Adipose-Derived Stem Cells: Methods and Protocols (Methods in Molecular Biology, 2783) [3 ed.] 1071637614, 9781071637616

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Methods in Molecular Biology 2783

Jeffrey M. Gimble · Bruce A. Bunnell Trivia Frazier · Cecilia Sanchez  Editors

Adipose-Derived Stem Cells Methods and Protocols Third Edition

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-bystep 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.

Adipose-Derived Stem Cells Methods and Protocols Third Edition

Edited by

Jeffrey M. Gimble Obatala Sciences Inc, New Orleans, LA, USA

Bruce A. Bunnell Department of Microbiology, Immunology and Genetics, University of North Texas Health Science, Fort Worth, TX, USA

Trivia Frazier Obatala Sciences Inc, New Orleans, LA, USA

Cecilia Sanchez Obatala Sciences Inc, New Orleans, LA, USA

Editors Jeffrey M. Gimble Obatala Sciences Inc New Orleans, LA, USA Trivia Frazier Obatala Sciences Inc New Orleans, LA, USA

Bruce A. Bunnell Department of Microbiology, Immunology and Genetics University of North Texas Health Science Fort Worth, TX, USA Cecilia Sanchez Obatala Sciences Inc New Orleans, LA, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-3761-6 ISBN 978-1-0716-3762-3 (eBook) https://doi.org/10.1007/978-1-0716-3762-3 © 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 For over two decades, adipose-derived stromal/stem cells (ASC) and stromal vascular fraction (SVF) cells have continued receiving attention from regenerative medicine due to their potential for differentiation, immunomodulation, and exosome/secretome-mediated paracrine actions. Since the publication of the second edition of this book, increased focus has been placed on the use of ASC and SVF cells in three-dimensional hydrogel-based scaffolds for the development of microphysiological systems (MPS) serving as in vitro humanized assays and alternatives to in vivo preclinical animal models. This Methods in Molecular Biology third edition volume focuses primarily, but not exclusively, on humanderived ASC and SVF; we have been fortunate to receive new and revised protocols from experts in the field. The contents are organized into four discrete sets of chapters. The initial section focuses on human ASC’s isolation, characterization, and differentiation. This is followed by a section describing the isolation and characterization of ASC and SVF from canine, feline, and murine tissues. Next, we present a new group of chapters on hydrogels, scaffolds, and microphysiological systems. The final section, consisting almost exclusively of new content, describes assays and applications using ASC. The chapters provide a “how to” protocol suitable for novices and experts alike to expand the research infrastructure exploring adipose tissue biology in the academic, biotech, and pharmaceutical communities. The editors are grateful to the many coauthors, many of whom are fellow members of the International Federation of Adipose Therapeutics and Science (IFATS), who took time out of their busy schedules and graciously contributed manuscripts to this body of work. We also thank Professor John Walker, Series Editor, and the editorial staff at Springer, whose direction, advice, and editorial support were both needed and appreciated. Finally, we all owe our appreciation to our families who sacrificed quality time on nights, weekends, and holidays, permitting us to complete this work on schedule. Fort Worth, TX, USA New Orleans, LA, USA New Orleans, LA, USA New Orleans, LA, USA New Orleans, LA, USA

Bruce A. Bunnell Trivia Frazier Jeffrey M. Gimble Katie Hamel Cecilia Sanchez

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Acknowledgements The editors express their sincere appreciation to Katie Hamel, PhD, for her expert assistance in organizing and managing the review process during the completion of the Third Edition of Adipose-Derived Stem Cells, Methods Molecular biology Vo. 2783.

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

PART I

METHODS TO ISOLATE, CHARACTERIZE, AND DIFFERENTIATE HUMAN ASC

1 Isolation of Human Adipose-Derived Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . Elizabeth Montano, Sara Al-Ghadban, and Bruce A. Bunnell 2 Isolation of Human Adipose-Derived Stem Cells from Lipoaspirates . . . . . . . . . . Yuejia Li, Ziyi Mei, Jie Li, and Jeffrey M. Gimble 3 Isolation of Perivascular Mesenchymal Progenitor Cells from Human Adipose Tissue by Flow Cytometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Neelima Thottappillil, Mario A. Gomez-Salazar, Mary Archer, Bruno Pe´ault, and Aaron W. James 4 Soft Tissue Reconstruction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mario Alessandri-Bonetti, Francesco M. Egro, and Kacey G. Marra 5 Adult Stem Cells Freezing Processes and Cryopreservation Protocols . . . . . . . . . Mohan Kumar Dey and Ram V. Devireddy

PART II

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3 13

25

35

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ANIMAL ASC MODELS

6 Isolation of Murine Adipose-Derived Stromal/Stem Cells for Adipogenic and Osteogenic Differentiation or Flow Cytometry-Based Analysis. . . . . . . . . . . . 93 Matthew C. Scott, Chul-Hong Park, Marilyn Dietrich, Xiying Wu, Jeffrey M. Gimble, Carrie M. Elks, Ji Suk Chang, and Z. Elizabeth Floyd 7 Isolation of Murine Adipose-Derived Stromal/Stem Cells Using an Explant Culture Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Ziyi Mei, Yuejia Li, Jeffrey M. Gimble, and Jie Li 8 Canine Adult Adipose Tissue-Derived Multipotent Stromal Cell Isolation, Characterization, and Differentiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Takashi Taguchi and Mandi J. Lopez 9 Feline Adult Adipose Tissue-Derived Multipotent Stromal Cell Isolation and Differentiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137 Catherine Takawira, Wei Duan, Takashi Taguchi, and Mandi J. Lopez

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PART III 10

11

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15

16

17

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19 20

HYDROGELS, SCAFFOLDS, AND 3 DIMENSIONAL MICROPHYSIOLOGICAL SYSTEMS

Manufacturing of a Human Adipose-Derived Hydrogel . . . . . . . . . . . . . . . . . . . . . Jordan Robinson, Haley Lassiter, Katie Hamel, Xiying Wu, Jeffrey M. Gimble, Trivia Frazier, and Cecilia Sanchez Quality Control Assessment of Human Adipose-Derived Hydrogels . . . . . . . . . . Haley Lassiter, Jordan Robinson, Katie Hamel, Jeffrey M. Gimble, Trivia Frazier, and Cecilia Sanchez Analysis of Biomechanical Properties of Adipose-Derived Hydrogels for Adipose-Derived Stem Cell-Based Hydrogel Culture. . . . . . . . . . . . . . . . . . . . . Jorge A. Belgodere Combination of Adipose Stem Cells and Decellularized Adipose Tissue Hydrogel for Osteogenic Applications. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kainat Ahmed, Haadia Tauseef, and Omair A. Mohiuddin Preparation of Decellularized Amniotic Membrane and Adipose-Derived Stem Cell Seeding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Haadia Tauseef, Kainat Ahmed, Asmat Salim, and Omair A. Mohiuddin Robust Generation of ASC Spheroids for Use as 3D Cultures and in Bioprinted Tissue Models. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Martin Watzling, Hannes Horder, Petra Bauer-Kreisel, and Torsten Blunk Generating and Characterizing Adipose Spheroids f rom Adipose Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Charles Amurgis, W. Vincent Nerone, and Lauren Kokai Spheroid Formation of Human Adipose-Derived Stem Cells Using a Liquid Overlay Technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sara Al-Ghadban and Bruce A. Bunnell Repurposing Decellularized Lung to Generate Vascularized Fat . . . . . . . . . . . . . . Lindsey K. Huff, Zihan Ling, Megan K. DeBari, Xi Ren, and Rosalyn D. Abbott In Vitro Culture of White Adipose Tissue. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jake J. Fontenot and Frank H. Lau Bioreactor Culture to Create Adipose Tissue from Human Mesenchymal Stromal Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katelyn E. Lipa, Meagan J. Makarcyzk, Sophie Hines, Celeste E. Lintz, Bruce A. Bunnell, and Hang Lin

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PART IV ANALYTICAL ASSAYS AND APPLICATIONS OF ASC 21

A Method for Screening a Kinase Inhibitor Drug Library on ASCs . . . . . . . . . . . 303 Caroline H. Rinderle and Bruce A. Bunnell

Contents

Analyses and Utilization of Selectively Tuned Human Adipose-Derived Stem Cell Exosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . John W. Ludlow and Benjamin M. Buehrer 23 Functional Assays in 3D ObaCell® Fat-on-a-Chip Cultures: Lipolysis and Glucose Uptake Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katie Hamel, Emma Rogers, Jeffrey M. Gimble, Trivia Frazier, and Cecilia Sanchez 24 Monitoring ADSC Metabolism Using the Seahorse Analyzer. . . . . . . . . . . . . . . . . Daniel Rittenhouse and Kevin J. Zwezdaryk 25 Mechanical Stimulation of Adipose-Derived Stem Cells for Functional Tissue Engineering of the Musculoskeletal System via Cyclic Hydrostatic Pressure, Simulated Microgravity, and Cyclic Tensile Strain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rachel C. Nordberg, Josie C. Bodle, and Elizabeth G. Loboa 26 Nuclear Transcription Factor Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Joseph A. Straub and Jamie J. Newman

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Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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349 367 391

Contributors ROSALYN D. ABBOTT • Department of Biomedical Engineering, Carnegie Mellon University, Pittsburgh, PA, USA; Department of Materials Science and Engineering, Carnegie Mellon University, Pittsburgh, PA, USA KAINAT AHMED • Dr. Panjwani Center for Molecular Medicine and Drug Research, International Center for Chemical and Biological Sciences, University of Karachi, Karachi, Pakistan MARIO ALESSANDRI-BONETTI • Department of Plastic Surgery, University of Pittsburgh, Pittsburgh, PA, USA SARA AL-GHADBAN • Department of Microbiology, Immunology and Genetics, School of Biomedical Sciences, University of North Texas Health Sciences Center, Fort Worth, TX, USA CHARLES AMURGIS • Department of Plastic Surgery, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA MARY ARCHER • Department of Pathology, Johns Hopkins University, Baltimore, USAMD PETRA BAUER-KREISEL • Department of Trauma, Hand, Plastic and Reconstructive Surgery, University Hospital Wu¨rzburg, Wu¨rzburg, Germany JORGE A. BELGODERE • Department of Biological and Agricultural Engineering, Louisiana State University and Agricultural Center, Baton Rouge, LA, USA; Section of Hematology and Medical Oncology, Department of Medicine, Tulane University School of Medicine, New Orleans, LA, USA TORSTEN BLUNK • Department of Trauma, Hand, Plastic and Reconstructive Surgery, University Hospital Wu¨rzburg, Wu¨rzburg, Germany JOSIE C. BODLE • Duke University, Durham, NC, USA BENJAMIN M. BUEHRER • Zen-Bio, Inc., Durham, NC, USA BRUCE A. BUNNELL • Department of Microbiology, Immunology and Genetics, School of Biomedical Sciences, University of North Texas Health Sciences Center, Fort Worth, TX, USA JI SUK CHANG • Pennington Biomedical Research Center, Baton Rouge, LA, USA MEGAN K. DEBARI • Department of Materials Science and Engineering, Carnegie Mellon University, Pittsburgh, PA, USA RAM V. DEVIREDDY • Bioengineering Laboratory, Department of Mechanical Engineering, Louisiana State University, Baton Rouge, LA, USA MOHAN KUMAR DEY • Bioengineering Laboratory, Department of Mechanical Engineering, Louisiana State University, Baton Rouge, LA, USA MARILYN DIETRICH • Pennington Biomedical Research Center, Baton Rouge, LA, USA WEI DUAN • Laboratory for Equine and Comparative Orthopedic Research, School of Veterinary Medicine, Louisiana State University, Baton Rouge, LA, USA FRANCESCO M. EGRO • Department of Plastic Surgery, University of Pittsburgh, Pittsburgh, PA, USA CARRIE M. ELKS • Pennington Biomedical Research Center, Baton Rouge, LA, USA Z. ELIZABETH FLOYD • Pennington Biomedical Research Center, Baton Rouge, LA, USA JAKE J. FONTENOT • Department of Surgery, School of Medicine, Louisiana State University Health Sciences Center-New Orleans, New Orleans, LA, USA

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Contributors

TRIVIA FRAZIER • Obatala Sciences, Inc., New Orleans, LA, USA JEFFREY M. GIMBLE • Obatala Sciences, Inc., New Orleans, LA, USA MARIO A. GOMEZ-SALAZAR • Department of Pathology, Johns Hopkins University, Baltimore, USAMD KATIE HAMEL • Obatala Sciences, Inc., New Orleans, LA, USA SOPHIE HINES • Department of Orthopaedic Surgery, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA HANNES HORDER • Department of Trauma, Hand, Plastic and Reconstructive Surgery, University Hospital Wu¨rzburg, Wu¨rzburg, Germany LINDSEY K. HUFF • Department of Biomedical Engineering, Carnegie Mellon University, Pittsburgh, PA, USA AARON W. JAMES • Department of Pathology, Johns Hopkins University, Baltimore, USAMD LAUREN KOKAI • Department of Plastic Surgery, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA HALEY LASSITER • Obatala Sciences, Inc, New Orleans, LA, USA FRANK H. LAU • Department of Surgery, School of Medicine, Louisiana State University Health Sciences Center-New Orleans, New Orleans, LA, USA JIE LI • College of Stomatology, Chongqing Medical University, Chongqing, China YUEJIA LI • College of Stomatology, Chongqing Medical University, Chongqing, China HANG LIN • Department of Orthopaedic Surgery, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA; Department of Bioengineering, University of Pittsburgh Swanson School of Engineering, Pittsburgh, PA, USA ZIHAN LING • Department of Biomedical Engineering, Carnegie Mellon University, Pittsburgh, PA, USA CELESTE E. LINTZ • Department of Orthopaedic Surgery, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA; Department of Bioengineering, University of Pittsburgh Swanson School of Engineering, Pittsburgh, PA, USA KATELYN E. LIPA • Department of Orthopaedic Surgery, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA; Department of Bioengineering, University of Pittsburgh Swanson School of Engineering, Pittsburgh, PA, USA ELIZABETH G. LOBOA • Southern Methodist University, Dallas, TX, USA MANDI J. LOPEZ • Laboratory for Equine and Comparative Orthopedic Research, School of Veterinary Medicine, Louisiana State University, Baton Rouge, LA, USA JOHN W. LUDLOW • Zen-Bio, Inc., Durham, NC, USA MEAGAN J. MAKARCYZK • Department of Orthopaedic Surgery, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA; Department of Bioengineering, University of Pittsburgh Swanson School of Engineering, Pittsburgh, PA, USA KACEY G. MARRA • Department of Plastic Surgery, University of Pittsburgh, Pittsburgh, PA, USA; McGowan Institute for Regenerative Medicine, University of Pittsburgh, Pittsburgh, PA, USA; Department of Bioengineering, University of Pittsburgh, Pittsburgh, PA, USA ZIYI MEI • College of Stomatology, Chongqing Medical University, Chongqing, China OMAIR A. MOHIUDDIN • Dr. Panjwani Center for Molecular Medicine and Drug Research, International Center for Chemical and Biological Sciences, University of Karachi, Karachi, Pakistan ELIZABETH MONTANO • Department of Microbiology, Immunology and Genetics, School of Biomedical Sciences, University of North Texas Health Sciences Center, Fort Worth, TX, USA

Contributors

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W. VINCENT NERONE • Department of Plastic Surgery, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA JAMIE J. NEWMAN • Department of Biological Sciences, Louisiana Tech University, Ruston, LA, USA RACHEL C. NORDBERG • University of California, Irvine, CA, USA CHUL-HONG PARK • Pennington Biomedical Research Center, Baton Rouge, LA, USA BRUNO PE´AULT • Center for Cardiovascular Research, University of Edinburgh, Edinburgh, UK; Orthopaedic Hospital Research Center and Broad Stem Cell Research Center, David Geffen School of Medicine, University of California, Los Angeles, CA, USA XI REN • Department of Biomedical Engineering, Carnegie Mellon University, Pittsburgh, PA, USA CAROLINE H. RINDERLE • Department of Microbiology, Immunology and Genetics, School of Biomedical Sciences, University of North Texas Health Science Center, Fort Worth, TX, USA DANIEL RITTENHOUSE • Department of Microbiology & Immunology, Tulane University School of Medicine, New Orleans, LA, USA JORDAN ROBINSON • Obatala Sciences, Inc, New Orleans, LA, USA EMMA ROGERS • Obatala Sciences, Inc., New Orleans, LA, USA ASMAT SALIM • Dr. Panjwani Center for Molecular Medicine and Drug Research, International Center for Chemical and Biological Sciences, University of Karachi, Karachi, Pakistan CECILIA SANCHEZ • Obatala Sciences, Inc., New Orleans, LA, USA MATTHEW C. SCOTT • Pennington Biomedical Research Center, Baton Rouge, LA, USA JOSEPH A. STRAUB • Department of Biology and Microbiology, Northwestern State University, Natchitoches, LA, USA TAKASHI TAGUCHI • Laboratory for Equine and Comparative Orthopedic Research, School of Veterinary Medicine, Louisiana State University, Baton Rouge, LA, USA CATHERINE TAKAWIRA • Laboratory for Equine and Comparative Orthopedic Research, School of Veterinary Medicine, Louisiana State University, Baton Rouge, LA, USA HAADIA TAUSEEF • Dr. Panjwani Center for Molecular Medicine and Drug Research, International Center for Chemical and Biological Sciences, University of Karachi, Karachi, Pakistan NEELIMA THOTTAPPILLIL • Department of Pathology, Johns Hopkins University, Baltimore, USAMD MARTIN WATZLING • Department of Trauma, Hand, Plastic and Reconstructive Surgery, University Hospital Wu¨rzburg, Wu¨rzburg, Germany XIYING WU • Obatala Sciences, Inc, New Orleans, LA, USA KEVIN J. ZWEZDARYK • Department of Microbiology & Immunology, Tulane University School of Medicine, New Orleans, LA, USA; Tulane Center for Aging, Tulane University School of Medicine, New Orleans, LA, USA; Tulane Brain Institute, Tulane University School of Medicine, New Orleans, LA, USA

Part I Methods to Isolate, Characterize, and Differentiate Human ASC

Chapter 1 Isolation of Human Adipose-Derived Stem Cells Elizabeth Montano, Sara Al-Ghadban, and Bruce A. Bunnell Abstract Human adipose-derived stromal/stem cells (hASCs) are a promising source of adult stem cells used in numerous applications in regenerative medicine. We present the protocols from our laboratory for isolating and expanding hASCs. The isolation of hASCs involves the enzymatic digestion of adipose tissue and subsequent culturing of the isolated cells. Key words Adipose-derived stromal/stem cells (ASCs), Biopsy, Collagenase, Expansion, Human, Lipoaspirate, Mesenchymal stem cells (MSCs), Stromal vascular fraction (SVF)

1

Introduction ASCs are an excellent source of adult stem cells for regenerative medicine due to their accessibility, abundance, and multi-lineage differentiation potential [1–3]. Isolation of ASCs from adipose tissue is critical for their use in various applications. The resulting ASCs can be characterized by their surface marker expression, differentiation potential, and gene expression profiles [4–8]. This protocol provides isolation methods for primary in vitro cultures of human ASCs, including the enzymatic digestion protocol, culture conditions, and characterization methods.

2

Materials

2.1

Tissue

2.2

Supplies

Subcutaneous adipose tissue collected from liposuction aspirates. 1. 50 mL conical tubes. 2. Stericup quick release-GP sterile vacuum filtration system. 3. 50 mL Steriflip filter unit. 4. 100 mL borosilicate glass storage bottle.

Jeffrey M. Gimble et al. (eds.), Adipose-Derived Stem Cells: Methods and Protocols, Methods in Molecular Biology, vol. 2783, https://doi.org/10.1007/978-1-0716-3762-3_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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5. 250 mL polypropylene beaker. 6. 2 mL borosilicate Pasteur pipettes. 7. Parafilm. 8. Sterile 70 μm cell strainers. 9. Sterile 100 μm cell strainers. 10. 175 cm2 cell culture flasks. 11. 0.25% Trypsin-EDTA. 12. Hemocytometer. 13. Mr. Frosty freezing container. 14. Fluorochrome-conjugated monoclonal antibodies. 2.3

Equipment

1. Analytical balance. 2. Biosafety cabinet with vacuum suction. 3. Incubator shaker. 4. Centrifuge with 50 mL conical inserts. 5. Brightfield microscope. 6. CO2 incubator. 7. Microcentrifuge for 1.5 mL tubes. 8. Magnetic stir plate. 9. pH meter. 10. Orbital shaker. 11. Microplate reader.

