation can be done using a short 3.5h processing cycle on a tissue processor. Cut 5 μm serial sections for immunohistochemical analysis.

3.5.2 Whole Mount Staining of Trophoblast Organoids

1. Remove medium from the organoid cultures and fix in 10% formalin for 30 min at room temperature. 2. Remove fixative and rinse organoids in PBS twice for 5 min each. Give organoids 10 min after the addition of each wash volume to fully settle by gravity (see Note 17). 3. Permeabilize organoids with 0.1% saponin for 15 min. 4. Remove and add blocking buffer containing 5% BSA/0.1% saponin for 30 min at room temperature. 5. Remove blocking buffer and sequentially add primary antibody and incubate overnight at 4 °C and secondary antibodies conjugated with fluorophores for 2 h at room temperature the following day (see Note 18). 6. Between steps, wash organoids in PBS three times for 15 min each on a shaker at slow speed. 7. Image-stained organoids intact by confocal microscopy, incubate with 10 μg/mL of Hoechst 33342 for 30 min at room temperature, then wash in PBS three times as above. Organoids can be imaged in PBS. Alternatively, if organoids are too large to enable adequate confocal imaging of the center of the structures, organoids can be mounted and sectioned as in

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Subheading 3.5.1, and nuclei can be stained using a mounting medium with DAPI. 8. Slides can be stored covered in a foil at 4 °C until ready to visualize the immunoreactivity by fluorescence or confocal microscopy.

4

Notes 1. Keep this medium on ice at the beginning of Subheading 3.3 and transfer to a 37 °C water bath at the end of step 6 once the master mix is complete. 2. Accumulate dissected villous tissue in a small pile in a dry Petri dish as you work rather than transferring immediately into the tube. This allows excess PBS to be excluded from the final wet weight and allows for easier transfer of a greater volume of tissue fragments at a time. 3. If the sample is especially fragmented, villous fragments can be collected in PBS and strained through a 70 μm mesh filter to collect tissue. This is particularly valuable for precious and/or small samples (i.e., 5–6 weeks gestation). 4. Depletion of villous mesenchymal cells is especially important as these cells will rapidly proliferate and overtake the gel dome. 5. This generates a population of mononuclear villous trophoblasts, consisting of both TSCs and cytotrophoblasts. Organoids can also be generated from TSC only (see modifications in Subheadings 3.3 and 3.4). To use the side-population technique to directly isolate TSC from this population by flow cytometry refer to previously published methods [12, 13]. Alternately, cells could be plated under the conditions of Okae et al. to generate TSCs via prolonged culture [11]. 6. Each 40 μL dome consists of 60% (24 μL) Cultrex® BME and 40% trophoblasts (105 cells per dome) suspended in ice-cold TOM (16 μL). Calculate the volume of Cultrex® BME and TOM cell suspension required for the master mix depending on the number of domes desired. We recommend a minimum of 12 domes to ensure successful passaging, but this will depend on the intended downstream analyses (for 12 domes, the master mix would contain 288 μL of Cultrex ® BME (24 μL of Cultrex per dome × 12) and 192 μL of TOM containing cells (16 μL of cell suspension per dome × 12). Calculate the volume required from the 1 mL cell suspension using the following equation: Number of cells required × 1000 = XXX μL cell suspension Cell concentration cells mL

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7. If the volume calculated in step 3 is less than the volume of TOM cell suspension required in the master mix, centrifuge the required cells at 100 × g for 3 min and remove as much residual medium as possible using a P20 pipette, then resuspend in the required volume of ice-cold TOM. 8. Thaw the Cultrex® BME solution at 4 °C overnight on ice and place 200 μL pipette tips in a -20 °C freezer. Always handle Cultrex® BME and the master mix with ice-cold tips to prevent premature polymerization. It may be necessary to place an ice box sprayed with 70% ethanol in the biological hood to keep the master mix cold as much as possible throughout the following steps. 9. It is crucial to prevent bubble formation in the Cultrex® BME and master mix as bubbles can interfere with organoid formation and visibility, always pipette gently with the tip below the surface of the solution. 10. The domes tend to spread into the edges of wells in 24-well plates and thus we recommend a 12-well plate. To further combat this issue, place the plate into a 37 °C incubator for 10 min prior to addition of organoid domes. 11. PBS can be added to spare wells or the reservoir surrounding the plate to prevent excess evaporation. 12. The time required for organoids to reach 200–500 μm in diameter varies between different placental samples and is slower with increasing gestational age. Cultures are passaged or harvested for further processing when most organoids have reached this size for ease of manipulation, rather than using a culture time point as a landmark. 13. Incubate on a horizontal shaker or lightly tap the plate every 20 min to improve depolymerization. 14. Coat pipette tips with FBS for 10 min to prevent organoids sticking to plastic. 15. Glass tubes are preferred here to prevent organoids sticking to plastic, alternatively, plastic tubes can be coated with FBS for 10 min at room temperature before use. 16. Pre-warm pipette tips prior to using them for Histogel preparation. 17. To maximize organoid retention, ensure that starting organoids are mostly between 200 and 500 μm in diameter and avoid disturbing the tube until organoids have fully settled by gravity. Coat tubes and pipette tips with FBS for 10 min at room temperature before use to prevent organoids sticking to plastic.