2.4 Media and Solutions

All the solutions are filtered through a Stericup vacuum filtration system. 1. 0.1% Collagenase solution: Weigh 0.1 grams of type I collagenase and place it in a 50 mL conical tube; add 1 gram of powdered bovine serum albumin (BSA). Dissolve these in 50 mL phosphate-buffered saline (PBS). Next, filter with a 250 mL Stericup and wash the filter with another 50 mL of PBS (see Note 1). The volume of the collagenase solution used will be equal to the volume of the lipoaspirates (1:1). This solution can be stored at 4 °C for 1–2 weeks. 2. Stromal medium: Remove 50 mL of medium from a 500 mL bottle of DMEM/Ham’s F-12 medium, add 50 mL of heatinactivated fetal bovine serum (10%) and 5 mL of antibioticantimycotic 100× stock solution (1%). Use a 500 mL Stericup for filtration. This stromal medium should be used within 4 weeks of its preparation (see Note 2).

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3. AdipoQual adipogenic differentiation medium: The differentiation medium used for adipogenesis of hASCs is provided by AdipoQual (Obatala Sciences, Inc., New Orleans, LA, USA). 4. Osteogenic differentiation medium: The following stock solutions must be prepared and stored at -20 °C until ready for use. (a) A 1 mM stock solution of dexamethasone (4000-fold concentration) dissolved in distilled water. (b) A 50 mM stock solution of L-ascorbic acid 2-phosphate (2000-fold concentration) dissolved in distilled water. (c) A 0.5 M stock solution of β-glycerophosphate must be made fresh in the stromal medium. (d) The osteogenic differentiation medium is prepared with 1 nM dexamethasone, 50 μM L-ascorbic acid 2-phosphate, and 20 mM β-glycerophosphate in the stromal medium. The differentiation medium can be stored at 4 °C for up to 3 weeks. 5. Cell detachment: 0.25% Trypsin-EDTA (Thermo Fisher Scientific). 6. Oil Red O staining solution: Weigh out 0.5 g Oil Red O in a 100 mL borosilicate glass storage bottle. Dissolve in 100 mL isopropanol. Store at room temperature as a stock solution (see Note 3). At the time of use, take 3 parts of Oil Red O stock solution. Add 2 parts distilled water. Filter with a 250 mL Stericup. The working solution must be used within 2 h before precipitate builds up. 7. Alizarin Red staining solution: Weigh 0.5 g of alizarin red in a borosilicate glass storage bottle. Add 50 mL of distilled water and place on a stir plate with a magnetic stirrer for 2 min. Use a pH meter to adjust the pH of the solution to a range between 4.1 and 4.4. Increase the pH to the appropriate range with ammonium hydroxide or decrease it with hydrochloric acid, carefully adding small drops at a time. Cover the bottle with foil and store for up to 3 months. Filter the solution with a 50 mL filtration unit before use. 8. Crystal violet staining solution: Weigh 3 g of crystal violet in a clean disposable plastic container with a screw-top lid. Add 100 mL of methanol and stir on a stir plate with a magnetic stir bar for 5 min. 9. Freezing medium: The freezing medium comprises 90% heatinactivated fetal bovine serum and 10% dimethyl sulfoxide (DMSO). Use a freshly made freezing medium for cryopreservation.

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10. Medium sterility test procedure: Maintain 10 mL of the medium in a 10 cm2 tissue culture plate alongside your cultures to ensure the sterility of the medium. Everyday, check the plate using a brightfield microscope for any evidence of contamination, which can appear as cloudiness in the stromal medium or very apparent circular colonies with the naked eye and are more evident as bacterial or fungal colonies under the microscope. Immediately add 15% bleach solution to all contaminated plates if any contamination is observed and discard.

3

Methods The lipoaspirate sample should be processed as close to the acquisition time as possible and left no longer than 24 h at room temperature before cells are isolated. Before beginning the protocol, turn on the shaking incubator and set it to 37 °C at 100 rpm. Before transferring them into the biosafety cabinet, all materials must be cleaned with 70% ethanol. All isolation procedures are conducted in a biosafety cabinet by investigators trained in handling human tissue (see Note 4).

3.1 Processing Liposuction Aspirates

1. Wipe the biosafety cabinet working area with 70% ethanol and clean the collagenase solution, PBS, and stromal medium bottles before placing them in the biosafety cabinet. 2. Clean the outside of the lipoaspirate container with 70% ethanol and open it under the biosafety cabinet. 3. Pour up to 100 mL of the lipoaspirate into a 250 mL polypropylene beaker. 4. Add an equal volume of PBS to the lipoaspirate, and allow the fat and blood layers to separate for 3–5 min. The fat layer will float to the top and the blood layer will collect at the bottom. Next, aspirate the lower layer with a glass Pasteur pipette, careful not to aspirate any top fat later (see Note 5). Repeat the wash until the lipoaspirate loses its bloody appearance. 5. Divide the clean adipose tissue into 50 mL conical tubes by adding 15 mL of tissue to each conical. 6. Add an equal volume (15 mL) of collagenase solution to the 50 mL conical tubes and secure the caps with parafilm. Invert the conical tubes a few times, then secure them in the prepared shaking incubator for 1 h at 100 rpm. 7. Isolation of stromal vascular fraction (SVF): After 1 h of digestion, transfer the conical tubes back to the biosafety cabinet. Neutralize the digestion solution with 15 mL of stromal medium. Centrifuge the neutralized mixture at 300 × g (1200 rpm) for 5 min at room temperature.

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8. Carefully pour the fat layer into a disposable cup. Aspirate the remaining supernatant, leaving approximately 3 mL and being careful not to aspirate the resulting SVF pellet (see Note 6). 9. Resuspend the cells in the remaining supernatant and filter through a 100 μm cell strainer into a 50 mL conical tube. (a) Add PBS to the filtrate and adjust the final volume to ~25 mL. (b) Centrifuge at 300 × g (1200 rpm) for 5 min. 10. Aspirate the supernatant and resuspend the pellet in 5 mL of stromal medium. Filter the suspension through a 70 μm cell strainer into a new 50 mL conical tube. Rinse the filter with an additional 2–3 mL of stromal medium. 11. Plate the cells using 175 cm2 cell culture flasks in the stromal medium. Add approximately 3 mL of cell suspension per flask. 12. Twenty-four hours after plating, aspirate the medium from the flask. Wash the flask with 1× PBS and refeed with the stromal medium. The medium is then changed every 3 days until the cells are at 70–80% confluence. 13. Remove the medium from the flasks and add 4 mL of 0.25% Trypsin-EDTA. Place in a CO2 incubator for 3 min. Verify cell detachment under a microscope, and then deactivate the trypsin with double the volume of the stromal medium (see Note 7). 14. Transfer the cell suspension into a 15 mL or 50 mL conical. Centrifuge for 5 min at 180 × g. Aspirate the supernatant and resuspend the cell pellet in 2–6 mL stromal medium. 15. Use a 1:1 dilution of cell suspension and Trypan Blue (see Note 8). Then, count the cells with a hemocytometer. 16. The following are suggested next steps for characterization and expansion. 3.2

Cryopreservation

1. Ensure Mr. Frosty, or a similar freezing container, is filled with 250 mL of isopropanol. 2. Gather cryogenic vials and freezing medium. 3. Centrifuge the cell suspension at 180 × g for 5 min and resuspend in the freezing medium at a concentration of at least 5 × 105 cells/mL and up to 1 × 106 cells/mL. 4. Place 1 mL of the cell suspension into each cryovial, then place the labeled cryovials into Mr. Frosty and freeze at -80 °C overnight. 5. Transfer cryovials to a liquid nitrogen container within 72 h (see Note 9).

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3.3 Seeding Cells for Expansion

1. To expand the number of ASCs at subsequent passages, use a seeding density of 400 cells/cm2. 2. Seed 175 cm2 cell culture flasks with approximately 7 × 104 cells per flask. 3. When cells reach 70–80% confluence, then freeze or split into further passages.

3.4 Characterization of hASCs

Flow cytometry: 1. Spin trypsinized cells at 12,000 rpm for 5 min and decant the supernatant. 2. Add 1 mL of PBS at room temperature and transfer to a 1.5 mL microcentrifuge tube. 3. Centrifuge at 3000 rpm for 3 min and decant the supernatant. 4. Add 400 μL of human FC block and incubate at room temperature for 10 min. 5. Prepare amber 1.5 mL microcentrifuge tubes with 1 μL of fluorochrome-conjugated monoclonal antibodies, ensuring to leave one without antibody as a control. 6. Aliquot 100 μL of cell suspension into each amber microcentrifuge tube and vortex. Incubate at room temperature for 15 min. 7. Wash cells with 1 mL of PBS per tube and centrifuge at 3000 rpm for 3 min. 8. Decant the supernatant and add 500 μL of 1% paraformaldehyde for fixation. 9. Store the samples at 4 °C until flow cytometry can be conducted (see Note 10). Adipocyte differentiation and staining: 1. Seed a 6-well cell culture plate with 2 × 105 cells/well and a 12-well cell culture plate with 8 × 104 cells/well with the stromal medium. 2. Once cells are at 70% confluence, they can be induced for differentiation using AdipoQual adipogenic differentiation medium. 3. Remove the stromal medium and add AdipoQual, leaving at least one well per plate as undifferentiated control. 4. Change the AdipoQual every 3 days and differentiate for 21 days. 5. Remove the medium from all the wells and rinse once with PBS. 6. Fix with 4% paraformaldehyde (PFA) for 1 h.

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7. Rinse with distilled water and add enough Oil Red O to the working solution to cover the bottom of the wells. 8. Place the plate on an orbital shaker for 20 min. 9. Rinse with distilled water until the wells run clear and image immediately. 10. Let the plate dry overnight with the lid off. 11. To quantify the extent of Oil Red O staining, elute the stain from the wells by adding 500 μL of isopropanol to each well and placing it on an orbital shaker for 20 min. 12. Transfer 100 μL per well into a 96-well plate in duplicate. 13. Read the OD 584 using a microplate reader to determine the relative stain intensity of osteocytes compared to non-differentiated controls. Osteocyte differentiation and staining: 1. Seed 12-well cell culture plate with 8 × 104 cells/well in stromal medium. 2. Once cells are at 70% confluence, they may be induced for differentiation using an osteogenic differentiation medium. 3. Remove the stromal medium and add an osteogenic differentiation medium, leaving at least one well as an undifferentiated control. 4. Change the medium every 3 days and differentiate for 28 days. 5. Collect the plates after rinsing them once with PBS. Fix the cells with 4% PFA for 1 h. 6. Rinse the wells with distilled water. 7. Add enough Alizarin Red stock solution to cover the bottom of the well and place it on an orbital shaker for 30 min. 8. Rinse the cells with distilled water until the wells clear and image immediately. 9. Let the plate dry overnight with the lid off. 10. To quantify Alizarin Red staining, make 10% Cetylpyridinium chloride (CPC) by dissolving 1 g of CPC into 10 mL of distilled water under a chemical fume hood. CPC is stored with parafilm and kept under the fume hood. 11. Elute the alizarin red by adding 300 μL of 10% CPC to each well and place it on an orbital shaker for 1 h. 12. Transfer 100 μL per well to a 96-well plate in duplicate. 13. Read the OD 544–584 using a microplate reader. Compare the staining intensity of the adipocytes to the undifferentiated controls.

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Colony-forming unit-fibroblast (CFU): 1. Seed a 6-well plate with 1000 cells in each well using the stromal medium. 2. After 7 days, change the stromal medium. Rinse the wells with PBS after 14 days. 3. Add enough 3% crystal violet solution to cover the bottom of the wells and place it on an orbital shaker for 20 min. 4. Next, rinse the wells with distilled water until the wells run clear. 5. Leave the plate uncovered to dry overnight. 6. The following day, count each colony larger than 2 mm in diameter.

4

Notes 1. Washing the filter with 50 mL of PBS will yield 100 mL of the collagenase solution at the desired concentration (0.1%). 2. Place the bottle in a 50 °C water bath for 30 min to heat inactivate FBS. 3. The Oil Red O stock solution is good for 1 year; however, accumulating a precipitate is a significant issue. Therefore, when making working solutions, it may be necessary to undergo two filtration steps with a Stericup. Alternatively, a 50 mL filtration unit may make working solutions much smaller in volume. 4. Proper training for blood-borne pathogens, contaminants, and biological hazards should be completed before working with human tissues. 5. Alternatively, a sterile disposable serological pipette may remove the bottom layer. 6. The resulting pellet after centrifugation is the SVF from which hASCs are isolated. 7. If less than 90% of cells appear detached, tap the sides of the flask with your palm to encourage further detachment of the cells. 8. An aliquot of 10–20 μL of cells and 10–20 μL of Trypan Blue may be used to achieve the 1:1 dilution. 9. The cell count determines the number of cryovials to be frozen. Therefore, for earlier passages (P0 and P1), it is best to freeze as many cryovials as possible while maintaining the appropriate cell concentration. For example, freezing 8 vials of 5 × 105 cells/mL is preferable to only freezing 4 at a concentration of 1 × 106 cells/mL.

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10. Use 100 μL of FC block reagent per antibody panel. The number of amber microcentrifuge tubes depends on what monoclonal antibodies are chosen and whether the fluorochromes are read at different wavelengths. References 1. Gimble JM, Katz AJ, Bunnell BA (2007) Adipose-derived stem cells for regenerative medicine. Cir Res 100(9):1249–1260 2. Al-Ghadban S, Bunnell BA (2020) Adipose tissue-derived stem cells: immunomodulatory effects and therapeutic potential. Physiology (Bethesda) 35(2):125–133 3. O’Donnell BT, Al-Ghadban S, Ives CJ et al (2020) Adipose tissue-derived stem cells retain their adipocyte differentiation potential in threedimensional hydrogels and bioreactors. Biomol Ther 10(7):1070 4. Li Z, Lin Z, Liu S et al (2022) Human mesenchymal stem cell-derived miniature joint system for disease modeling and drug testing. Adv Sci (Weinh) 9(21):e2105909 5. Yoshimura K, Shigeura T, Matsumoto D et al (2006) Characterization of freshly isolated and

cultured cells derived from liposuction aspirates’ fatty and fluid portions. J Cell Physiol 208(1): 64–76 6. Li J, Curley L, Floyd EZ, Wu X et al (2018) Isolation of human adipose-derived stem cells from lipoaspirates. Methods Mol Biol 1773: 155–165 7. Harrison MAA, Al-Ghadban SI, O’Donnell BT et al (2022) Establishing the adipose stem cell identity: characterization assays and functional properties. In: Kokai L, Marra K, Rubin JP (eds) Scientific principles of adipose stem cells. Elsevier 8. Zuk PA, Zhu M, Ashjian P et al (2002) Human adipose tissue is a source of multipotent stem cells. Mol Biol Cell 13(12):4279–4295

Chapter 2 Isolation of Human Adipose-Derived Stromal/Stem Cells from Lipoaspirates Yuejia Li, Ziyi Mei, Jie Li, and Jeffrey M. Gimble Abstract Adipose tissue is an abundant and accessible source of stem cells with multipotent properties suitable for tissue engineering and regenerative medical applications. Adipose-derived stromal/stem cells (ASCs) have been widely used in tissue engineering and cell therapy. In addition, the clinical application of ASCs in the treatment of inflammation and injury has been proven a success. Here, we describe methods from our own laboratory and the literature for the isolation and expansion of Adipose-derived stromal/stem cells (ASCs). We present a large-scale procedure suitable for processing >100 mL volumes of lipoaspirate tissue specimens by collagenase digestion, a related procedure suitable for processing adipose tissue aspirates without digestion, and a procedure suitable for intact human adipose tissue, such as buccal fat pads in the maxillofacial region. Key words Adipose-derived stromal/stem cells (ASCs), Biopsy, Collagenase, Expansion, Human, Isolation, Lipoaspirate, Mesenchymal stromal/stem cells (MSCs), Stromal vascular fraction (SVF)

1

Introduction Mesenchymal stromal/stem cells (MSCs) were initially described in bone marrow and have been found subsequently in multiple tissues, including subcutaneous adipose tissue [1–4]. Recently, mesenchymal stromal/stem cell-based therapy became a key focus of regenerative medicine [5, 6]. In mammals, adipose tissue is mainly divided into white adipose tissue, which specifically stores energy, and brown adipose tissue, which maintains body temperature [7]. It has been demonstrated that there are brown adipocyte-like cells in White Adipose Tissue (WAT), which are called beige or brite adipocytes [8]. Although adipose-derived stromal cells had been termed “pre-adipocytes” [2, 3], multiple independent investigators have demonstrated that they are multipotent, with adipogenic, chondrogenic, neuronal-like, and osteogenic differentiation capability [9–12]. Consequently, they have now been identified as

Jeffrey M. Gimble et al. (eds.), Adipose-Derived Stem Cells: Methods and Protocols, Methods in Molecular Biology, vol. 2783, https://doi.org/10.1007/978-1-0716-3762-3_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Adipose-derived stromal/stem cells or ASCs [13]. This protocol, which includes information presented in earlier volumes of this series [14], describes the isolation of primary in-vitro cultures of ASCs from human adipose tissue. After isolation, the quality of ASCs are assessed by characterization of self-renewal ability, proliferation rate, and multi-lineage differentiation potential [6]. Furthermore, we introduced the isolation of ASCs from Buccal Fat Pad (BFP), which excision has become a more common aesthetic surgical procedure [15, 16]. BFP removed in maxillofacial surgery is also a reliable source of ASCs.

2 2.1

Materials Supplies

1. Tissue: Subcutaneous adipose tissue samples obtained from liposuction aspirates (see Note 1). 2. 200 mL plastic centrifugation bottles (Nalgene). 3. 0.2 mm filter units. 4. 50 mL conical tubes. 5. 2 mL tubes. 6. Hematocytometer. 7. Freezing apparatus (Mr. Frosty™ Freezing Container). 8. Fluorochrome-conjugated monoclonal antibodies against stromal (CD44, CD73, CD90, CD105), hematopoietic (CD27, CD34, CD38, CD48, CD117, CD150), endothelial (CD31, CD34, CD45, CD54), pericyte (CD31, NG2, PDGFR beta, CD146, Nestin), and related cell surface antigens.

2.2

Equipment

1. Inverted microscope—Nikon Eclipse TS100 with Epi-Fluorescence Attachment (Mercury Lamp Illuminator model name: C-SHG) (Nikon Instruments Incorporation, Melville, NY) and equipped with a camera photometric coolsnap (Nikon). 2. MetaMorph Imaging Corporation).

Software

(Universal

Imaging

3. Shaking water bath. 4. Centrifuge. 5. Biosafety hood. 6. CO2 incubator. 2.3 Media (See Note 2)

All the media solutions are filtered through a 0.2 mm filter unit (see Note 3). 1. 0.1% Collagenase solution: Weigh out 0.1 g of type I collagenase and 1 g of powdered bovine serum albumin (fraction V).

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Dissolve these in 100 mL of phosphate-buffered saline (PBS) supplemented with 2 mM calcium chloride (CaCl2). After sterile filtration, warm the solution to 37 °C. This solution should be used within 1 h of preparation. 2. Erythrocyte lysis buffer: Prepare 155 mM ammonium chloride (NH4Cl), 10 mM potassium carbonate (K2CO3), and 0.1 mM EDTA using sterile, distilled water. This solution should be used within 24 h of its preparation. 3. Stromal medium: Add 55 mL of 10% fetal bovine serum and 5.6 mL of 100X antibiotic (penicillin/streptomycin)/antimycotic (amphotericin) to 500 mL of DMEM/Ham’s F-12 medium. This solution should be used within 4 weeks of its preparation. All fetal bovine serum should be pre-screened prior to purchase for its ability to support both cell proliferation and adipocyte differentiation. 4. Adipogenic differentiation medium: Prepare the differentiation medium containing the following final concentrations in DMEM/Ham’s F-12: 3% fetal bovine serum, 0.25 mM IBMX, 66 μM biotin, 34 μM D-panthothenate, 5 μM rosiglitazone (or equivalent PPARγ2 ligand), 1 μM dexamethasone, 200 nM human insulin. Use this solution within 2 weeks of its preparation. In advance, prepare and aliquot the following stock solutions: 66 mM biotin (2000-fold concentration) dissolved in 1 N sodium hydroxide, 34 mM solution D-pantothenate (2000-fold concentration) dissolved in water, 1 mM dexamethasone (1000-fold concentration) dissolved in water or ethanol depending on its formulation, 250 mM solution of methylisobutylxanthine (1000-fold concentration) dissolved in dimethyl sulfoxide, 200 mM solution of human insulin (2000-fold concentration) dissolved in PBS, 5 mM stock solution of rosiglitazone or equivalent PPARg agonist dissolved in dimethyl sulfoxide. Store frozen at -20 °C until required within 2 weeks. 5. Adipocyte maintenance medium: This solution is prepared in an identical manner as differentiation medium except that it does not contain either the isobutylmethylxanthine or the PPARγ agonist; these two stock solutions should be omitted. Use this solution within 2 weeks of its preparation. 6. Oil Red O staining stock solution: Weigh out 0.5 g Oil Red O. Dissolve in 100 mL isopropanol. Filter through a 0.2 μm filter. Store at room temperature as stock solution. 7. Oil Red staining working solution: At the time of use, take 6 mL of Oil Red O stock solution. Add 4 mL of distilled water. Stand 1 h at room temperature before use. Use this solution within 24 h of its preparation.

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8. Freezing medium: The freezing medium consists of 80% fetal bovine serum or 80% calf serum, 10% DMEM/Ham’s F-12, and 10% dimethyl sulfoxide. Use this solution within 2 weeks of its preparation.

3

Methods After transportation to the laboratory, the liposuction sample can be kept at room temperature for no more than 24 h prior to use. Before performing the experiment, warm up the water bath to 37 °C. All the following procedures are performed in biosafety hoods. Investigators should be trained and certified in the handling of human tissues and human pathogens prior to initiation of any studies (see Note 4).

3.1 Digestion and Cell Isolation from >100 mL Liposuction Aspirate Samples

1. Warm up the buffer (500 mL or more of PBS or KRB). Line the surface of the biosafety hood with a disposable bench protector. 2. Warm up the freshly prepared collagenase solution in the 37 °C water bath. The volume of the collagenase solution used will be equal to the volume of the lipoaspirates (1:1). 3. Prepare the PBS (or KRB) solution with 1% BSA, filter the solution, and warm it in the 37 °C water bath. 4. To maintain optimal sterile conditions, open the surgical container used for the liposuction procedure under the biosafety hood (see Note 5). Dispense a volume of adipose tissue in sterile plastic bottles for each 175 cm2 flask (0.16 mL tissue/cm2). It is recommended that you distribute about 33 mL of tissue; each bottle can accommodate ~100 mL of tissue. We routinely process a total of 200 mL of tissue to be plated in six 175 cm2 flasks. Add an equal volume of warm PBS. Agitate to wash the tissue and then allow phase separation for 3–5 min. Alternatively, you can spin the samples at 300× g in an appropriate centrifuge for 1 min at room temperature. Suction off the infranatant solution (lower liquid phase) while retaining the supernatant solution containing the floating tissue. The wash is repeated several times until a clear infranatant solution is obtained (usually 3–4 times). 5. Add an equal volume (60–70 mL) of warm collagenase solution into the plastic bottles containing the clean adipose tissue sample. Wrap the bottles with parafilm and place them in a 37 ° C shaking water bath at ~200 rpm for 60–80 min until the tissue appears smooth upon visual inspection (see Note 6). 6. After digestion, spin the samples at 300× g in an appropriate centrifuge for 5 min at room temperature. Take the samples out of the centrifuge and shake them vigorously to thoroughly

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disrupt the pellet and to mix the cells. This is to complete the separation of the stromal cells from the primary adipocytes. Repeat the centrifugation step. 7. After spinning, the stromal vascular fraction will form a pellet at the bottom of the bottle or tube (this will usually include a layer of dark red cells). Carefully remove the top layer of oil and fat, the primary adipocytes (a yellow layer of floating cells), and the underlying layer of collagenase solution. Leave behind a small volume of collagenase solution above the pellet so that the cells are not disturbed. 8. Resuspend the cells in 10 mL of warm PBS (or KRB) solution and transfer the solution containing the cells into a 50 mL conical tube. Centrifuge the cells at 300× g in an appropriate centrifuge for 5 min at room temperature. 9. Aspirate the remaining collagenase solution. When aspirating, the tip of the pipette should aspirate from the top so that the oil is removed as thoroughly as possible. The cell pellet should be at the bottom of the tubes. 10. Resuspend the cells in 15 mL of stromal medium (see Note 7). Divide the cells according to the number of flasks. The cells are plated at a density equivalent to approximately 0.18 mL of liposuction tissue aspirate per cm2 of surface area (volume of ~33 mL of tissue for a 175 cm2 flask). Divide the cells according to the number of flasks. In this protocol, we use about 200 mL of liposuction tissue. Thus, to each 175 cm2 flask (×6), we add 2.5 mL of cell suspension and 32.5 mL of stromal medium (see Note 8). 11. Forty-eight hours after plating, aspirate the medium from the flask. Wash the cells with prewarmed PBS (see Note 9). Add 35 mL of fresh stromal medium (see Note 10). 3.2 Harvesting the Isolated Cells

1. Remove the medium from the flasks and save the sterile “conditioned media” in a sterile tube for future cell culture application (this should be sterile filtered prior to such use and can be cryopreserved). Add 10 mL of sterile warm PBS to the flasks and allow the PBS to remain on cells for 2 min while the flasks are in a horizontal position. Remove the PBS by aspiration and replace the PBS with 5 mL of 0.5% Trypsin/EDTA solution (see Note 11). Incubate in the incubator for 5–10 min. Verify under microscope that more than 90% of the cells have detached and then add 5–10 mL of stromal medium to allow the serum contained in the solution to neutralize the trypsin reaction. 2. Proceed to cell counting by taking an aliquot of cells diluted in trypan blue (for a 1:1 dilution: add 10 μL of suspended cells to 10 μL of trypan blue). Count the cells using the hematocytometer.

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1. Suspend the cell pellet in a room temperature freezing medium at a concentration of 1–2 × 106 cells per mL.

3.3 Cryopreservation of the Isolated Cells

2. Dispense 1 mL aliquots of the cell suspension to sterile cryovials. 3. Place the cryovials in the appropriate freezing apparatus. Freeze the cells to -80 °C overnight. 4. The next day, transfer the cells on dry ice or other frozen material to a liquid nitrogen storage container. 1. Harvest ~1.5 × 106 ASCs and centrifuge at 300× g for 5 min at room temperature in a 50 mL tube.

3.4 Flow Cytometry Analysis of the Isolated Cells

2. Wash the ASCs twice with 10 mL cold PBS (Ca and Mg free) and resuspend the cells in 500 μL cold PBS. 3. Aliquot 50 μL of the cells into ten 1.5-mL microcentrifuge tubes. 4. Add 50 μL volume of PBS containing a fluorochromeconjugated monoclonal antibody or isotype control antibody (usually 5–10 μL) to each tube and mix well. 5. Incubate the samples for 20–30 min at room temperature according to the manufacturer’s instructions (see Note 12). 6. Wash the cells with 1 mL PBS with 1% BSA and pellet the cells at 300× g for 3 min at room temperature. Repeat this step three times. 7. Resuspend the cells in 500 μL of 1% formaldehyde in PBS to fix the cells. 8. Keep the tubes at 4 °C protected from light exposure until they can be analyzed on a flow cytometer within a 48 h period. 1. After cell count, suspend the cell pellet in stromal medium following the different concentrations listed in Table 1 to achieve a confluent culture within 24 h of re-plating.

3.5 Replating the Isolated Cells

Table 1 Table for plating Plate

Area per plate 2

6-well plate

60 cm

24-well plate

48 cm2

96-well plate

2

31 cm

Cells per plate

Cells per well

Media per well

1.8 × 10

30 × 10

2.5 mL

6 × 104

1 mL

6

4

1.44 × 106 0.93 × 10

6

10

4

200 μL

Isolation of hASCs from Lipoaspirates

3.6 Adipogenic Differentiation, Fixation, and Staining of the Isolated Cells

19

1. When the cells reach between 80% and 90% confluence (before or after harvesting the cells), the ASCs are induced to differentiate. 2. Aspirate the medium, add a small volume (about 1.5 mL for a 6-well plate) of prewarmed PBS + 1% antibiotic to wash the cells, and finally remove the PBS by aspiration (see Note 13). 3. Add the differentiation medium. 4. The cells will be maintained in the differentiation medium for 3 days (Day +3 differentiation). 5. Aspirate the differentiation medium and wash the cells with prewarmed PBS + 1% antibiotic (see Note 14). Then add a volume (2.5–3 mL for a 6-well plate) of adipocyte maintenance medium. 6. Change the adipocyte maintenance medium every 3 days until mature adipocytes are obtained (Day +9 to +12 differentiation) (see Note 15). 7. After 12 days of differentiation, the cells can be fixed either using 10% formalin solution, 4% paraformaldehyde, or 70% ethanol (see Note 16). After removing the medium and washing the cells with PBS, immerse the cells in the fixative solution: 10% formalin, 4% paraformaldehyde, or 70% ethanol for 30 min, 10 min, or 1 h, respectively. 8. Remove the fixative before staining (fixed cells can be stored at 4 °C for as long as several months although shorter times are recommended). 9. Add 50 μL Oil Red O to each well for 15 min at room temperature. 10. Rinse 3× or more with 50 μL distilled water (see Note 17). 11. Elute the stain from the cells by adding 50 μL isopropanol per well. Elution is immediate. 12. Read the OD540 using a plate reader. 13. Subtract the background staining determined in blank wells (no cells) from the experimental points. 14. Determine the relative staining intensity of the differentiated wells compared to the preadipocyte controls.

3.7 Digestion and Isolation of Cells from Lipoaspirate Fluid (See [17])

1. Aspirate the lipoaspirate fluid fraction using pipette, separating it from the lipoaspirate tissue fraction. 2. Transfer the lipoaspirate fluid (LAF) to a sterile plastic bottle. 3. Centrifuge for 10 min at 400× g at room temperature. 4. Resuspend the LAF cell pellet in erythrocyte lysis buffer. Stand at room temperature for 5 min. 5. Filter the LAF cell suspension through 100 μm filter.

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6. Load the cells onto a Ficoll density gradient. Centrifuge for 20 min at 800× g at room temperature. 7. Aspirate the cells from gradient interface. 8. Wash with PBS prewarmed to 37 °C. 9. Filter the LAF cell suspension through a 100 μm filter. 10. Count the nucleated cell number using trypan blue staining and hemocytometer. 11. Suspend the LAF cells in stromal medium. Plate at density of 6.4 × 104 cells/cm2 on gelatin coated T175 cm2. Maintain in culture and proceed as described under Subheadings 3.2, 3.3, 3.4, 3.5, and 3.6 (see Note 18). 3.8 Digestion and Isolation of Cells from the Buccal Fat Pad

4

1. Wash the buccal fat pad (BFP) twice with PBS and absorb the excess PBS. 2. Transfer the BFP to a dry container and mince into small pieces. The subsequent steps are as described under Subheadings 3.2, 3.3, 3.4, 3.5, and 3.6.

Notes 1. Prior to the implementation of this protocol, all personnel involved in the processing of human lipoaspirate material or primary ASC cultures should complete safety training for the use of blood-borne pathogens. Regardless of whether tissue donors have been screened for evidence of infection by hepatitis, HIV, or other transmissible agents, these precautions are mandatory. In addition, no glass containers or pipettes should be used and the use of sharp objects (scissors, needles) during the processing steps should be minimized. All procedures involving the human tissue or cells should be conducted in a biological safety cabinet and with appropriate personnel protection gear. 2. Before purchase, the fetal bovine serum should be assayed to test for its ability to support robust cell proliferation and adipogenesis. 3. Medium sterility test procedure: Prior to use, it is wise to test the sterility of the medium by removing a single milliliter from each bottle, placing it in a single well of a 24-well plate, and incubating it for 48 h in a humidified, 37 °C, CO2 incubator. After this period, examine the plate using a phase contrast microscope for any evidence of contamination. If contaminated, immediately inactivate all bottles and test plates with 15% bleach solution and discard. 4. Insuring sterility and minimizing microbial contamination: In order to keep optimal sterile conditions, it is recommended

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that you open and close the container properly to avoid any potential contaminations. It is vital that the culture be examined regularly to confirm the absence of microbial contamination. To avoid this problem, 5% iodine solution can be added to the initial wash solution. The disinfectant is then washed away in subsequent washes. In order to avoid this problem in small adipose tissue samples, it is recommended to add in the PBS solution 1% of antibiotic solution and wash thoroughly the tissue with this solution. 5. Cellular heterogeneity within ASC cultures: The blood cells could be a source of contamination and may reduce or prevent adhesion of stromal cells; it is then important to wash thoroughly the cells with PBS (1% of antibiotic solution can be added). When removing the PBS from the cells, aspirate the solution up and down until the cells appear clean and free of red cells. An erythrocyte lysis buffer (155 mM NH4Cl, 5.7 mM K2HPO4, 0.1 mM EDTA at pH 7.3) can serve to remove red blood cells. Endothelial cells (EC) could be another source of contamination. Intra-abdominal depots are more subject to this type of contamination when compared to subcutaneous adipose tissue, which is relatively free of EC. Therefore, a filtration procedure can be performed by using a nylon mesh filter with a small pore size (25 mm). 6. After 1 h digestion, if pieces of undigested tissue are still observed in the tube, make sure that the collagenase solution is fresh and has not been maintained at room temperature for an extended period. This is necessary to maximize the enzyme efficiency. The collagenase solution can also be stored at -20 ° C for a few days, with a minor loss of enzyme activity. Prior to use, the frozen solution is slowly thawed at room temperature and prewarmed to 37 °C. 7. A filtration procedure can be performed by using a nylon mesh filter with a small pore size (250–350 mm). The suspension is then centrifuged at 300× g at room temperature to allow separation of the stromal vascular fraction from the mature adipocytes. The filtration can be performed after removing the floating mature adipocytes cells. This step will remove tissue fragments and is used by some investigators. However, the filtration procedure is not recommended for small amounts of adipose tissue. 8. To accelerate cell adhesion, the culture dishes can be precoated with extracellular matrix proteins, such as gelatin or Matrigel. 9. If the cells do not grow very well and do not appear healthy, there are a few options. The percent of preadipocytes obtained from the stromal vascular fraction after digestion is patient-dependent. Supplementation of the culture medium

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with 20–30% conditioned medium (saved from previous cultures) should facilitate the growth of the cells. If you do not have conditioned medium, another alternative is to increase the FBS contained in the stromal medium to 15–20%; however, this may promote premature adipogenesis. Signs of deterioration such as granularity around the nucleus, cytoplasmic vacuolations, and/or detachment of the cells from the plastic surface may indicate inadequate or toxic medium, microbial contamination or senescence of the primary cells. 10. This period can vary from 24 to 72 h depending on the number of cells attached to the plastic surface (observed under microscope). The medium is then changed every 2–3 days until the cells achieve 80–90% confluence. 11. It is important to not overexpose the cells to the trypsin/ EDTA solution. This could decrease the cell viability. 12. Keep the tubes protected from light exposure to avoid bleaching of the fluorochrome. This can be accomplished by wrapping the tubes in aluminum foil. 13. If the adipocytes are detaching, do not dry the well when changing the medium since adipocytes tend to float when new medium is added. 14. According to the protocol, the medium is changed every 2–3 days. However, as the adipocytes mature, you may observe a yellowing of the culture medium: a drop in pH may account for this. As the pH falls from 7 to 6.5, cell growth will decline and cell viability falls at pH between 6.5 and 6. You can observe this change of pH by looking at the medium color change, going from red (pH 7) through yellow (pH ≤ 6), indicating the need for an immediate change of the medium. 15. If the cells do not differentiate very well, consider that the differentiation process may be patient dependent. The age of the donor can be a factor, since some studies suggest that the differentiation capacity is higher in culture from younger subjects compared to older people. To further enhance adipogenesis, the following alternatives are proposed: • You may try different PPARg agonists (troglitazone, pioglitazone among others). • The addition of 5% rabbit serum (RS) can be added to the differentiation medium enhance differentiation (the ethyl acetate contain in the RS has been found 35-fold more abundant than in FBS [18]. • Another alternative would be to perform the addition of the differentiation medium multiple times after a 3-day rest period; i.e., 3 days on in the presence of the differentiation

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medium and 3 days off in the presence of the adipocyte medium. Repeat this cycle until mature adipocytes are obtained. 16. Using ethanol, there is a risk that the lipids will be eluted from the cells. 17. The rinse should become completely clear (no red coloring). Do not rinse with a volume larger than 50 μL. It will raise the level of the solution in the plastic well and cause staining of the wall of the well, resulting in artifactually high background. 18. Ultrasound should be avoided in liposuction, which might lead to the reduction of the number of cells recovered from tissue digests and the proliferative capacity [12].

5

Final Comments Recently, there has been increased appreciation for the use of primary cell culture models in the study of human adipocyte differentiation in vitro. This protocol on the cultivation of human adipocyte precursor cells can be used by laboratories with access to human tissues in a scalable manner. Very small volumes of tissue can be cultured using tissue explants as described in Chap. 14.

References 1. Rangwala SM, Lazar MA (2000) Transcriptional control of adipogenesis. Annu Rev Nutr 20:535–559 2. Deslex S, Negrel R, Vannier C et al (1987) Differentiation of human adipocyte precursors in a chemically defined serum-free medium. Int J Obes 11:19–27 3. Hauner H, Entenmann G, Wabitsch M et al (1989) Promoting effect of glucocorticoids on the differentiation of human adipocyte precursor cells cultured in a chemically defined medium. J Clin Invest 84:1663–1670 4. Halvorsen YD, Bond A, Sen A et al (2001) Thiazolidinediones and glucocorticoids synergistically induce differentiation of human adipose tissue stromal cells: biochemical, cellular, and molecular analysis. Metabolism 50:407– 413 5. Hoang DM, Pham PT, Bach TQ et al (2022) Stem cell-based therapy for human diseases. Sign Transduct Targ Ther 7:272 6. Al-Ghadban S, Bunnell BA (2020) Adipose tissue-derived stem cells: immunomodulatory effects and therapeutic potential. Physiology 35:125–133

7. Nahmgoong H, Jeon YG, Park ES et al (2022) Distinct properties of adipose stem cell subpopulations determine fat depot-specific characteristics. Cell Metab 34:458–472.e456 8. Li J, Curley JL, Floyd ZE et al (2018) Isolation of human adipose-derived stem cells from lipoaspirates. Methods Mol Biol:155–165 9. Zuk PA, Zhu M, Ashjian P et al (2002) Human adipose tissue is a source of multipotent stem cells. Mol Biol Cell 13:4279–4295 10. Zuk PA, Zhu M, Mizuno H et al (2001) Multilineage cells from human adipose tissue: implications for cell-based therapies. Tissue Eng 7: 211–228 11. Guilak F, Lott KE, Awad HA et al (2006) Clonal analysis of the differentiation potential of human adipose-derived adult stem cells. J Cell Physiol 206:229–237 12. Gimble JM, Katz AJ, Bunnell BA (2007) Adipose-derived stem cells for regenerative medicine. Circ Res 100:1249–1260 13. Mitchell JBMK, Zvonic S, Garrett S, Floyd ZE, Kloster A, Halvorsen YD, Storms RW, Goh B, Kilroy GS, Wu X, Gimble JM (2006) The immunophenotype of human adipose derived cells: Temporal changes in stromal- and stem

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cell-associated markers. Stem Cells 24:376– 385 14. Dubois SG, Floyd EZ, Zvonic S et al (2008) Isolation of Human Adipose-derived Stem Cells from Biopsies and Liposuction Specimens. Methods Mol Biol 449:69–79 15. Dehghani Nazhvani F, Mohammadi Amirabad L, Azari A et al (2021) Effects of in vitro low oxygen tension preconditioning of buccal fat pad stem cells on in Vivo articular cartilage tissue repair. Life Sci 280 16. Traboulsi-Garet B, Camps-Font O, TraboulsiGaret M et al (2021) Buccal fat pad excision for

cheek refinement: a systematic review. Med Oral:e474–e481 17. Yoshimura K, Shigeura T, Matsumoto D et al (2006) Characterization of freshly isolated and cultured cells derived from the fatty and fluid portions of liposuction aspirates. J Cell Physiol 208:64–76 18. Diascro DD Jr, Vogel RL, Johnson TE et al (1998) High fatty acid content in rabbit serum is responsible for the differentiation of osteoblasts into adipocyte-like cells. J Bone Miner Res 13:96–106

Chapter 3 Isolation of Perivascular Mesenchymal Progenitor Cells from Human Adipose Tissue by Flow Cytometry Neelima Thottappillil, Mario A. Gomez-Salazar, Mary Archer, Bruno Pe´ault, and Aaron W. James Abstract Perivascular cells represent an in vivo counterpart of mesenchymal stromal/stem cells that populate the outer layer of blood vessels. Pericytes in capillaries and microvessels and adventitial cells of large arteries and veins give rise to stem/progenitor cells when isolated and cultured in vitro. These cells have been considered candidate cell types for cell therapy. Adipose tissue, being highly vascularized, dispensable, and easily accessed, is a viable option to obtain perivascular cells for use in research and in clinical trials. Here, we describe our established protocol to extract perivascular cells from human fat through fluorescence-activated cell sorting, which allows for the isolation of defined populations of progenitor cells with high reproducibility. Key words Perivascular stem cell, Pericyte, Adventitial cell, Mesenchymal stromal/stem cell, Flow cytometry, Adipose tissue, Cell therapy

1

Introduction Presumptive mesenchymal stromal/stem cells (MSCs) are ubiquitously present in all human tissues, so as in blood vessels [1– 3]. Diverse perivascular cells enclose the endothelium in blood vessel anatomy [3]. Pericytes are in close contact with endothelial cells in capillaries, arterioles, and venules, and adventitial cells reside in the outermost layer of large arteries and veins [3, 4]. When isolated and cultured in vitro, these cells demonstrate the remarkable ability to (i) adhere to tissue culture plastic; (ii) differentiate to multiple mesodermal cell lineages; (iii) express markers like CD73, CD105, and CD90; and (iv) participate in tissue repair and regeneration. These cells can be collectively termed as perivascular stem/ progenitor cells (PSCs) [1, 5].