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18. Antigens useful to distinguish cytotrophoblasts and syncytiotrophoblast are outlined in Table 1. These markers are useful in dual-immunofluorescence labelling experiments to characterize the organoids obtained, or to distinguish the cell type that other functional markers of interest are expressed in.

Acknowledgments This work was supported by a RANZCOG Mercia Barnes Trust Project Grant. C. Sun is the recipient of an AMRF Doctoral Scholarship (1219006). References 1. Gordijn S et al (2016) Consensus definition of fetal growth restriction: a Delphi procedure. Ultrasound Obstet Gynecol 48(3):333–339 2. Beune IM et al (2018) Consensus based definition of growth restriction in the newborn. J Pediatr 196:71–76. e1 3. Barker DJP (2006) Adult consequences of fetal growth restriction. Clin Obstet Gynecol 49(2): 270–283 4. Figueras F, Gardosi J (2011) Intrauterine growth restriction: new concepts in antenatal surveillance, diagnosis, and management. Am J Obstet Gynecol 204(4):288–300 5. Sun C et al (2020) The placenta in fetal growth restriction: what is going wrong? Placenta 96: 10–18 6. Kno¨fler M et al. (2019) Human placenta and trophoblast development: key molecular mechanisms and model systems. Cell Mol Life Sci 76(18):3479–3496 7. Burton G et al (2009) Rheological and physiological consequences of conversion of the maternal spiral arteries for uteroplacental blood flow during human pregnancy. Placenta 30(6):473–482 8. Surico D et al (2019) Preeclampsia and intrauterine growth restriction: role of human umbilical cord mesenchymal stem cellstrophoblast cross-talk. PLoS One 14(6): e0218437

9. Haider S et al (2018) Self-renewing trophoblast organoids recapitulate the developmental program of the early human placenta. Stem Cell Rep 11(2):537–551 10. Turco MY et al (2018) Trophoblast organoids as a model for maternal–fetal interactions during human placentation. Nature 564(7735): 263–267 11. Okae H et al (2018) Derivation of human trophoblast stem cells. Cell Stem Cell 22(1): 50–63. e6 12. James JL et al (2015) Isolation and characterisation of a novel trophoblast side-population from first trimester placentae. Reproduction 150(5):449–462 13. Gamage TKJB et al (2020) Side-population trophoblasts exhibit the differentiation potential of a trophoblast stem cell population, persist to term, and are reduced in fetal growth restriction. Stem Cell Rev Rep 16(4):764–775 14. Korhonen M et al (1991) Distribution of the alpha 1-alpha 6 integrin subunits in human developing and term placenta. Lab Invest 65(3):347–356 15. Po¨tgens AJG et al (2004) Syncytin: the major regulator of trophoblast fusion? Recent developments and hypotheses on its action. Hum Reprod Update 10(6):487–496

INDEX A

H

Adhesion ...............................................64, 132, 154, 156, 160, 161, 239 Angiostamp700 ........................................... 132, 133, 135 Apparent permeability coefficient (Papp).....113, 115–116

Human.................................3–11, 25–33, 45–50, 58, 59, 70, 71, 88, 89, 96, 97, 99, 123–128, 166, 174, 183–193, 195, 223, 224, 236, 237, 241 Human placenta ........................48, 49, 88, 96, 106, 107, 166, 223, 225, 236 Human term placenta ......................................4, 106, 124

B BeWo................................. 50, 58, 88, 89, 91, 93, 95–97, 99–103, 109–113, 115–117, 124, 137, 184, 197

C Cell fusion ............................................ 13, 20, 21, 87–89, 91–93, 95, 101 Cell isolation................................... 25–33, 188, 190, 192 Cholesterol ................................. 123–128, 167, 174, 176 Cholesterol efflux ................................................. 123–128 Co-culture ..................................... 50, 55, 58, 59, 63–65, 72, 88, 93, 183–193 Cytotrophoblast (CTB) .........................4, 27, 32, 48, 87, 124, 125, 137–146, 192, 239

D Di-8-ANEPPS ................................................88, 100–103

E EDC/NHS.................................................................... 175 Embryos ...................................25–33, 83, 105, 133, 195 Extravillous trophoblast (EVT) cells.................. 150, 151, 183–193, 196, 201, 211–214, 220, 223, 224, 236, 239, 241