Authors Neelima Thottappillil and Mario A. Gomez-Salazar have contributed equally to this work. Jeffrey M. Gimble et al. (eds.), Adipose-Derived Stem Cells: Methods and Protocols, Methods in Molecular Biology, vol. 2783, https://doi.org/10.1007/978-1-0716-3762-3_3, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Adipose tissue is highly vascularized and represents an abundant source of perivascular mesenchymal progenitor cells [6]. Moreover, adipose tissue is readily available and can be obtained with minimal surgical intervention, posing low risk for donors [1, 6]. A clinically relevant number of PSCs can be easily isolated and even used without in vitro expansion [1, 7]. Several studies from our group indicate that PSCs isolated from adipose tissue display therapeutic benefit to treat bone fractures and osteoporosis [8–14]. Several studies indicate that perivascular progenitor cells have increased regeneration potential even at low numbers when compared to the total population [15–17]. Perivascular mesenchymal progenitors—adventitial cells and pericytes—are defined by immunophenotype, which helps isolate these cells from a heterogeneous progenitor population [1]. Our previous studies reported for the first time that pericytes can be prospectively isolated from adipose tissue as CD146+CD34-CD31-CD45cells [1]. Similarly, CD34+CD146-CD31-CD45- perivascular adventitial progenitor cells can also be purified [1, 12]. Therefore, the present chapter describes a standardized protocol to simultaneously isolate perivascular adventitial and pericyte progenitor cells. This is primarily achieved by careful enzyme-based digestion of human adipose tissue followed by advanced multicolor flow cytometry. Thus, the method described here allows for the isolation of much defined populations from a heterogeneous progenitor cell pool of major therapeutic potential.

2 2.1 2.1.1

Isolation of Adipose-Derived Perivascular Progenitor Cells by Flow Cytometry Material Equipment

1. Dissection tray. 2. Sterile fine forceps. 3. Sterile fine scissors. 4. Disposable sterile scalpels. 5. Sterile scoops. 6. Shaking water bath. 7. Sterile cell filters/strainers (22 μm, 100 μm, 70 μm, and 40 μm). 8. Centrifuge tubes (15 and 50 mL). 9. Eppendorf tubes (1.5 mL). 10. Filter capped FACS tubes. 11. Propylene tubes (5 mL). 12. Invitrogen Countess II automated cell counter. 13. Centrifuge. 14. MoFlo XDP cell sorter (Beckman Coulter, USA).

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2.2

Solutions

27

1. 70% (v/v) ethanol. 2. Dulbecco’s modified Eagle’s medium high glucose (DMEM) with L- glutamine. 3. Endothelial cell growth medium-2 Bullet Kit (EGM-2). 4. Alpha minimum essential medium (αMEM). 5. Hanks’ balanced salt solution (HBSS). 6. Phosphate-buffered saline (PBS). 7. Fetal calf serum (FCS). 8. Penicillin-streptomycin (P/S). 9. Collagenase type II-S. 10. Bovine serum albumin (BSA). 11. Red blood cell (RBC) lysis buffer (ACK lysis). 12. 0.2% gelatin solution. 13. Digestion solution: DMEM with 0.5% (v/v) BSA and 1 mg/ mL collagenase type II-S. 14. Flow cytometry staining buffer: HBSS with 0.5% (v/v) BSA and 1% (v/v) P/S.

2.3 Dyes and Antibodies

1. 0.4% trypan blue. 2. Vybrant™ DyeCycle™ violet. 3. Propidium iodide (PI). 4. Mouse anti-human CD34-APC (1:100). 5. Mouse anti-human CD146-FITC (1:100). 6. Mouse anti-human CD31-APC-Cy7 (1:200). 7. Mouse anti-human CD45-APC-Cy7 (1:200).

3

Methods

3.1 Tissue Processing and Digestion

Permission for the collection of tissue and subsequent research was approved by Johns Hopkins University Institutional IRB approval (IRB Number#00119905). Patients were informed about the tissue collection and wrote consent. Fresh tissues are collected in sterile vacuum-sealed containers and stored at 4 °C in PBS supplemented with 5% (v/v) FCS until processed (see Note 1). Whole fat from surgical abdominoplasty needs to be minced prior to digestion, while lipoaspirates (tissue retrieved following liposuction) can be processed for digestion immediately (Fig. 1). 1. Make incisions in the adipose tissue using a scalpel to remove the Scarpa’s fascial layer.

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Fig. 1 Lipoaspirate sample prior to cell isolation

2. Mechanically mince the adipose tissue using scissors, a scalpel, or other dissociation methods. 3. Combine approximately 150 mL of dissociated adipose tissue with 100 mL of PBS. 4. Mix thoroughly by manually shaking the tubes for 30 s. 5. Transfer 50 mL tissue (processed whole fat or lipoaspirate) into a 50 mL tube. 6. Centrifuge the tube at 2000 rpm for 10 min, room temperature (RT) (see Note 2). 7. Carefully transfer the middle layer into a new 50 mL tube. 8. Mix approximately 25 mL of tissue with 25 mL of PBS/2% (v/v) FCS in a 50 mL tube. 9. Centrifuge the tube at 2000 rpm for 10 min, RT. 10. Remove the supernatant and add 25 mL of the digestion solution to the remaining stromal vascular fraction pellet (SVF). 11. Transfer the tube to the shaking water bath at 37 °C, 150 rpm for 45 min. 12. After incubation, centrifuge the tubes at 2000 rpm for 10 min, RT. 13. Aspirate the supernatant (containing oily fat and adipocytes) and resuspend each pellet in pre-warmed 25 mL of PBS/2% (v/v) FCS (Fig. 2).

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Fig. 2 Lipoaspirate after collagenase digestion. The top yellow layer is fat/oil. The middle red part is digestion debris and blood. The pellet is the stromal vascular fraction prior to RBC lysis (red arrow)

14. Disrupt clumps as much as possible by pipetting up and down. Discard any large, visibly persistent clumps manually with a sterile 3 mL Pasteur pipette. 15. Filter the tissue suspension sequentially through 100 μm and 70 μm cell strainers. 16. Top up the filtered solution with PBS/2% (v/v) FCS and centrifuge at 2000 rpm for 10 min, RT. 17. Aspirate/discard the supernatant and resuspend the pellet in 10 mL of RBC lysis buffer. Incubate at RT for 5–10 min. Add 20 mL of PBS/2% (v/v) FCS to the tube and filter the resulting suspension through a 40-μm cell strainer (see Note 3). Top up the filtered solution with PBS/2% (v/v) FCS and centrifuge at 1500 rpm for 10 min, RT. 18. Resuspend the pellet in 1–2 mL of HBSS 0.5% BSA, 1% P/S depending on pellet size. Stain the cells with Vybrant™ DyeCycle™ violet stain according to manufacturer’s protocol. 19. After incubation, add an excess amount of HBSS and centrifuge at 1500 rpm for 5 min, 4°C. 20. Aspirate/discard the supernatant and resuspend the pellet (comprised of the SVF) in 1 mL (or less) of HBSS 0.5% BSA to count the cells.

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3.2 Antibody Staining

All staining procedures must be done on ice. 1. Prepare Eppendorf tubes as outlined below: Tube 1: Unstained cells. Tube 2: Full stained sample. Tube 3: Fluorescence minus one (FMO) for CD34. Tube 4: FMO for CD146. Tube 5: FMO for CD31/CD45. Add 100 μL of cell suspension (105 cells/mL) to Tubes 1, 3, 4, 5. 2. Add 400 μL of the cell suspension to Tube 2. 3. Add the antibodies as detailed below to give a final dilution of 1:100. Tube 1: no antibodies. Tube 2: CD34, CD146, CD31, CD45. Tube 3: CD31, CD45, CD146, isotype control for CD34. Tube 4: CD31, CD45, CD34, isotype control for CD146. Tube 5: CD146, CD34, isotype controls for CD31, CD45. 4. Place the stained cell suspension on ice for 30 min. 5. After staining, add twice the amount of HBSS and wash the cells by centrifuging at 1500 rpm for 5 min at 4°C. Repeat the washing once. 6. Discard the supernatant and resuspend the pellets from each tube to an appropriate volume (107 SVF cells/ mL) of HBSS. 7. Transfer the cells to their respective FACS tubes, leaving them on ice until sorting.

3.3 Flow Cytometry Pre-sort Preparation: Instrument Details and Preparation

The MoFlo XDP cell sorter has violet (405 nm), blue (488 nm), yellow-green (561 nm), and red (640 nm) lasers for fluorescence excitation and 16 fluorescence channels for detection. The blue laser is used for the detection of forward scatter (FSC) and side scatter (SSC) in the 671/28 channel. The V457/50 channel from the violet laser is used to detect nucleated cells (405 nm, DCV). The 530/40 channel is used to detect CD146 (AF488). For live/ dead cells, the 670/30 channel (PI). The lineage channel is 795/68 (APC-Cy7). This leaves the yellow-green and red lasers available for further analysis if necessary. Instrument preparation: 1. Perform the fluidics start-up and wait a short time for the stream to stabilize into droplets. 2. Calculate the drop delay manually.

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Fig. 3 Gating strategy and isolation of human adipose-derived perivascular progenitors—adventitial cells and pericytes

3. Clean the sample line and flow cell using bleach, distilled water, and 70% (v/v) EtOH. 4. Restart the stream to achieve by drop break-off point of the liquid stream. 3.4 Cell Analysis and Sorting Gating Strategy (Fig. 3)

1. Adjust voltages using an unstained cell sample and check the staining on a fully stained sample to make sure all signals are on scale. 2. Run the sample with a steady flow rate to achieve 80% cell recovery. 3. Check the single color compensation controls to ensure each single stain is the brightest in its own channel, then run the automatic compensation. 4. Run the isotype controls to check the gates. 5. Gate on the cell population excluding debris (see Note 4). 6. Gate for the single cells only using FSC and/or SSC to discriminate doublets. 7. Gate for the live nucleated cells using the DCV staining together with PI.

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8. Exclude the hematopoietic (CD45 positive) and endothelial (CD31 positive) cells. 9. Select the adventitial population by gating on CD34 positive and CD146 negative cells. 10. Select the adventitial population by gating on CD34 negative and CD146 positive cells (see Note 4). 3.5 Cell Collection and Culture

1. Collect the perivascular cells into 5-mL polypropylene tubes containing collection medium (EGM-2) on ice. 2. Following FACS, collected pericytes and adventitial cells are cultured on 0.2% gelatin pre-coated wells at 2 × 104 cells per cm2 density. To pre-coat the wells with gelatin, add the 0.2% gelatin solution to cover the bottom of well and incubate for at least 10 min at 4 °C. 3. Plate the pericytes in EGM-2 and adventitial cells in αMEM with 15% FCS and 1% P/S. 4. Change the medium every 4 days until 100% confluence is reached (approximately 2 weeks).

4

Notes 1. Adipose tissue may be stored for up to 48 h prior to processing. 2. After centrifugation, the contents of the tube will separate into 3 phases. At the top is liquid fat/oil, and the blood/fluid is at the bottom. The adipose tissue layer of interest is the central phase found between blood/fluid at the bottom and liquid fat/oil at the top. 3. Sample preparation is critical. Ensure efficient RBC lysis depending on sample size to minimize debris and RBC contamination. 4. Check gating at each stage using the isotype controls and single stains.

5

Conclusion Perivascular progenitor represents a subset of conventional mesenchymal stromal/ stem cells with high therapeutic and regeneration potential. The protocol described above employs multicolor flow cytometry approach to isolate pericytes and progenitor cells with ease from human adipose tissue.

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Acknowledgments We thank the Hao Zhang and JHU flow cytometry facility. References 1. Corselli M, Crisan M, Murray IR et al (2013) Identification of perivascular mesenchymal stromal/stem cells by flow cytometry. Cytometry 83(8):714–720. https://doi.org/10. 1002/cyto.a.22313 2. Caplan AI (2007) Adult mesenchymal stem cells for tissue engineering versus regenerative medicine. J Cell Physiol 213:341–347. https://doi.org/10.1002/jcp.21200 3. Crisan M, Yap S, Casteilla L et al (2008) A perivascular origin for mesenchymal stem cells in multiple human organs. Cell Stem Cell 3(3): 301–313. https://doi.org/10.1016/j.stem. 2008.07.003 4. Corselli M, Chen CW, Sun B et al (2012) The tunica adventitia of human arteries and veins as a source of mesenchymal stem cell. Stem Cells Dev 21:1299–1308. https://doi.org/10. 1089/scd.2011.0200 5. Crisan M, Corselli M, Chen WC et al (2012) Perivascular cells for regenerative medicine. J Cell Mol Med 16:2851–2860. https://doi. org/10.1111/j.1582-4934.2012.01617.x 6. Si Z, Wang X, Sun C et al (2019) Adiposederived stem cells: sources, potency, and implications for regenerative therapies. Biomed Pharmacother 114:108765. https://doi.org/ 10.1016/j.biopha.2019.108765 7. Gomez-Salazar M, Gonzalez-Galofre ZN, Casamitjana J et al (2020) Five decades later, are mesenchymal stem cells still relevant? Front Bioeng Biotechnol 8:148. https://doi.org/10. 3389/fbioe.2020.0014 8. Xu J, Wang Y, Hsu CY et al (2020) Lysosomal protein surface expression discriminates fat-from bone-forming human mesenchymal precursor cells. elife 9:e58990 9. Ding L, Vezzani B, Khan N et al (2020) CD10 expression identifies a subset of human perivascular progenitor cells with high proliferation and calcification potentials. Stem Cells 38: 261–275. https://doi.org/10.1002/stem. 3112 10. Hsu GC, Cherief M, Sono T et al (2021) Divergent effects of distinct perivascular cell

subsets for intra-articular cell therapy in posttraumatic osteoarthritis. J Orthop Res 39: 2388–2397. https://doi.org/10.1002/jor. 24997 11. Negri S, Wang Y, Sono T et al (2020) Human perivascular stem cells prevent bone graft resorption in osteoporotic contexts by inhibiting osteoclast formation. Stem Cells Transl Med 9:1617–1630. https://doi.org/10. 1002/sctm.20-0152 12. Xu J, Wang Y, Gomez-Salazar MA et al (2021) Bone-forming perivascular cells: cellular heterogeneity and use for tissue repair. Stem Cells 39:1427–1434. https://doi.org/10. 1002/stem.3436 13. Wang Y, Xu J, Meyers CA et al (2020) PDGFRalpha marks distinct perivascular populations with different osteogenic potential within adipose tissue. Stem Cells 38:276–290. https://doi.org/10.1002/stem.3108 14. Selich A, Daudert J, Hass R et al (2016) Massive clonal selection and transiently contributing clones during expansion of mesenchymal stem cell cultures revealed by Lentiviral RGB-barcode technology. Stem Cells Transl Med 5:591–601. https://doi.org/10.5966/ sctm.2015-0176 15. Galipeau J, Sense´be´ L (2018) Mesenchymal stromal cells: clinical challenges and therapeutic opportunities. Cell Stem Cell 22:824–833. https://doi.org/10.1016/j.stem.2018. 05.004 16. Capoccia BJ, Robson DL, Levac KD et al (2009) Revascularization of ischemic limbs after transplantation of human bone marrow cells with high aldehyde dehydrogenase activity. Blood 113:5340–5351. https://doi.org/ 10.1182/blood-2008-04-154567 17. Gomez-Salazar MA, Wang Y, Thottappillil N et al (2023) Aldehyde dehydrogenase, a marker of normal and malignant stem cells, typifies mesenchymal progenitors in perivascular niches. Stem Cells Transl Med 12(7): 474–484. https://doi.org/10.1093/stcltm/ szad024

Chapter 4 Soft Tissue Reconstruction Mario Alessandri-Bonetti, Francesco M. Egro, and Kacey G. Marra Abstract Autologous fat transplantation has revolutionized soft tissue reconstruction, but conventional methods remain unpredictable as graft resorption rates are high due to lack of vascularization. The advent of adiposederived stem cells (ASCs) has led to improvement of fat grafting outcomes, in part to their ability to undergo facile differentiation into adipose tissue, their angiogenic properties, and their ability to express and secrete multiple growth factors. This chapter discusses the isolation and characterization of human ASCs, its expansion in vitro, and relevant in vivo models for adipose tissue engineering. Key words Adipose tissue, Mesenchymal stem cells, Reconstruction

1

Introduction “We restore, repair, and make whole those parts which nature has given but fortune has taken away. Not so much that they may delight the eye but that they may buoy up the spirit and help the mind of the afflicted.” These are the words of Gaspare Tagliacozzi, who already in 1597 [1] appreciated the importance of soft tissue reconstruction, not so much for the aesthetic benefit, but for the patient’s overall physical and psychological wellbeing. Soft tissue reconstruction still remains one of the most significant challenges for plastic surgeons. A lack of soft tissue can occur due to trauma, congenital diseases (e.g., Romberg disease [2] or Poland syndrome [3]), or oncologic surgery. A variety of techniques have been employed to reconstruct such defects, including autologous composite tissue flap surgery where vascularized tissue is transferred from another region of the body, insertion of prosthetic implants, and transplantation of autologous adipose tissue. Each technique has its own disadvantages. Autologous composite tissue flap procedures allow a natural reconstruction of bone, adipose tissue, and skin, but are highly invasive with significant donor site morbidity, and potentially require multiple costly surgical

Jeffrey M. Gimble et al. (eds.), Adipose-Derived Stem Cells: Methods and Protocols, Methods in Molecular Biology, vol. 2783, https://doi.org/10.1007/978-1-0716-3762-3_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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procedures [4–7]. Implant reconstruction has associated long-term problems of migration, leakage, infection, and capsular contracture [6]. Transplantation of autologous adipose tissue minimizes these complications, and due to its minimally invasive approach, it has increasingly gained popularity throughout the U.S. [8]. From the initial report by Dr. Neuber in 1893 [9], autologous fat transplantation has undergone an evolution which has aimed to reduce resorption rates and improve clinical outcomes. Various advances in techniques and instrumentation have been achieved, and the Coleman method marked a very important milestone for the use of autologous transfer of adipose tissue in soft tissue reconstruction [10]. This worldwide-used method utilizes the principles of careful lipoaspiration, centrifugation, and subsequent grafting of the concentrated fat tissue [11]. Unfortunately, despite these advances the clinical outcome of adipose tissue transplantation remains unpredictable as there is graft resorption due primarily to lack of vascularization [12]. Adipose tissue is highly vascularized with extensive capillary networks surrounding adipocytes; therefore, it is imperative that the newly transplanted adipocytes have access to a vascular source. Furthermore, there is a constant resorption rate that can vary from 20% to 90% over time [13–17]. Immediately after harvesting of the fat graft, the suctioned aspirate can be composed of various degrees of viable fat (10–90%) [10]; and it has been hypothesized that the number of viable adipocytes at the time of transplantation correlates with the ultimate fat graft survival volume [18, 19]. The viability of adipocytes can be correlated to the extent of vascularization. Scientists have now considered the influence of precursor cells, such as adipose-derived stem cells (ASCs), normally found in fatty tissue, as a variable that can affect graft integration [10]. ASCs are the mesenchymal stem cells that are contained within the stromalvascular fraction (SVF) of enzymatically digested adipose tissue. Dr. Yoshimura popularized a technique called cell-assisted lipotransfer (CAL), which admixes SVF isolated from adipose tissue with fat graft material from the same patient with the aim of improving fat grafting outcomes [20]. ASCs have the ability to differentiate into many different cell types, such as osteoblasts [21, 22], chondrocytes [23, 24], myocytes [21, 25, 26], neuronal-like cells [27, 28], cardiovascular cells [29–32], hepatocytes [33–35], and adipocytes [21, 36–40]. ASCs are also capable of dividing and renewing for extended periods of time [41]. Of particular interest to reconstructive surgery is the ability of ASCs to undergo facile differentiation into the mesenchymal phenotype, particularly adipose tissue. ASCs have been shown to possess angiogenic characteristics and to differentiate into vascular endothelial cells [42, 43]. Furthermore, ASCs were shown to

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express and secrete multiple growth factors including IGF, HGF, TGF-b1, and VEGF, all of which are important in tissue regeneration [32, 44–47].

2

Materials 1. Dulbecco’s Modified Eagle’s Medium (DMEM/Fl2). 2. Fetal bovine serum. 3. Penicillin/Streptomycin. 4. Gentamycin. 5. Dexamethasone. 6. Collagenase solution: Type II collagenase, M8B10274 (0.1%), with bovine serum albumin (BSA; 3.5%) in 1× Hank’s solution. 7. Human regular medium: DMEM and DMEM/F12 at a 1:1 ratio, 10% fetal bovine serum, 0.1 mM penicillin, 0.06 mM streptomycin, 0.1 mM dexamethasone, and gentamycin sulfate (10 mg/L). 8. Erythrocyte lysis buffer. 9. Centrifuge tubes. 10. CyQUANT assay. 11. Fatty acid-free BSA. 12. PKH26. 13. Hematoxylin and eosin. 14. PBS. 15. Formalin.

3

Methods

3.1 Adipose Tissue Removal

1. Adipose tissue (100–200 g) is harvested from the superficial abdominal depots of females or males undergoing elective abdominal reduction surgery. 2. The patient age range is 18–80 years old and all are healthy according to clinical examination and laboratory tests. The University of Pittsburgh Institutional Review Board (IRB) approved the procedure of collecting the samples of adipose tissue. 3. Samples are not pooled, with each experiment using cells from different patients.

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ASC Isolation

1. Abdominal adipose tissue is placed in sterile 50-mL conical tubes. 2. A collagenase solution is sterile filtered then added at a 3:1 ratio of solution to adipose tissue for digestion. 3. This solution is then filtered through double-layer gauze (4 × 4 in.), to remove unprocessed debris. 4. The resulting filtered solution is centrifuged at 9200× g for 10 min at 20 °C. The supernatant is removed and the pellet is resuspended in 10 mL of erythrocyte lysis buffer (see Note 1). 5. Resuspended pellets are vortexed to lyse RBCs and debris, and then recentrifuged at 9200× g for 10 min at 20 °C. 6. The supernatant is then removed and the pellet resuspended in 10 mL of human regular media. 7. After 6 h, the ASCs are adherent to the plates and then washed extensively with PBS to remove any contaminants. 8. Cells are passaged at confluency, and characterized as previously described by our laboratory [48, 49].

ASC Culture

1. Plating media is changed three times a week, and continued until the ASCs reach approximately 80% confluence (see Note 2).

3.4 Measurement of Proliferation

1. Approximately 2.5 × 103 cells from each depot are seeded in triplicate into 24-well plates.

3.3

2. After 48 and 96 h, the plates are then frozen at -80 °C for a minimum of 24 h. 3. The plates are then thawed to room temperature and a CyQUANT® cell proliferation assay kit is used to quantify cell number. 4. Fluorescence is measured at 480-nm excitation and 520-nm emission. 5. A standard curve is created and cell number obtained. 3.5

In Vivo Model

A common in vivo model for assessment of human ASCs for soft tissue reconstruction is described. 1. A dorsal subcutaneous injection is made in 6-week-old male athymic nude mice (see Note 3). 2. Each injection is 1 mL in volume and performed through a hypodermic syringe (see Note 4). 3. For histological evaluation, animals are sacrificed, and excised gel implants are rinsed in PBS and fixed in 10% buffered formalin at 37 °C. 4. The gel explants and surrounding tissues are dehydrated in a graded series of alcohols at 37 °C, and embedded in paraffin.

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5. The transverse paraffin is sectioned through the center of gel implants at 5-μm thickness, and histologically processed using hematoxylin and eosin (H & E) stains. 6. Identification of implanted ASCs can be achieved through a variety of techniques, described in Subheading 3.6. 3.6 Identification of ACSs

Labeling of ASCs prior to animal implantation can occur using: 1. Viral transfection with green fluorescent protein (GFP). 2. Cell membrane dyes such as PKH26. 3. Nuclear dyes such as CMFDAse or DiR. 4. Gender mismatch (e.g., implantation of male cells into female animals and identification of the Y chromosome via FISH).

4

Notes 1. Erythrocyte lysis buffer is composed of 154 mM NH4Cl, 10 mM KHC03, 103 poise) such that diffusion is insignificant over less then geological time spans [42].

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1.3 Choice of Cryoprotective Agents (CPAs)

Starting with a serendipitous observation by Polge and colleagues in the 1940s the use of chemicals, denoted as cryoprotective agents (CPAs), in the freezing media has been shown to be a pre-requisite for the successful outcome of a freezing-storage process [43]. Chemicals that exhibit and/or confer some form of cryoprotection range from low molecular weight permeating solutes like dimethylsulfoxide (DMSO), glycerol, ethylene glycol (EG), etc., to high molecular weight non-permeating solutes like polyvinylpyrrolidone (PVP), hydroxyethylstarch (HES), methylcellulose (MC), etc. [44– 47]. The exact nature and the extent of cryoprotection offered by these chemicals to a specific cell type is still a matter of conjecture and is ill-understood [48–51]. But the explanations range from: (i) Enhanced colligative properties of solutions with CPAs [52–54]; (ii) To modifying the cell membrane transport properties [40, 55–61]; (iii) To preventing denaturing of membranes by elevated concentrations of extracellular salts at low temperatures [62–65]; (iv) To forming a protective coating on sensitive plasma membranes [66–69]; (v) To the prevention of seeding of the supercooled intracellular water and thus, the formation of damaging intracellular ice [70–72]; and. (vi) To the ability of the CPAs to alter the physical properties of solutions during freezing rate than in any direct effects on cellular membranes [73–80]. Thus, the choice and the composition of an optimal cryopreservation media is an area of intense research in the field of cryobiology and is not, as yet, completely understood. Our experiments with SVF and P1 adult stem cells were conducted with the media listed later in this chapter [36–38].

2

Materials Where appropriate, materials should be sterile and all manipulations performed using a laminar flow bench (fume hood). Adequate safety equipment (gloves, face shield) will be used during all liquid nitrogen handling procedures. 1. Sterile plastic dishes/pipets/micropipettes. 2. -80 °C freezer. 3. 37 °C CO2 incubator and 43 °C CO2 incubator. 4. Normal and cooling centrifuges.

Adult Stem Cells Freezing Processes and Cryopreservation Protocols

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5. Commercially available and programmable controlled rate freezer (Planer Series Kryo 560–16, TS Scientific, Perkasie, PA) able to cool at rates of 1 °C/min to 40 °C/min to -80 ° C (see Note 2). 6. Type-T hypodermic thermocouples and precision temperature data logger. 7. Directional solidification stage, capable of controlled cooling rates ranging from 1 °C/min to 40 °C/min to -80 °C. 8. Liquid nitrogen storage dewars (long-term storage containers). 9. 37 °C warm water bath. 10. Digital image analyzer—typhoon laser imaging scanner. 11. Glass microslides with central grooves. 12. Warm copper block maintained at 50 °C. 13. Cryovials, such as 2-mL plastic cryotubes. 14. Diluent: Dulbecco’s Modified Eagle Medium (DMEM), DMEM-F12, fetal calf serum (FCS) and human serum. 15. Supplies for sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) including distilled water, Tris (pH 8.8 and pH 6.8), acrylamide-bisacrylamide, ammonium persulfate (APS), TEMED, and EtOH. Gel casting apparatus including gel casting plates, comb, power pack, and plate holder/ container. 16. Lysis buffer—RIPA buffer with protease cocktail inhibitors. 17. BCA assay kit for protein concentration. 18. Loading buffer (6×) containing Tris, glycerol, bromophenol blue, and SDS. 19. Western blotting antibodies (HSP-70, HSP-90, HSP-27, and β-actin) from Santa Cruz Biotechnology, Dallas, TX. 20. Western blot chemiluminescence substrate. 21. Running buffer or 1× Tris-glycine buffer. 22. Transfer buffer or 1× Tris-glycine buffer and methanol. 23. Blocking buffer or 5% non-fat milk with 1× Tris-buffered saline with Tween® 20 (TBST). 24. Washing buffer or 1× Tris-buffered saline with Tween® 20 (TBST). 25. Cell culture medium containing 10% FBS at equivalent of 1 mL of tissue per mL of media (see Note 3). 26. Adipogenic induction cocktail containing DMEM/F-12 Ham’s with 3% FBS, 33 μM biotin, 17 μM pantothenate, 1 μM bovine insulin, 1 μM dexamethasone, 0.5 mM isobutylmethylxanthine (IBMX), 5 μM rosiglitazone, and 100 U penicillin/100 μg streptomycin/0.25 μg fungizone.

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27. Osteogenic induction cocktail containing DMEM/F-12 Ham’s, 10% FBS, 10 mM β-glycerophosphate, 50 μg/mL sodium ascorbate 2-phosphate, and 100 U penicillin/100 μg streptomycin/0.25 μg fungizone. 28. Cryoprotectants (CPAs): Dimethylsulfoxide (DMSO) (see Note 6), methylcellulose (MC), polyvinylpyrrolidone (PVP). All cryoprotectant solutions prepared using DMEM supplemented with either fetal calf serum or human serum. PVP and MC were individually autoclaved at 121 °C for 30 min before being added to DMEM (see Note 4). The DMEM-PVP solutions and DMEM-MC solutions were prepared by dissolving weighted PVP and MC in DMEM at room temperature and the solutions were then stored overnight at 4 °C to obtain a homogeneous preparation. Concentrations above 1% MC and 40% PVP were found to be highly viscous and hard to handle and hence, were not used [36–38]. 29. Fluorescence-activated cell sorting (FACS) machine and associated chemicals (Etoposide, hydrogen peroxide, annexin V-FITC/PI kit, propidium iodide): Cell Quest software to acquire the FACS data. Apoptosis control was cells incubated in fresh medium enriched with 40 mM Etoposide for 24 h. Necrotic control was cells incubated in fresh medium with 5 mM hydrogen peroxide (H2O2) for 24 h.

3

Methods

3.1 Collection and Isolation of ASCs

The process of adult stem cell isolation (and culture/expansion/ differentiation) from human liposuction aspirate (adipose tissue) is described elsewhere [81–90] and is only briefly described below: 1. Obtain liposuction aspirate from plastic surgeon. 2. Wash tissue and collagenase digest at 37 °C with continuous shaking for 1 h. 3. Centrifuge digest to obtain “stromal vascular fraction (SVF)” pellet (see Note 12). 4. Suspend SVF in culture media. 5. Seed flasks at a density of 0.156 mL tissue digest per sq. cm surface area as “Passage 0 (P0)”. Incubate at 37 °C, 5% CO2. 6. Wash cultures to remove non-adherent cells after 24 h of incubation. 7. Feed cultures every 2–3 days. 8. Harvest P0 by trypsin/EDTA digestion when it reaches 80–90% confluency (mean, 5.5 days).

Adult Stem Cells Freezing Processes and Cryopreservation Protocols

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9. Seed “Passage 1 (P1)” and subsequent passages of undifferentiated ASCs at a density of 5 × 103 cells per sq. cm and maintain according to steps 6 and 7. 10. Harvest P1 by trypsin/EDTA digestion when it reaches 80–90% confluency (mean, 5.5 days) (see Note 1). 3.2 Freezing Media for SVF of Adipose Tissue (Those with *’s Are Sub-optimal)

The measured post-freeze-thaw viability is shown for each media. The detailed freeze-thaw procedures are described in Subheading 3.3. Additional (and corresponding) data on post-thaw apoptotic and necrotic cells is presented elsewhere [37].

1. DMEM) with 80% FCS and 10% DMSO

65.3 ± 3.3%

2. DMEM with 80% human serum (HS) and 10% DMSO 61.9 ± 2.4%

3.3 Freezing Media for P1 ASCs (Those with *’s Are Suboptimal)

3. *DMEM with 0% DMSO

11.4 ± 7.4%

4. DMEM with 2% DMSO

60.1 ± 4.4%

5. DMEM with 4% DMSO

62.8 ± 4.5%

6. DMEM with 6% DMSO

65.8 ± 2.4%

7. DMEM with 8% DMSO

72.9 ± 3.3%

8. DMEM with 10% DMSO

69.8 ± 2.5%

9. *DMEM with 1% MC

37.3 ± 4.1%

10. *DMEM with 1% MC and 10% FCS

22.2 ± 8.3%

11. *DMEM with 1% MC and 10% HS

20.0 ± 6.8%

12. DMEM with 1% MC and 10% DMSO

66.0 ± 2.1%

13. DMEM with 10% PVP

54.6 ± 1.7%

14. DMEM with 10% PVP and 10% FCS

52.3 ± 5.7%

15. DMEM with 10% PVP and 40% FCS

49.7 ± 6.7%

16. DMEM with 10% PVP and 80% FCS

58.6 ± 3.2%

17. DMEM with 10% PVP and 10% HS

53.9 ± 5.5%

The measured post-freeze-thaw viability is shown for each media. Additional (and corresponding) data on post-thaw apoptotic and necrotic cells is presented elsewhere [36, 38].

1.

DMEM with 80% FCS and 10% DMSO

84.1 ± 7.7%

2.

DMEM with 80% HS and 10% DMSO

82.5 ± 8.3%

3.

*DMEM with 0% DMSO

25.3 ± 5.7% (continued)

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3.4 Process of Adding CPAs and Cooling to Liquid Nitrogen Temperatures and Thawing to Room Temperature

4.

DMEM with 2% DMSO

83.8 ± 5.3%

5.

DMEM with 4% DMSO

80.6 ± 3.1%

6.

DMEM with 6% DMSO

86.0 ± 4.6%

7.

DMEM with 8% DMSO

85.6 ± 2.8%

8.

DMEM with 10% DMSO

87.9 ± 2.9%

9.

*DMEM with 1% MC

46.8 ± 7.6%

10.

*DMEM with 1% MC and 10% FCS

41.5 ± 7.6%

11.

*DMEM with 1% MC and 10% HS

39.8 ± 7.6%

12.

DMEM with 1% MC and 10% DMSO

79.8 ± 2.8%

13.

*DMEM with 1% PVP

5.7 ± 0.5%

14.

DMEM with 5% PVP

49.3 ± 8.0%

15.

DMEM with 10% PVP

69.7 ± 8.2%

16.

DMEM with 20% PVP

55.5 ± 11.0%

17.

*DMEM with 40% PVP

18.

DMEM with 5% PVP and 10% HS

53.2 ± 8.7%

19.

DMEM with 10% PVP and 10% HS

72.1 ± 8.1%

20.

DMEM with 20% PVP and 10% HS

61.8 ± 12.5%

21.

DMEM with 5% PVP and 10% FCS

55.4 ± 11.0%

22.

DMEM with 10% PVP and 10% FCS

69.2 ± 6.7%

23.

DMEM with 20% PVP and 10% FCS

64.3 ± 6.4%

24.

DMEM with 10% PVP and 40% FCS

70.5 ± 3.5%

25.

DMEM with 10% PVP and 80% FCS

72.5 ± 2.5%

4.6 ± 1.3%

1. Suspend the SVF or P1 ASCs at a concentration of 1.0 × 106 cells/mL in the various cryopreservation medium, described above (see Note 5). 2. Place the cells in an ethanol-jacketed closed container overnight at -80 °C. 3. The temperature/time history experienced by the cells in the ethanol-jacketed container can be measured using a type-T hypodermic needle thermocouples. Thermocouple voltages can be read by a precision temperature data logger and transferred to a personal computer for further reduction and data analysis. The representative cooling rates experienced by the cells in the ethanol-jacketed container placed in a -80 °C freezer is shown in Fig. 1 (see Note 14).

Adult Stem Cells Freezing Processes and Cryopreservation Protocols

B

30

30

20

CR1= 1.22 bC/min

20

CR1=1.31 bC/min

10

CR2=0.41 bC/min

10

CR2=0.42 bC/min

0

CR3=1.23 bC/min

0

-10

CR4=0.37 bC/min

CR1

CR 2

Temperature (ºC)

Temperature (ºC)

A

-20 -30

CR3

-40 -50 -60

CR4

CR1

EHR

CR3=1.76 bC/min CR2

-10

CR4=0.45 bC/min

-20 CR3

-30 -40 -50 -60

-70

-70

-80

-80

CR4

-90

-90 0.0

61

20.0

40.0

60.0

80.0

100.0

Time (min)

120.0

140.0

160.0

0.0

20.0

40.0

60.0

80.0

100.0

120.0

140.0

Time (min)

Fig. 1 Representative cooling rates experienced by the P1 ASCs in the ethanol-jacketed closed container placed in the -80 °C freezer with 80% FCS + 10% DMSO in DMEM (Fig. 1a) and 10%PVP in DMEM (Fig. 1b). The measurements were measured using type-T thermocouple coupled with a precision temperature data logger. The plot suggests that the cells are subjected to different cooling rates at different time points in different freezing media within the ethanol-jacketed container. For media with 80% FCS + 10% DMSO in DMEM, the ice nucleation was observed around -8 (±1.1) °C and subsequently, a cooling rate of ~0.4 °C/min was imposed to a temperature of ~ -18 °C. The cooling rates experienced by the cells then further drops to ~1.2 °C/min until ~ -50 °C and then to ~0.4 °C/min, before reaching ~ -80 °C (Fig. 1a). Alternatively, for media with 10% PVP in DMEM, there was significant super cooling before any ice nucleation was observed around -17 (±2.3) °C. Due to the enthalpic heat release (EHR) the sample temperature was abruptly raised to -5 °C and a subsequent cooling rate of ~0.4 °C/min was imposed to a temperature of -10 °C. The cooling rates experienced by the cells then further drops to 1.8 °C/min until ~ -50 °C and then to ~0.4 °C/min, before reaching ~ -80 °C (Fig. 1b)

4. Note that the exact magnitude of the cooling rates experienced by the cells in the ethanol-jacketed container is a function of the cryopreservation media. 5. Store the cells in liquid nitrogen vapor and/or immersed in liquid nitrogen (see Notes 7, 8, 9, and 13). 6. Rapidly thaw individual cryovials of cells in a 37 °C water bath (1–2 min of agitation (see Note 10). 7. Re-suspend the cells in stromal culture media, and seed into the separate wells of a 6-well plate for a 24-h incubation period at 37 °C (see Note 11). 3.5 Evaluation of Frozen-Thawed Cells

The post-thaw viability and functionality of the cells was analyzed using flow cytometry [91] and histo-chemical staining for adipogenic and osteogenic differentiability [35, 83, 84, 86–88].

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3.5.1 Cell Viability and Apoptosis/Necrosis Assessment

1. A well-established annexin V apoptosis assay was analyzed by quantitative flow cytometry. 2. Chemically induced apoptotic control was cells incubated in fresh medium enriched with 40 mM Etoposide for 24 h. Apoptosis is characterized by phosphatidylserine (PS) translocation from the inner leaflet to the outer leaflet of the lipid bilayer, while the cell membrane remains intact. Annexin V-positive cells correspond to cells that have experienced PS translocation. 3. Necrotic control was cells incubated in fresh medium with 5 mM hydrogen peroxide (H2O2) for 24 h. PI staining of the cells indicates that the integrity of the cell membrane has been compromised and is used to distinguish living and early apoptotic cells from necrotic cells. 4. The no-treatment control was cells in fresh medium, free from inducing agents. 5. For each treatment, detached and attached cells were pooled, harvested by trypsinization (0.25% trypsin), washed with 10 mL of culture medium, and re-suspended in 100 μL of 1× annexin-binding buffer (included in annexin V-FITC/PI kit). 6. Approximately 100 μL of the cell suspension was mixed with 8 μL of annexin-V-FITC and 8 μL of 100 μg/mL propidium iodide (PI) and incubated in the dark at room temperature for 15 min. 7. Liquid volume was removed by centrifugation and aspiration, and the cells were re-suspended by gentle vortexing in 300 μL of 1× annexin-binding buffer and subsequently analyzed on the flow cytometer. 8. Apoptotic analyses for ASCs were performed on a FACS Caliber flow cytometer (BD Biosciences, San Jose, CA) utilizing 488-nm laser excitation and fluorescence emission at 530 nm (FL1) and > 575 nm (FL3). 9. Forward and side scatter measurements were made using linear amplification, and all fluorescence measurements were made with logarithmic amplification. A total of 20,000 cells per sample were acquired using Cell Quest software (BD Biosciences, San Jose, CA). 10. Quadrant analysis was performed on the fluorescence dotplot to quantify the percentage of live, necrotic, and apoptotic cell populations [36–38, 91]. The fluorescent dotplots show three cell populations: live (annexin V-FITC-negative and PI-negative; annexin V- and PI-), necrotic (annexin VFITC-positive and PI-positive; annexin V+ and PI+), and apoptotic (annexin V-FITC-positive and PI-negative; annexin V+ and PI-). The quadrants positions were placed according to the no treatment control and 5 mM H2O2 necrotic control (Fig. 2).