I Immunofluorescence .................... 52, 61, 62, 72, 87–97, 188, 192, 212, 230 Impedance ............................................................ 150, 154 Invasion ......................................... 46, 48–50, 56, 65, 68, 69, 73, 132, 150, 151, 158–161, 165, 183, 195, 224, 236 In vitro cell culture.................................... 26, 29, 91, 184 In vivo angiogenesis...................................................... 132 In Vivo Imaging...............................................79, 82, 133

L Leucine uptake ..................................................... 105–119 Lipoproteins ........................................................ 123, 124, 127, 128 Liposomes............................................................ 167–171, 173–179 Live cell imaging ............................................................. 65 L-[3H]-leucine transport .............................................. 111 Long term culture............................................................. 4

M

Fetomaternal interface ......................................... 131, 132 Fluorescence imaging .........................102, 132, 133, 135 Follow-up of choriocarcinoma dissemination ............... 78 Fusion index (FI) ................................88, 95, 97, 99, 102

Membrane labeling .................................................99–103 Methodology...................................................... 6–10, 160 Microfluidics.................................................224–227, 230 Migration.............................. 50, 65, 66, 69, 73, 78, 132, 150, 156–161, 224 Migratory trophoblast (MTB) ....................26–28, 31, 32 Mitochondria........................................................ 137–146

G

N

Gestational choriocarcinoma (CC) ..........................77, 78 Green fluorescent protein (GFP) ..................... 55, 88–93, 95, 97, 103

Na+-independent amino acid transport .............. 106, 107

F

Sandeep Raha (ed.), Trophoblasts: Methods and Protocols, Methods in Molecular Biology, vol. 2728, https://doi.org/10.1007/978-1-0716-3495-0, © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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248 Index O

Organotypic...............................................................35, 38 Orthotopic model .....................................................78, 81 Oxidative phosphorylation ........................................... 146 Oxygen consumption rate (OCR) ..................... 137, 138, 143, 144, 146

Spheroids .................................................... 50, 55, 59, 65, 66, 73, 224 Split protein complementation.................................87–97 Syncytiotrophoblast (STB) .............................3, 4, 10, 13, 26–28, 32, 47, 48, 54, 78, 87, 89, 92, 93, 95, 97, 99, 124, 125, 127, 166, 169, 174, 196, 197, 207, 212, 236, 239, 245

P

T

Percoll gradient ............................. 4, 14, 17–20, 23, 125, 184, 186, 189, 193, 200, 202, 203 Peri-implantation ......................................................25–33 Placenta........................................ 3, 6, 10, 13–25, 35–41, 45, 46, 48, 49, 57, 58, 71, 78, 80–83, 125, 131, 132, 135, 165–171, 173, 174, 183, 187, 189, 195, 196, 198, 200, 202, 219, 223, 236, 239, 241 Placental chondroitin sulfate A (plCSA)............. 174, 175 Placental development ................................. 87, 149, 197, 223, 236, 237 Placental drug delivery ........................................ 173–179 plCSA binding peptide (plCSA-BP) .................. 174, 175, 177, 178 Primary cells ........................................25, 50, 88, 96, 197 Primary culture.................................................4, 109, 124 Primary trophoblast isolation ....................................... 124 Proliferation......................................69, 79, 96, 150–152, 154, 156, 160, 169, 198, 233

Three-dimensional (3D)............................................... 224 Trans-epithelial electrical resistance (TEER)...... 113–117 Transfection........................................................ 13–24, 97 Transporters .......................................106–109, 123, 124, 128, 197, 236 Transwell.................................................68, 70, 109, 110, 113–117, 192, 224 Trophoblast ...............................3, 19, 25, 45, 78, 87, 99, 124, 132, 137, 149, 165, 174, 184, 195, 223, 236 Trophoblast cells .................................. 19, 25–33, 45–52, 55, 56, 58, 59, 63–70, 77–84, 100, 105–119, 123–128, 132, 137–146, 184, 189, 192, 193, 197, 223–225 Trophoblast organoids (TB-ORGs) .................. 195–198, 202, 206–221, 236, 237, 240 Trophoblast syncytialization...................................99–103 Trypsin digestion ............................. 16, 17, 31, 125, 228

Q Quantitative assessment of gestational choriocarcinoma ............................................77–84

S Seahorse XF24 Extracellular Flux analyzer......... 137–146 Single-cell collection ....................................................... 32 SiRNA .......................................................... 15, 16, 20, 22 Slice cultures..............................................................35–41

U Uterine natural killer (uNK) cells .......................... 46, 59, 183–193

V Villous cytotrophoblastic cell (vCTB) .....................13–24

X xCELLigence........................................................ 149–161