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Fig. 2 Characteristic flow cytometer fluorescence dotplots showing fluorescence-activated cell sorting (FACS) analysis of SVF cells frozen/thawed in the presence various concentrations (0%, 2%, 4%, 6%, 8%, or 10%) of DMSO in DMEM. Corresponding plots for other media and P1 ASCs are available in the literature [36, 38]. Figure 2a–f represent 0%, 2%, 4%, 6%, 8%, or 10% DMSO in DMEM, respectively. The fluorescent dotplots show three cell populations: live (annexin V-FITC-negative and PI-negative; annexin V- and PI-) necrotic (annexin V-FITC-positive and PI-positive; annexin V+ and PI+), and apoptotic (annexin V-FITC-positive and PI-negative; annexin V+ and PI-). The quadrants positions were placed according to the no treatment control, 40 μm Etoposide apoptotic control and 5 mM H2O2 necrotic control. (Reprinted with permission from Thirumala et al. [37]) 3.5.2

Adipogenesis

1. Confluent cultures of ASCs were induced to adipogenesis by replacing the medium with an adipocyte induction cocktail (see Subheading 2, item 5). 2. After 72 h, the adipocyte induction medium was replaced with adipocyte maintenance media, which contains the same components as the induction medium except IBMX and rosiglitazone. 3. Induced cells were maintained in culture for 9 days, with adipocyte maintenance medium replacement every 3 days. Upon the ninth day, the cultures were washed twice with pre-warmed PBS and fixed in formalin at 4 °C. Adipocyte quantification was determined by staining neutral lipids with Oil Red O (Fig. 3).

3.5.3

Osteogenesis

1. Confluent cultures of ASCs were induced to osteogenesis by replacing the medium with an osteogenic induction cocktail (see Subheading 2, item 6).

64

Mohan Kumar Dey and Ram V. Devireddy 80%HS+10%DMSO +10%DMEM

80%FCS+10%DMSO +10%DMEM

0%FCS+10%DMSO +90%DMEM

0%FCS+2%DMSO +98%DMEM

Control

Adipogenesis

Osteogenesis

Fig. 3 Representative phase contrast photomicrographs of P1 ASCs cultured under untreated (first row; Toludine blue staining), adipogenic (second row; oil red O staining), or osteogenic (third row; alizarin red staining) conditions. Corresponding plots for other media and SVF cells are available in the literature [37, 38]. Adipogenic cultures were stained with oil red O 14 days after induction while osteogenic cultures were stained with alizarin red after 21 days of culture. Images in column 1 represent cells that were cryopreserved in media containing 80% FCS with 10% DMSO in DMEM. Images in column 2 represent cells that were cryopreserved in media containing 80% HS with 10% DMSO in DMEM. Images in column 3 represent cells that were cryopreserved in media containing 0% FCS with 10% DMSO in DMEM. And finally, images in column 4 represent cells that were cryopreserved in media containing 0% FCS with 2% DMSO in DMEM. (Reprinted from Thirumala et al. [36])

2. The induced cells were fed fresh osteogenic induction media every 3 days for 3 weeks. The cultures were then washed with 0.9% sodium chloride solution and fixed in 70% ethanol. 3. Osteoblast quantification was determined by Alizarin red staining for calcium phosphate (Fig. 3). 3.6 Some Observations on the Effect of Various Cryoprotectants on the Post-Freeze/Thaw Response of SVF of Adipose Tissue

1. The data suggests that the choice of the serum (human serum, HS or fetal calf serum, FCS) does not significantly alter the SVF cell survival when frozen/thawed in 10% DMSO and DMEM. 2. The highest % of post-thaw survival was achieved with a concentration of 8% DMSO (~73%), the post-thaw survival values obtained with DMSO concentrations of 2%, 4%, 6%, and 10% are comparable and are ~60%, ~63%, ~66%, and ~70%, respectively. The only significant differences in the data were found when the cells were frozen with DMSO and without DMSO, i.e., even the presence of 2% DMSO was significant and this extremely low concentration was able to cryoprotect SVF cells.

Adult Stem Cells Freezing Processes and Cryopreservation Protocols

65

3. Replacing DMSO with MC as the CPA lead to a significant reduction (~50%) in the post-thaw cell viability of SVF cells. Intriguingly, the addition of either HS or FCS with MC is extremely deleterious to post-thaw cell viability. 4. DMSO is more effective in preserving cell viability than MC, i.e., the maximum % of cell survival obtained with MC (~37%) is still only half of that obtained with media containing various concentrations of DMSO (~ 60–70%). 5. The presence or the absence of FCS did not significantly alter the post-thaw viability results for SVF cells frozen in the presence of 10% PVP. 6. The % of post-thaw cell viability (~55%) obtained with 10% PVP and DMEM is comparable to that obtained (~64%) with control media (10% DMEM with 10% DMSO and either 80% FCS or 80% HS) and also comparable with the values obtained with DMSO and DMEM (~65%). 3.7 Some Observations on the Effect of Various Cryoprotectants on the Post-Freeze/Thaw Response of P1 ASCs

1. The viability of ASCs cryopreserved in the control media containing 80% serum with 10% DMSO in DMEM was ~83%. 2. The highest % of post-thaw cell survival was found for cells frozen in DMEM with 2–10% DMSO (~80% to ~88%) and the lower values were found for cells frozen in DMEM with either 40% PVP or 1% PVP (~5%), DMEM with 0% DMSO (~25%) and in DMEM with 1% MC and either 10% HS or 10% FCS (~40%). 3. Freezing P1 ASCs in the absence of DMSO was detrimental to cell survival. 4. The addition of 1% MC significantly lowers the cell viability, i.e., the post-thaw cell viability with 1% MC in DMEM (~47%) is significantly lower than that obtained with DMEM containing 1% MC and 10% DMSO (~80%). 5. The use of 1% and 40% PVP in DMEM caused a dramatic loss in cell viability (~5%) whereas the use of 10% PVP produced a maximum viability of ~65%. Although the viability of ASCs decreased when the % of PVP was increased from 10 to 20, the data analysis revealed that this increase is statistically not significant (>95% confidence level). 6. The ASCs frozen/thawed in the presence of PVP (and in the absence of both DMSO and serum) displayed similar morphology and growth/differentiation characteristics when compared to the ASCs cryopreserved in 80% serum with 10% DMSO in DMEM (Fig. 3). An important (and unique) feature of our ASC freezingstorage protocols is the development of a cryopreservation media without serum (and without DMSO) that simplifies their use and application in in vivo tissue engineering applications.

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Fig. 4 Photographs of the custom-built directional solidification stage (DSS) on the left-hand side and the commercially available controlled rate freezer (CRF) on the right-hand side (source: “Bioengineering Laboratory, Department of Mechanical Engineering, Louisiana State University, Baton Rouge, LA”). As described in the text, the choice of the freezing device has a significant impact on the post-thaw viability of ASCs

3.8 Effect of Freezing Device on the PostFreeze/Thaw Response of ASCs

1. The effect of directional cooling on the immediate post-thaw membrane integrity of adipose tissue-derived adult stem cells (ASCs) was investigated using a directional solidification stage (DSS) and a commercially available controlled rate freezer (CRF)—Fig. 4. 2. P0, P1, and P2 ASCs were cooled at either 1, 5, 20 or 40 °C/ min to an end temperature of -80 °C in the presence and absence of a cryoprotective agent (DMSO). For experiments in the presence of DMSO, the cell suspensions were gently mixed with 10% DMSO as a cryoprotective solution (CPA), i.e., a one-step addition of CPA. The samples were equilibrated in the cryoprotective solution for ~10 min before the DSS and CRF experiments were performed. 3. After freezing to -80 °C, the samples were thawed at 200 °C/ min and the ability of the frozen/thawed ASCs to exclude fluorescent dyes was assessed.

3.9 Directional Solidification Stage (DSS) Experiments

1. The directional solidification stage (DSS) built by Rubinsky & Ikeda [92] consisted of hot and cold temperature copper bases (set at Th and Tc, respectively) separated by a distance (Dgap). 2. Freezing was achieved by placing the sample on a glass slide and moving the glass slide from the hot base to the cold base at a known velocity (V). The cooling rate (B) experienced by the h -Tc sample is then described as: B = T D •V. gap 3. For the DSS freezing experiments, ~100 μL of ASC suspension was placed in a grooved microslide and traversed between the precisely controlled copper blocks at a constant velocity. Note that the first copper block was held at +4 °C and the second at -80 °C. The samples were then cooled at a nominal cooling

Adult Stem Cells Freezing Processes and Cryopreservation Protocols

67

rates of 1, 5, 20 or 40 °C/min to an end temperature of -80 ° C. Due to interactions with the ambient atmosphere the actual cooling rates experienced by the cells between 0 and -40 °C were ~ 1.1, ~5.2, ~10.6, ~21.7, and ~ 43.6 °C/min, respectively [34]. 4. All the glass microslides had a circular milled well, perpendicular to the direction of microslide travel, within which the cell suspension was placed. The well was 3.2 mm wide (in the direction of travel) and of 1 mm depth (into the microslide surface). 5. The thawing rate of ~200 °C/min was achieved by quickly bringing the bottom of the glass slide into contact with a warm copper block maintained at 50 °C. 3.10 Controlled Rate Freezer (CRF) Experiments

1. A programmable controlled rate freezing (CRF) machine (Planer Series Kryo 560–16, TS Scientific, Perkasie, PA). 2. The samples were loaded into a 6-well cell culture cluster. 3. The cluster was then loaded into the cryo-machine (which was precooled to +4 °C) and kept for 1 min for equilibration. The samples were then cooled at either 1, 5, 20 or 40 °C/min to an end temperature of -80 °C. 4. The thawing rate 200 °C/min was obtained by removing the 6-well cluster from the CRF machine and quickly bringing the bottom of the cluster into contact with a warm water bath maintained at 37 °C [41].

3.11 Some Observations on the Effect of Freezing Device on the PostFreeze/Thaw Response of ASCs

1. The viability of cell suspensions was measured using calcein AM and propidium iodide (PI) [93]. 2. A 10 μL sample of cell suspension was removed from the microslide and incubated with 2 μmol/L calcein AM and 3 μmol/L PI for 15–30 min at 37 °C. 3. After incubation, cells were scored as either live or dead under a fluorescent microscope. 4. Viability measurements of cells taken directly from the stock suspension without freezing were determined before and after every freeze-thaw experiment as a control; all experimental viability values were normalized to the average control viability. 5. A comparison of the P1 ASCs post-freeze viability in the presence and absence of DMSO obtained using the DSS and the CRF is shown in Fig. 5. Corresponding data for P0 and P2 cells are available elsewhere [34]. 6. The post-freeze viability of ASCs is significantly higher (confidence level > 95%) when they are frozen in the presence of DMSO, than in its absence.

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Fig. 5 Comparison of post-thaw viability of P1 ASCs frozen in the absence (a) and presence (b) of 10% (v/v) DMSO is shown. In each figure, the post-thaw viability obtained using the DSS (unfilled columns) and CRF (filled columns) is also shown. The error bars represent standard deviation in the data (n = 18). In both the figures, the immediate post-thaw ASC membrane integrity is shown on the Y-axis while the cooling rate imposed on the sample (°C/min) is shown on the X-axis. (Redrawn from Fuller and Devireddy [34]

7. A cooling rate of 1 °C/min results in the highest post-thaw cell survival for ASCs frozen using either devices. 8. The post-freeze viability of ASCs is uniformly higher (confidence level > 95%) when they are frozen using the CRF as opposed to using the DSS. This significant reduction in cell viability for the cells frozen using the DSS might be related to previously postulated mechanical/damaging interactions between the ice crystals and the cell membranes [94–97]. 3.12 Effect of PreHeating (Inducing Heat Shock Proteins) on the Post-Freeze/Thaw Response of ASCs

1. Heat shock proteins (HSPs) are known to be upregulated when the cells encounter unfavorable conditions such as elevated temperatures, hypoxia, heavy metals, ethanol, oxidative phosphorylation inhibitors, and other chemicals [98]. They function as molecular chaperones in proper folding of proteins, reduce protein aggregation and also alleviate oxidative stress that occurs in various pathological conditions [99, 100]. 2. Literature suggests that HSP70, HSP90, and HSP27 inhibit the apoptosis pathway in stressed cells and may provide cytoprotection [101–106]. For example, studies conducted on hela cells indicate that heat shock treatment significantly improves the survival rate in comparison to control hela cells [107] whereas in human fibroblasts the induction of HSPs did not provide cytoprotection [108].

Adult Stem Cells Freezing Processes and Cryopreservation Protocols

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3. Thus, the hypothesis of these experiments is that the induction of HSPs prior to the freeze-thaw insult can improve the postthaw viability of ASCs by attenuating the effects of freezing associated stresses. 3.12.1 Collection and Isolation of ASCs 3.12.2 Heat Shock Treatment

The culturing of ASCs, preparation of freezing media, evaluation of post-thaw viability was done as described earlier. 1. The P1 ASCs after reaching 80% confluence were placed in a cell culture incubator that was adjusted to 43 (± 0.2) °C for 1 hour to induce the HSPs expression. 2. After 1 h of heat shock, the cells were transferred back to cell culture incubator at 37 °C and incubated for 18 h prior to freezing.

3.12.3 Freezing/Thawing Protocol

1. The heat-shocked ASCs were suspended at a density of 1.0 × 106 cells/mL with DMSO at concentrations of 10% or 1% and PVP at 10% and without any CPA in DMEM media containing 10% FBS in 1.5 mL cryovials. The ASCs without heat shock were prepared in the same way and used as controls. 2. Using a CRF the cryovials were cooled down to -35 °C from room temperature at a cooling rate of 1 °C/min followed by plunging in liquid nitrogen. The cells were stored in liquid nitrogen for 1 day. 3. The next day the cryovials were taken out from liquid nitrogen and thawed in 37 °C water bath for 1–2 min by gentle shaking. 4. The thawed ASCs were resuspended in stromal media and cultured for 24 h before assessing the post-thaw viability using Annexin V/Propidium iodide flow cytometer-based method as described earlier.

3.12.4 Casting Polyacrylamide Gels for SDS-Page

1. Resolving gel master mix: 400 mL distilled water, 250 mL of 1.5 mL Tris pH 8.8 with 10 mL of 10% SDS. 2. Stacking gel master mix: 340 mL of distilled water, 62.5 mL 1.0 M Tris pH 6.8 with 5 mL of 10% SDS. 3. Pouring resolving gel: To make 6 mL of resolving gel, pour 4 mL of resolving gel master mix into a 50 mL tube followed by 2 mL of 30% acrylamide and 60 μL of 10% APS. Vortex the mixture quite vigorously for 1–2 min and add 2.5 μL of TEMED. Vortex or shake the mixture vigorously again for another 1 min. 4. Immediately load gel mixture into the casing with a pipette— fill to the line on the casing. Add distilled water on top of the gel to eliminate or mitigate the presence of any air bubbles. Allow 30 min to polymerize at room temperature (25–27 °C).

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Remove water with a pipette or a blotting paper after the polymerization process. Use the gel immediately or store for later use at 4 °C in 1% Tris-glycine buffer. 5. Stacking gel: To make ~3 mL of stacking gel, add 2.5 mL of stacking gel master mix, 0.5 mL of 30% acrylamide, 30 μL of 10% APS. Vortex the mixture quite vigorously for 1–2 min and add 2.5 μL of TEMED. Vortex or shake the mixture vigorously again for another 1 min. 6. Immediately or before the gel solidifies (~2 min) insert the gel comb carefully to avoid incorporating air bubbles (see Note 15). 7. Allow the gel to polymerize or solidify (~ 15–30 min). Carefully remove the gel comb from the top of the stacking gel (see Note 16). 3.12.5

Western Blotting

1. The ASCs recovered for 18 h post-heat shock were lysed with ice cold RIPA buffer with protease cocktail inhibitor. 2. The collected cell lysates were centrifuged at 8000 × g for 10 min at 4 °C using a cooling centrifuge and the supernatants were collected. 3. The total protein concentration was estimated using BCA assay. 4. Denature the proteins by adding the loading buffer. Heat the cell lysate and loading buffer for 3–5 min at 95 °C (see Note 17). 5. Let the sample cool to room temperature (25–27 °C) or ~ 5–10 min. Apply the appropriate power pack (160 V) until the bromophenol blue dye permeates to the bottom of the gel or approximately 40–45 min (see Note 18). 6. The protein samples were separated using 10% polyacrylamide gel using SDS PAGE and then transferred onto nitrocellulose membrane via the application of electrical field (70 V for 2 h) (see Note 19). 7. The membrane was then blocked using 5% non-fat dry milk prepared in 1× TBST for 1 h. Later the membrane was incubated overnight with antibodies for HSP90, HSP70, and β-actin at dilution of 1:200 and HSP27 with 1:5000 dilution at 4 °C. 8. The membrane was washed several times with 1× TBST before incubation of HRP-conjugated secondary antibody at dilution of 1:2000 for 1 h. 9. One hour after incubation of secondary antibody the membrane was washed again several times and treated with chemiluminescent substrate.

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10. The detection of chemiluminescence signal was done using typhoon laser imaging scanner (GE Life Sciences, Pittsburgh, PA). 3.12.6

qPCR

1. The total RNA of heat-shocked ASCs was isolated using Purelink RNA mini kit (Life Technologies, Grand Island, NY). 2. The isolated RNA was quantified using nanodrop spectrophotometer and reverse transcribed using high capacity cDNA reverse transcription kit (Applied Biosystems, Grand Island, NY). 3. The synthesized cDNA was later used to carry out qPCR using SYBR select master mix (Applied Biosystems, Grand Island, NY). The qPCR data was analyzed using Δ ΔCt method.

3.12.7 Osteogenesis: Alizarin Red Staining and Quantification

To determine the effect of heat shock on osteogenesis, ASCs were given heat shock on 3rd, 9th, and 15th day of osteogenic differentiation. Osteogenic induction and heat shock were performed as described in Subheadings 3.4 and 3.9, item 1. 1. 40 mM alizarin red S solution was prepared in distilled water and the pH was adjusted to 4.1 using ammonium hydroxide. 2. On the 21st day of osteogenic induction the cells were and fixed with 4% paraformaldehyde for 30 min and stained with the 40 mM alizarin red S solution for 30 min. 3. The stained wells were washed with distilled water to remove excess unbound stain and examined with inverted microscope. 4. The alizarin red S stained wells were treated with 10% cetylpyiridium chloride to elute the calcium bound stain and the absorbance of that solution was measured at 530 nm using spectrophotometer. 5. A standard plot was also generated with various concentration of alizarin red S stain which was used to quantify the amount of eluted dye.

3.13 Some Observations on the Effect of Inducing Heat Shock Proteins Prior to Freeze-Thaw Process

1. Western blotting expression of HSPs in ASCs is shown in Fig. 5. Heat treatment for 1 h at 41 °C did not significantly enhance the expression of HSPs studied, i.e., HSP27, HSP70, and HSP 90. Increasing the pre-heat treatment temperature to 43 °C for 1 hr. showed an upregulation of HSP70 with no significant change in either HSP27 or HSP90. 2. The post-thaw viability of heat-shocked (at 43 °C for 1 h) ASCs in comparison to controls is shown in Table 1. The lack of significant differences between the heat treated cells and control, suggests that the upregulation of HSP70 alone is not sufficient to provide cryoprotection in ASCs; with the caveat that the “time point” of freezing after the heat shock treatment might not be optimal.

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Table 1 Post-freeze-thaw cell viability of ASCs with (43 °C for 1 h) and without heat treatment in various cryopreservation media % Post-thaw cell viability normalized to pre-freeze cells Cryoprotectant

No heat shocked

Heat shocked

No cryoprotectant

34.7 (±7.25)

26.2 (±2.56)

1% DMSO

70.9 (±9.68)

72.1 (±4.63)

10% DMSO

91.6 (±1.67)

93.6 (±1.33)

74.92 (±3.86)

79.33 (±3.65)

10% PVP A)

HSP90

HSP70

HSP27

41˚ C 37 ˚C

β-Actin

B)

HSP90

HSP70

HSP27

C)

HSP90

43˚ C 37 ˚C

β-Actin

HSP70

HSP27

β-Actin

Fig. 6 Western blotting of HSP90, HSP70, and HSP27 of heat-shocked ASCs at different temperatures: 41 °C for 1 h (a) and 43 °C for 1 h (b). The elevated expression level of HSP70 is conserved in heat-shocked ASCs after cryopreservation (c)

3. To determine the time point where maximum expression of HSPs occur after heat shock, qPCR was performed on RNA isolated from ASCs at 3, 9, 18, 48, and 72 h after heat shock treatment and is shown in Fig. 6. 4. The data indicates that only HSP70 and HSP27 mRNA expression levels decreased with time whereas HSP90 remained virtually unchanged. Specifically, 3 h after the heat shock HSP70 and HSP27 mRNA levels reached their peak levels (Fig. 7). At 18 h after the heat shock treatment the mRNA levels of both HSP70 and HSP27 were at the basal level. However, western blotting data indicates that HSP70 protein levels are still elevated 18 h post-heat shock whereas little change in HSP27 is observed.

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Fig. 7 The expression levels of HSP90, HSP70, and HSP27 at various time points after heat shock treatement at 43 °C for 1 h

5. The data in Fig. 7 suggests that freezing ASCs after 18 h of HSP induction is not optimal and that freezing ASCs 3 h after heat treatment might be more efficacious. 6. No conclusions can be drawn, as yet, regarding the relationship between pre-freeze HSP induction and the post-freeze-thaw viability of ASCs. 7. Further studies, including western blotting are currently being pursed to assess the protein levels of HSP70 and HSP27 at various time points after the heat shock treatment (and to identify the peak time points and optimal time points for applying the freezing insult). 8. Future studies will also assess the effect of combinatorial upregulation of HSPs on the post-freeze-thaw viability of ASCs by overexpressing or knocking down certain HSPs using genetic engineering. 3.14 Some Observations on the Effect of Periodic Heat Shock on ASCs During Osteogenesis

1. The hypothesis was that the periodic upregulation or elevation of HSPs might be beneficial or detrimental to ASC osteogenesis. 2. ASCs that were heat treated (43 °C for 1 h) were periodically heat treated (on the 3rd, 9th, and 15th day) of osteogenic

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Fig. 8 Photograph of the commercially available differential scanning calorimeter, DSC (source: “Bioengineering Laboratory, Department of Mechanical Engineering, Louisiana State University, Baton Rouge, LA”). As described in the text, the DSC can be utilized to measure freezing process in adult stem cell suspensions; this biophysical knowledge of the freezing processes can subsequently be used to predict a priori the optimal rates of freezing

differentiation. The data suggests a decrease in osteogenesis staining with periodic heat treatment and decreased the quantified alizarin red value from 1.35 to 1.08 mM Alizarin red stain/9.5 cm2 well (control value was 0.4 mM). 3.15 Measurement of Slow Freezing Processes in Adult Stem Cells

1. The effect of freezing on the biophysical process during freezing (loss of water and cell shrinkage at “slow” cooling rates, denoted as water transport and the formation of intracellular ice at “fast” cooling rates, denoted as intracellular ice formation or IIF) of ASCs was investigated using a differential scanning calorimeter (DSC)—Fig. 8. 2. DSC experiments were carried out both in the absence and presence of CPAs on ASCs. For both SVF and ASCs (P0 and P2) cells, six separate DSC experiments were conducted in the absence of any CPAs and in the presence of two permeating CPAs: glycerol (10%, v/v) and DMSO (10%, v/v). 3. Approximately 10 μL of cell suspension was loaded in the DSC standard aluminum sample pan (Perkin Elmer Corporation, Norwalk, CT, USA) with ~0.5 mg of Pseudomonas syringe (ATCC, Rockville, MD, USA), a naturally occurring ice nucleating agent.

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4. The DSC dynamic cooling protocol used to measure the water transport out of the adipose-derived cells was the same as that reported in earlier studies on other cell and tissue systems [38, 39, 55–61, 109–112] and will only be briefly stated here. (i) Cool the sample along with 0.5–1 mg of P. syringae bacteria from 4 °C to -5 °C at a cooling rate 1 °C/min until extra cellular ice nucleates (observed as a sharp exothermic peak in the DSC). (ii) At the time of nucleation manually thaw the sample at a warming rate of 5 °C/min such that Tph (-0.53 °C) is reached (but not overshot). Note that -0.53 °C represents the phase change temperature of 0.3 mOsm isotonic solution (see Note 20). (iii) Cool the sample to -50 °C at a pre-selected cooling rate (5 °C/min or 10 °C/min or 20 °C/min). (iv) Thaw the sample at 100 °C/min to -0.53 °C. (v) Re-equilibrate the sample at -0.53 °C for 5 min. (vi) Cool the sample to -150 °C at a pre-selected high cooling rate (200 °C/min) to lyse all the cells (see Note 21). (vii) Repeat iv, v, and iii. 5. In the DSC technique, heat releases from the same cell suspension are measured: (i) during freezing of osmotically active (live) cells in media and (ii) during freezing of osmotically inactive (dead) cells in media. The difference in heat release measured between the two cooling runs was correlated to water transport and was denoted as Δqdsc. Should the cells be osmotically inactive or lysed prior to the start of the experiment, the DSC cooling protocol will measure no difference in heat release. 6. The heat release measurements of interest are Δqdsc and Δq(T)dsc which represent the total and fractional difference between the heat releases measured by integration of the heat flows during freezing of osmotically active (live) cells in media. Note that q(T) represents the temperature dependence of the measured heat release difference. 7. The heat release measurements qdsc and Δq(T)dsc obtained using the DSC software can now be used to determine the volume of the freezing cell (V) as a function of temperature, Δq ðT Þ T as: V ðT Þ = V o - Δq dsc ðV o - V b Þ ; here Vo represents the dsc initial isotonic cell volume and the final volume (Vb) in the medium with no CPAs was assumed to be the osmotically inactive cell volume, Vb (= 0.6Vo). 8. Once the volumetric shrinkage data has been determined from the calorimetric experiments, a well-established model of water

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transport developed by Kedem and Katchalsky was used to determine the membrane permeability parameters [113, 114]. Briefly, the Kedem and Katchalsky (K-K) model is dependent on three cell membrane parameters: Lp or the hydraulic conductivity which measures the volume flow induced by a hydrostatic pressure difference (ΔP); ω or the solute permeability which measures the solute flux induced by a concentration difference (Δπ); and δ or the reflection coefficient which describes the ‘relative permeability’ of the membrane to the solute (and can be expressed in terms of Lp and ωv ω as: δ = 1 - Lcpa , where vcpa is the mole fraction of the cryop protective agent). These three transport coefficients characterize the passive flux through a membrane via the K-K equations as: JV =

1 dV ðT Þ = L p þ Δp - RT ΔC w þ δΔC cpa A c dt

and J cpa =

1 dN cpa = ΔC cpa,ave ð1 - δÞJ V þ RT ωΔC cpa A c dt

where Jv represents the total flux across the cell membrane (m/s), Jcpa represents the solute (CPA) flux across the cell membrane (m/s), R is the universal gas constant (J/molK), T is the temperature (Kelvin), Δp represents the hydrostatic pressure difference between the extracellular and intracellular space, generally assumed to be zero in mammalian cells, ΔCw and ΔCcpa represent the concentration difference between the intracellular and extracellular space for water and CPA, respectively. Initially, both ΔCw and ΔCcpa are zero. As time progress, CPA starts to diffuse into the tissue establishing a gradient for both water and CPA. And finally, the ΔCcpa.ave represents the “log mean” osmolality of the permeable solute as: ΔC cpa , where C ocpa and C icpa represent the conΔC cpa,ave = ln

C ocpa C icpa

centration of the CPA outside and inside of the cell, respectively. The various assumptions made in the development of Mazur’s model of water transport are discussed in detail elsewhere [109–112]. 9. During freezing in the presence of extracellular ice, the value of Jcpa is zero as ω approaches zero and δ approaches one. 10. Additionally, Lp is described a function of Lpg and ELp as L p = L pg exp -

E Lp R

1 T

-

1 TR

where Lpg is the permeability

of the membrane to water at a reference temperature (TR = 273.15 K) and ELp is the apparent activation energy

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Table 2 Sub-zero permeability parameters of ASC cells when cooled at 20 °C/min in the presence of extracellular ice with an assumed value of 0.6Vo as the osmotically inactive volume. The predicted optimal cooling rates using the optimal cooling rate equation are also shown in the last column Lpg or Lpg[cpa] × 1015 m3/Ns (μm/min-atm)

ELp or ELp[cpa] kJ/ mol (kcal/mole)

R2 value

Bopt oC/min

43.4 (9.66) 63.3 (15.25) 57.4 (13.40)

0.992 0.994 0.994

59.2 25.8 33.5

41.0 (0.24) 53.7 (0.314) 55.1 (0.322)

43.1 (10.30) 64.0 (15.3) 58.9 (14.1)

0.997 0.994 0.994

96.7 25.3 30.2

No CPA 10% Glycerol 10.0% DMSO

111.5 (0.652) 50.5 (0.295) 51.3 (0.300)

168.8 (40.37) 64.6 (15.45) 61.1 (14.61)

0.995 0.993 0.994

22.1 23.3 24.2

No CPA 10% Glycerol 10.0% DMSO

120.0 (1.09) 39.0 (0.230) 50.0 (0.300)

177.8 (42.5) 51.0 (12.2) 61.5 (14.7)

0.995 0.993 0.994

21.1 29.3 22.1

Cell passage

Freezing media

SVF

No CPA 10% Glycerol 10.0% DMSO

23.1 (0.135) 56.4 (0.330) 59.0 (0.345)

P0

No CPA 10% Glycerol 10.0% DMSO

P2

P4

for the reference membrane permeability [38, 39, 55–61, 109– 112]. 11. A nonlinear least squares curve fitting technique was implemented using a computer program to calculate the membrane permeability parameters that best fit the volumetric shrinkage data as previously described [38, 39, 55–61, 109–112]. 12. All the curve fitting results presented have an R2 value greater than or equal to 0.99 indicating that there was a good agreement between the experimental data points and the fit calculated using the estimated membrane permeability parameters. 13. The cell membrane permeability parameter values water transport data in the absence and presence of CPAs are shown in Table 2 along with the model-simulated optimal cooling rates. 14. The model-simulated optimal cooling rates (Bopt) were determined using a generic optimal cooling rate equation developed by Thirumala and Devireddy [115] as: - 0:0546E Lp Þ ð SA B = 1009:5  exp L  = . opt

pg

WV

15. Figure 9 shows the water transport data and simulation for SVF cells using the best fit parameters in K-K equations at a cooling rate of 20 °C/min in culture medium with 10% glycerol (lefthand side figure) and in culture medium with 10% DMSO (right-hand side figure). The model-simulated equilibrium cooling response (equilibrium is achieved at each temperature

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Fig. 9 Volumetric response of P0 ASCs as a function of subzero temperatures obtained using the DSC in the presence of extracellular ice and glycerol (left-hand side figure), and in the presence of extracellular ice and DMSO (right-hand side figure). The filled circles represent the experimentally obtained water transport (volumetric shrinkage) at a cooling rate of 20 °C/min. The model-simulated dynamic cooling response at 20 °C/min is shown as a solid line and was obtained by using the best-fit membrane permeability parameters (Lpg and ELp or Lpg[cpa] and ELp[cpa]) (Table 1) in the water transport K-K equation (see text for further details). The model-simulated equilibrium cooling response obtained is shown as a dotted line in all the figures. The nondimensional volume is plotted along the y-axis and the subzero temperatures are shown along the x-axis. The error bars represent the standard deviation for the mean values of six separate DSC experiments (n = 6)

when the internal and external osmotic pressures are equal) is also shown as reference as dotted line (------). The equilibrium cooling response represents the volumetric shrinkage response of an ASC cooled infinitely slowly and is significantly different (>99% confidence level using the student’s t-test) from the dynamic water transport data obtained at 20 °C/min. 3.16 Measurement of Fast Freezing Processes in Adult Stem Cells

1. The calorimetric protocol developed to measure fast freezing processes or intracellular ice formation (IIF) is identical to the one detailed in earlier studies to measure water transport in cells and tissues [38, 39, 55–61, 109–112]. The only exception will be that the two heat releases of interest (initial, with live and final, with lysed tissue cells) will be measured at “higher or faster” cooling rates ranging from 10 to 50 °C/min (see Fig. 10). Note that the choice of the “high or fast” cooling rates is currently limited by the machine capabilities. 2. The DSC heat release for P2 ASCs frozen at 50 °C/min consists of three separate heat releases: (1) due to water transport or cellular dehydration, (2) IIF due to surface-catalyzed nucleation (SCN), and (3) secondary heat release due to residual water in the cell (or IIF due to volume-catalyzed nucleation or VCN). Expressing this mathematically: dqtot = dqdehyd + dqscn + dqvcn, where dqvcn is known and can be found from the DSC thermograms. Further, dqtot can be expressed as: dq dehyd dq tot

=

ðV i - V e Þ . ðV i - V f Þ

Adult Stem Cells Freezing Processes and Cryopreservation Protocols

1

KDSC IIF or h CM IIF

0

q [mJ/mg]

79

-5

-10

0.8 0.6

(h) 50 °C/min

0.4 0.2

(d) 50 °C/min -15

0

-10

-20 -30 -40 Temperature [°C]

0 0

-50

-10

-20 -30 -40 Temperature [°C]

-50

Fig. 10 Heat thermograms for qWT + IIF are shown for ASCs cooled at 50 °C/min (left-hand side image) in the absence of any CPAs along with a comparison of the IIF data at 50 °C/min using the heat thermograms to the corresponding data obtained from the cryomicroscopy experiments (right-hand side image). Within the figure three different cytocrit (α) values (0.2; filled circles, 0.4; filled triangles and 0.8; filled squares) are shown. Error bars represent standard error obtained from three separate experiments. See text for further details

3. Using previously reported DSC measured water transport parameters for P2 ASCs (see Table 2) of Lpg = 0.652 μm/ min-atm and ELp = 40.37 Kcal/mol it is possible to estimate the end volume at -15 °C, Ve as 0.84  Vi and as before Vb is assumed to be 0.6Vo. Simplifying and normalizing with respect and simplifying we have: to dqscn, 30c/min dq  ðT Þ  2 3dq tot

=

dq ðT Þdehyd 2dq dehyd

þ

dq ðT Þscn dq scn .

4. In this equation, the last term on the RHS represents the heat release ratio of interest, i.e., the percentage of tissue cells that underwent IIF (due to SCN) at each temperature T. The other two terms are known a priori: (1) the term on the LHS is calculated from the DSC thermograms and (2) the first term on the RHS is calculated from simulated water transport response (or from non-dimensionalized volume,VVðT0 Þ) at a cooling rate of 50 °C/min using the following relation: dq ðT Þdehyd,50c= min dq dehyd,50c= min

=

ðV o - V ðT Þ ðV o - V e Þ .

5. We can now predict the probability of ice formation (PIFhet) due to SCN from calorimetric data as: dq ðT Þ V bÞ %cells ðSCNÞor PIFhet = dq scn  ðVðVðTe . Note that V , V b e, Þ - V bÞ scn V ðT Þ and V 0 are known a priori, from either experimental data or prior numerical simulations. The % of cells undergoing IIF due to SCN at various subzero temperatures are shown in Fig. 10 (right-hand side image) along with corresponding cryomicroscopy data. 6. A mechanistic IIF model by Toner [116] states that for a thermodynamic system composed of identical biological cells,

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the total probability of intracellular ice formation (PIFtot) is a function of heterogeneous or surface-catalyzed (PIFSCN or PIFhet) and homogeneous or volume-catalyzed (PIFVCN or PIFhom) ice nucleation mechanisms, as: PIFtot = PIFSCN + (1 PIFSCN)  PIFVCN. 7. The probability of ice formation due to heterogeneous ice nucleation (PIFhet) also called surface-catalyzed nucleation PIFSCN = 1 (SCN) is given by [116–119]: exp -

1 B

T

Ns η N so ηo

AΩSCN T seed

T T fo

1=2

ðT f =T fo Þ

4

exp - κSCN

ΔT 2 T 3

dT ,

where T is the temperature, ΔT is the amount of super cooling, B is the cooling rate, V is the tissue cell volume, A is the surface area of the embedded tissue cell (all the other variables are described elsewhere [116–119]). In order to evaluate IIF (SCN) during freezing of cells, both the concentration and the temperature dependence of the viscosity, η is needed [116–119]. However, the dependence of viscosity on the concentration of cytoplasm is usually neglected and a power law dependence of water viscosity on temperature, T (K) is applied: ηðT Þ = 0:139 • ððT=225Þ - 1Þ - 1:64 × 10 - 3 Kg=ms. 8. The probability of ice formation due to homogeneous ice nucleation (PIFhom), also called volume-catalyzed nucleation (VCN), is described by the following equation [116–119]: PIFVCN = 1 - exp -

1 B

T T seed

ΩVCN V exp -

κVCN ΔT 2 T 3

dT .

PIFVCN or PIFhom occurs when the heterogeneous ice nucleation mode is suppressed by rapid cooling or in cases where cooling proceeds in the absence of extracellular ice. The two unknown homogeneous nucleation parameters (Ωhom and κhom) will be based on the widely accepted values obtained by Franks et al. [120] using saline droplets and a calorimeter. 3.17 Some Observations on the Measured Freezing Processes in ASCs

1. The dynamic portion of the cooling curve (region where the initial and final heat release thermograms are distinct and separate) is found to be between -0.6 °C and -14 °C in medium with no CPAs, between -3.1 °C and -30 °C in medium with 10% glycerol and between -3.2 °C and -30 °C in medium with 10% DMSO for SVF cells. 2. For SVF cells, although the predicted Lpg and Lpg[cpa] values using Vb = 0.4Vo and 0.8Vo are significantly different from each other and from the values obtained earlier using Vb of 0.6Vo, the values of ELp or ELp[cpa] are not significantly different and are within ±10% of each other. The only exception being the value of ELp[cpa] with 10% DMSO at an assumed Vb value of 0.8Vo. A similar result (significant difference in Lpg values and

Adult Stem Cells Freezing Processes and Cryopreservation Protocols

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negligible difference in ELp values for Vb = 0.4, 0.6, and 0.8Vo) was also observed with P0 and P2 ASCs. 3. Despite the differences in the predicted permeability (Lpg and Lpg[cpa]) values of SVF cells for different Vb values, a very similar value (±20%) is obtained for the predicted optimal cooling rate. For P0 ASCs frozen in the absence of CPAs, varying the assumed value of Vb from 0.6Vo to 0.8Vo had a significant effect (~50% lower) on the predicted optimal cooling rate while lowering the value of Vb from 0.6Vo to 0.4Vo had essentially no effect (~10% lower). Intriguingly, no such effect is seen in P2 ASCs frozen in the absence of CPAs. 4. For P0 and P2 ASCs frozen in the presence of CPAs, lowering or increasing the assumed value of Vb from 0.4Vo to 0.8Vo had some effect (±25% variation in the predicted optimal cooling rates; although the magnitudes are within ±6 °C/min of each other for corresponding freezing media). 5. The variation in the assumed value of Vb does not significantly alter the model-predicted optimal rates of freezing and further studies are needed to understand this lack of sensitivity in the value of predicted optimal cooling rates for ASCs to the assumed value of Vb. 6. To account for any errors and uncertainties in the measured cell diameter of the adipose-derived cells, we also investigated the effect of varying the surface area to volume ratio (S:V) by ±50%, from 0.12 to 0.06 and 0.18. As expected, the membrane permeability parameters decrease with increasing S:V ratio. This inverse relationship between S:V ratio and the predicted membrane permeability parameters (Lpg or Lpg[cpa] and ELp or ELp[cpa]) is not surprising, since K-K equations show that the change in the volume of the ASCs as a function of temperature) is proportional to the product of Lp and Ac. 7. There is a significant increase in the measured values of reference membrane permeability, Lpg (or Lpg[cpa]) (~100–170%) and the activation energy, ELp (or ELp [cpa]) (~8–65%) values for SVF cells obtained in the presence of CPAs when compared to values obtained in their absence, for all the assumed values of Vb. 8. Similarly for P0 ASCs there is an increase in the measured values of Lpg (or Lpg[cpa]) (~18–34%) and the activation energy, ELp (or ELp [cpa]) (~15–50%) in the presence of CPAs when compared to the values obtained in the absence of CPA for an assumed value of Vb of either 0.6Vo or 0.4Vo. However, when the assumed value of Vb is increased to 0.8Vo, the converse is found to be true for P0 ASCs, i.e., adding either glycerol or DMSO significantly decreased the measured values of the activation energy, ELp (or ELp [cpa]) (~33–50%) and the

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reference membrane permeability, Lpg (or Lpg [cpa]) (~9–32%) when compared to values obtained in their absence. 9. P2 ASCs showed a significant decrease in measured values of Lpg (or Lpg[cpa]) (~50–60%) and the activation energy, ELp (or ELp [cpa]) (~56–67%) for all Vb values when compared to values obtained in the absence of CPAs. 10. Clearly, the relative effects of the composition of the freezing media and the assumed value of the inactive cell volume had different effects on the predicted permeability parameters for different passages of ASCs. 11. For SVF and P0 ASCs at Vb = 0.6Vo (as shown in Table 2), the optimal cooling rates in the presence of either 10% glycerol or 10% DMSO are significantly smaller (~43–74%) than the predicted optimal values in their absence. 12. Interestingly, at Vb = 0.6Vo, the addition of CPAs increases the optimal freezing rates by ~5–10% for P2 ASCs. At Vb = 0.4Vo, the presence of CPAs significantly decreased the optimal cooling rates by ~42–50% (for SVF cells), by ~66–68% (for P0 ASCs), and by ~15–18% (for P2 ASCs), whereas at Vb = 0.8Vo, the presence of CPAs the optimal cooling rates were reduced by ~30–50% (for SVF cells), by ~20–40% (for P0 ASCs), and by ~21–23% (for P2 ASCs). 13. The significant decrease in the predicted optimal cooling rates for adipose-derived cells (except for P2 ASCs cells at Vb = 0.6Vo) in the presence of CPAs is surprising as the presence of CPAs is expected to increase the ability of the cell membrane to dehydrate at faster cooling rates as shown for several mammalian cells [109–112]. 14. Biophysical properties used to characterize the thermodynamic state of intracellular solution during freezing, such as water transport (Lpg and ELp), heterogeneous (Ωhet and κhet), and homogeneous (Ωhom and κhom) ice nucleation, can be used to predict or narrow the range of optimal rates of freezing with and without the presence of CPAs.

4

Notes 1. The outcome of any cryopreservation procedure will depend on the quality of the pre-frozen sample. Therefore, cells harvested for cryopreservation should be at their optimum viability to ensure maximal survival during freezing and after thawing. 2. Commercially available programmable controlled rate freezer is costly ($20,000). Cheaper alternatives might be a -80 °C freezer.

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3. Tissue culture media are highly species specific. Cryoprotectants and growth media components must be of high purity (analar-grade and spectroscopic grade for DMSO). 4. Where possible cryoprotectant mixtures should be sterilized by filtration or for high viscosity solutions by autoclaving. 5. The concentration, duration, and temperature of the addition of the CPAs is a critical factor in the success of the whole procedure. Care should be taken to minimize the time of exposure to CPAs. Once the cells have been prepared for freezing, they should be frozen as soon as possible. 6. DMSO freely penetrates the skin, can be irritating to the eyes and skin. 7. Storage in the vapor phase of the liquid nitrogen is recommended. However, storage in the liquid phase does prevent the possibility of inadvertent warming. 8. Extreme caution should be exercised while removing the samples from the liquid phase as the penetration of liquid nitrogen into defective containers can cause excessive pressure (and possibly, explosive) buildup of gas in a confined space during the thawing process. To reduce this possibility, cryovials should be allowed to equilibrate in the vapor phase before being transferred to a warm water bath for thawing. It might also be advisable to place the cryovials in a “closed metal container” (to contain any exploded material for a simpler decontamination procedure). 9. Storage temperature needs to be maintained below -139 °C. 10. To retain maximum viability, cells should be thawed rapidly and uniformly. But carefully, so that the maximum temperature does not exceed normal temperature range. 11. After thawing it is necessary to remove or slowly dilute the cryoprotectant to prevent osmotic shock. DMSO will evaporate from the medium at 37 °C. 12. Minimal g forces need to be applied during centrifugation. 13. Storage time does not influence the viability of the stored material. Therefore, the whole duration of low temperature storage can be as little as 1 h, greatly reducing the length of the whole procedure. 14. Physical factors influencing the success of a cryopreservation procedure include the imposed cooling rate, the composition of the freezing media, the time of exposure to the cryoprotectants, the nature and sequence of the cryoprotectant addition and removal procedures, the storage temperature, and the thawing process. All of these variables need to be carefully controlled during the freezing process to ensure an optimal outcome.

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15. The presence of spurious air bubbles during the pouring of the resolving gel can be eliminated by the use of a small amount of distilled water or EtOH on the top surface of the resolving gel. However, the presence of spurious or damaging air bubbles during the stacking gel casting can be eliminated by removing some of the stacking gel and replenishing it. To eliminate the possibility of the gel solidifying during this process, it is imperative to insert as carefully as possible in the minimum amount of time. 16. Improper removal of the gel combs from the top of the polymerizer stacking gel will result in breakage or damage the structural integrity of the wells resulting in a catastrophic failure of the casting process and initiating the process once again from the initial pouring of the resolving gel step. 17. Heating the cell lysate (proteins) plus loading buffer for more than 5 min will result in precipitation of proteins or degraded proteins that are unsuitable for the western blot assay. 18. Gently centrifuge the sample to minimize loss of sample via condensation on the tube walls. The protein samples should be carefully loaded to avoid spillage and contamination between wells. 19. Extreme care should be taken to avoid any trapped air or air bubbles between the nitrocellulose membrane and the SDS-PAGE gel during the protein transfer process. Care should also be taken to avoid contaminating with spurious proteins onto the nitrocellulose membrane during the disassembly and subsequent handling process. 20. The phase change temperature is given by the equation: mOsm * –1.858. Hence for a 0.285 mOsm solution, the phase change temperature is -0.53 °C. 21. The choice of the high cooling rate is dependent on the cell type and is at a rate that induces damaging intracellular ice formation. This lyses or renders the cells osmotically inactive during subsequent DSC heat transfer measurements.

Acknowledgments The author thanks Dr. Elizabeth Clubb and Dr. James Wade at the Pennington Biomedical Research Center (PBRC) for supplying the liposuction aspirates and their many patients for consenting to participate in this protocol; Marilyn Dietrick of the LSU School of Veterinary Medicine Flow Cytometry Core Facility; Prof. Jeffrey Gimble, Gang Yu, Xiying Wu, of the Stem Cell Biology Laboratory at the Pennington Biomedical Research Center (PBRC), and the clinical nutrition research unit (CNRU) Molecular Mechanism

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Core at PBRC for their technical assistance. Acknowledgments are also due to Dr. Gimble, a longtime colleague and without his help this work would never have been initiated. In addition, acknowledgments are also due to Dr. S. Thirumala (freezing experiments), R. Fuller (comparing the freezing devices, CRF and DSS), Dr. S. Shaik (HSP experiments) results reported in this chapter. This work was supported in part by funding from the Louisiana Board of Regents, the Department of Mechanical Engineering at the Louisiana State University, the institute of Diabetes and Digestive and Kidney Diseases of the National Institutes of Health (NIH) under award number R21DK91852 and by the institute of General Medical Sciences of the NIH under award number R15GM141653. References 1. De Ugarte DA, Morizono K, Elbarbary A et al (2003) Comparison of multi-lineage cells from human adipose tissue and bone marrow. Cells Tissues Organs 174:101–109 2. Kang SK, Putnam L, Dufour J et al (2004) Expression of telomerase extends the lifespan and enhances osteogenic differentiation of adipose tissue-derived stromal cells. Stem Cells 22:1356–1372 3. Mitchell JB, McIntosh K, Zvonic S et al (2006) Immunophenotype of human adipose-derived cells: temporal changes in stromal-associated and stem cell-associated markers. Stem Cells 24:376–385 4. Rubio D, Garcia-Castro J, Martin MC et al (2005) Spontaneous human adult stem cell transformation. Cancer Res 65:3035–3039 5. Wall ME, Bernacki SH, Loboa EG (2007) Effects of serial passaging on the adipogenic and osteogenic differentiation potential of adipose-derived human mesenchymal stem cells. Tissue Eng 13:1291–1298 6. Gimble JM, Katz AJ, Bunnell BA (2007) Adipose-derived stem cells for regenerative medicine. Circ Res 100:1249–1260 7. Zhang FB, Li L, Fang B et al (2005) Passagerestricted differentiation potential of mesenchymal stem cells into cardiomyocyte-like cells. Biochem Biophys Res Commun 336: 784–792 8. Tsutsumi S, Shimazu A, Miyazaki K et al (2001) Retention of multilineage differentiation potential of mesenchymal cells during proliferation in response to FGF. Biochem Biophys Res Commun 288:413–419 9. Zeng X, Rao MS (2007) Human embryonic stem cells: long term stability, absence of senescence and a potential cell source for

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and human adipose-derived stromal cells. Biochem Biophys Res Commun 294:371–379 90. Gronthos S, Franklin DM, Leddy HA et al (2001) Surface protein characterization of human adipose tissue-derived stromal cells. J Cell Physiol 189:54–63 91. Thirumala S, Forman JM, Monroe WT et al (2007) Freezing and post-thaw apoptotic behaviour of cells in the presence of palmitoyl nanogold particles. Nanotech 18:195104 92. Rubinsky B, Ikeda M (1985) A cryomicroscope using directional solidification for the controlled freezing of biological mat. Cryobiology 22:55–68 93. Garner DL, Johnson LA (1995) Viability assessment of mammalian sperm using SYBR-14 and propidium iodide. Biol Reprod 53:276–284 94. Takamatsu H, Rubinsky B (1999) Viability of deformed cells. Cryobiology 39:243–251 95. Takamatsu H, Takeya R, Naito S, Sumimoto H (2005) On the mechanism of cell lysis by deformation. J Biomech 38:117–124 96. Wolfe J, Bryant G (1999) Freezing, drying, and/or vitrification of membrane- solutewater systems. Cryobiology 39:103–129 97. Rubinsky B (2000) Cryosurgery. Annu Rev Biomed Eng 2:157–187 98. Lindquist S (1986) The heat-shock response. Annu Rev Biochem 55:1151–1191 99. Young JC, Agashe VR, Siegers K et al (2004) Pathways of chaperone-mediated protein folding in the cytosol. Nat Rev Mol Cell Biol 5:781–791 100. Kalmar B, Greensmith L (2009) Induction of heat shock proteins for protection against oxidative stress. Adv Drug Deliv Rev 61:310– 318 101. Beere HM, Wolf BB, Cain K et al (2000) Heat-shock protein 70 inhibits apoptosis by preventing recruitment of procaspase-9 to the Apaf-1 apoptosome. Nat Cell Biol 2:469–475 102. Saleh A, Srinivasula SM, Balkir L et al (2000) Negative regulation of the Apaf-1 apoptosome by Hsp70. Nat Cell Biol 2:476–483 103. Li CY, Lee JS, Ko YG et al (2000) Heat shock protein 70 inhibits apoptosis downstream of cytochrome c release and upstream of caspase3 activation. J Biol Chem 275:25665–25671 104. Pandey P, Saleh A, Nakazawa A et al (2000) Negative regulation of cytochrome c-mediated oligomerization of Apaf-1 and activation of procaspase-9 by heat shock protein 90. EMBO J 19:4310–4322 105. Gorman AM, Szegezdi E, Quigney DJ et al (2005) Hsp27 inhibits 6-hydroxydopamine-

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Part II Animal ASC Models

Chapter 6 Isolation of Murine Adipose-Derived Stromal/Stem Cells for Adipogenic and Osteogenic Differentiation or Flow Cytometry-Based Analysis Matthew C. Scott, Chul-Hong Park, Marilyn Dietrich, Xiying Wu, Jeffrey M. Gimble, Carrie M. Elks, Ji Suk Chang, and Z. Elizabeth Floyd Abstract Murine models of obesity or reduced adiposity are a valuable resource for understanding the role of adipocyte dysfunction in metabolic disorders. Adipose tissue stromal vascular cells or primary adipocytes derived from murine adipose tissue and grown in culture are essential tools for studying the mechanisms underlying adipocyte development and function. Herein, we describe methods for the isolation, expansion, and long-term storage of murine adipose-derived stromal/stem cells, along with protocols for inducing adipogenesis to white or beige adipocytes in this cell population and osteogenic differentiation. Isolation of the adipose stromal vascular fraction cells for flow cytometric analysis is also described. Key words Murine, Adipose-derived stromal/stem cells (ASCs), Collagenase, Isolation, Mesenchymal stem cells (MSCs), Hematopoietic stem cells (HSCs), Stromal vascular fraction (SVF), Adipogenesis, Beige adipogenesis, Osteogenesis, Flow cytometry

1

Introduction Adipose tissue can be separated into mature adipocytes and a stromal vascular fraction (SVF) containing a heterogeneous mesenchymal cell population that includes hematopoietic stem cells, endothelial cells, erythrocytes, fibroblasts, pericytes, myeloid and lymphoid cells, and adipose stromal cells [1, 2]. When the heterogeneous SVF is cultured, the adherent cells form a more homogeneous cell population of mesenchymal stem/progenitor cells that are referred to as adipose-derived stromal/stem cells (ASCs). Murine ASCs are commonly used in basic research and can undergo adipogenic, osteogenic, chondrogenic, myogenic, and neuronal differentiation in vitro [3–6]. In particular, ASCs from murine inguinal adipose tissue were recently used to study the molecular control of beige adipocyte formation in white adipose tissue [7, 8].

Jeffrey M. Gimble et al. (eds.), Adipose-Derived Stem Cells: Methods and Protocols, Methods in Molecular Biology, vol. 2783, https://doi.org/10.1007/978-1-0716-3762-3_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Mesenchymal stem/progenitor cells (MSCs) were originally characterized from bone marrow. However, harvesting MSCs from adipose tissue has several advantages over acquiring these cells from the bone marrow, including better accessibility and the greater abundance of the MSCs in adipose tissue. Thus, murine ASCs are a valuable resource for studying mechanisms controlling the differentiation potential of MSCs or studies specifically focused on adipogenesis, the relationship between adipogenesis and osteogenesis, or the role of mature adipocytes in metabolic disorders related to obesity. Recent data clearly establishes that obesity is often accompanied by an influx of immune cells into the adipose tissue, leading to chronic inflammation of the adipose tissue and compromised metabolic health (reviewed in [9, 10]). In obesity, adipose tissue SVF cells may primarily be composed of immune cells, including macrophages and T and B lymphocytes. Consequently, understanding the contribution of these adipose tissue immune cells to obesity-related health problems has become an essential component of understanding adipose tissue biology. As these cells are commonly analyzed via flow cytometric analysis of cell surface markers, steps for isolating murine stromal vascular cells for flow cytometry are included as part of a protocol describing the isolation and expansion of murine ASCs and the differentiation of the ASCs into white or beige adipocytes or osteoblasts in vitro.

2

Materials

2.1

Tissue

2.2

Supplies

White adipose tissue depots: Perigonadal (epididymal in males, parametrial in females), inguinal, retroperitoneal (along the dorsal abdominal wall, near the kidney, but distinct from perirenal fat), mesenteric (along the intestines, containing multiple lymph nodes). The protocol described herein is used for adipose tissue from any of these depots. ASCs from the inguinal fat depot more readily differentiate to white adipocytes in vitro compared to perigonadal fat. Inguinal fat is also the preferred tissue for differentiation to beige adipocytes. A visual guide to dissecting specific fat depots in mouse models, including omental fat in the severely obese mouse, is provided by Bagchi and MacDougald [11]. 1. 50 mL polypropylene conical tubes, sterile. 2. 0.22 μm filter units. 3. 2 mL microtubes, autoclave to sterilize. 4. 35 mm tissue culture plates. 5. 100 μm cell strainer, sterile. 6. 15 mL polystyrene conical tubes, sterile.

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7. Freezing apparatus (Nalgene Cryo freezing container or similar). 8. Hemocytometer to count cells. 9. Scissors and forceps, autoclave to sterilize. 10. Additional supplies, if isolating the adipose tissue SVF for flow cytometry applications. (a) 12-well tissue culture plate, sterile. (b) 5 mL polystyrene round bottom tubes. (c) 5 mL polystyrene round bottom tubes with 40 μm strainer caps. 2.3

Equipment

1. Inverted microscope. 2. Shaking water bath or shaking incubator at 37 °C. 3. Centrifuge, 4 °C and room temperature. 4. Biosafety hood. 5. 5% CO2 humidified incubator. 6. Sterilized surgical instruments.

2.4 Reagents and Buffers

1. 70% Ethanol. 2. 0.25% Trypsin/EDTA solution. 3. Ammonium-Chloride-Potassium (ACK) Red Cell Lysis Buffer. 4. Ascorbic acid. 5. β-Glycerol phosphate. 6. Biotin. 7. Bovine Serum Albumin (BSA). BSA Fatty Acid Free for flow cytometry. 8. Collagenase Type 1. 9. Dexamethasone. 10. Dimethyl sulfoxide. 11. DMEM/F12 (1:1) media. 12. DMEM, high glucose media. 13. D-pantothenic acid. 14. Fc Receptor Block (anti-mouse CD16/CD32, purified; flow cytometry). 15. Fetal Bovine Serum (FBS). 16. Hanks Buffered Saline Solution (HBSS) (CaCl2 and MgCl2 free). 17. Indomethacin. 18. Insulin.

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19. Isopropanol. 20. MEM alpha. 21. 3-isobutyl 1-methyl xanthine (IBMX). 22. Oil Red O. 23. Paraformaldehyde (flow cytometry). 24. Penicillin/Streptomycin antibiotic solution (see Note 1). 25. Phosphate-buffered saline (PBS), pH 7.4 (CaCl2 and MgCl2 free). 26. Rosiglitazone. 27. 3,3′,5-Triiodo-L-Thyronine (T3). 28. Trypan Blue, 0.4% solution. All buffers are sterile-filtered using a 0.22 μm filter unit. 1. Phosphate Buffered Saline (PBS): 37 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4 pH 7.4 (Sterile solutions of PBS are available commercially). 2. Collagenase solution: Weigh out 10 mg of type 1 collagenase and dissolve in 10 mL HBSS or PBS with 2% BSA. After sterile filtration, warm the solution to 37 °C. This solution should be used within 1 h of its preparation. 3. FACS staining buffer: 0.1% Fatty Acid Free Bovine Serum Albumin (BSA, fraction V) in PBS. Sterile filtered using a 0.22 μm filter. Several flow cytometry staining buffers are commercially available. 2.5 Media and Staining Solutions

All solutions are sterile-filtered using a 0.22 μm filter unit. 1. Stromal medium: DMEM/F12 with 15% fetal bovine serum (FBS) and 2× antibiotic solution from 100× stock (penicillin/ streptomycin). 2. White adipocyte differentiation medium: Prepare and aliquot the following stock solutions in advance and store frozen at 20 °C until required. (a) A 66 mM stock solution of biotin (2000-fold concentration) dissolved in 1 N sodium hydroxide. (b) A 34 mM stock solution D-pantothenate (2000-fold concentration) dissolved in water. (c) A 1 mM dexamethasone (1000-fold concentration) dissolved in water or ethanol, depending on its formulation (see Note 2). (d) A 5 mM stock solution of rosiglitazone or equivalent PPARγ agonist dissolved in dimethyl sulfoxide (1000fold concentration).

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Prepare fresh before use: (e) A 500 mM stock solution of IBMX (1000-fold concentration) dissolved in 50 mM NaOH. (f) A 200 μM stock solution of insulin (2000-fold concentration) dissolved in 10 mM HCl, made in plastic tube, not glass. (g) The base medium for white adipocyte differentiation is DMEM, high glucose with 1× antibiotic solution (from 100× stock of penicillin/streptomycin) combined with DMEM/F12, 10% FBS with 1× antibiotic solution at a ratio of 7 parts DMEM, high glucose to 3 parts DMEM/ F12, 10% FBS to yield a 3% final concentration of FBS. (h) Prepare the white adipocyte differentiation medium by supplementing the base medium with 0.5 mM IBMX, 1 μM dexamethasone, 100 nM insulin, 5 μM rosiglitazone (or equivalent PPARγ full agonist), 33 μM biotin and 17 μM D-pantothenate. 3. White adipocyte maintenance medium: This solution is prepared in an identical manner as differentiation medium except that it does not contain either the IBMX or rosiglitazone; these two stock solutions should be omitted. Use this solution within 2 weeks of its preparation. 4. Beige adipocyte differentiation medium: In advance, prepare and aliquot the following stock solutions and store frozen at -20 °C until required. (a) A 5 mg/mL stock solution of insulin (1000-fold concentration) dissolved in 0.02 N sodium chloride. (b) A 10 mM T3 (107-fold concentration) dissolved in 0.02 N sodium hydroxide. (c) A 5 mM dexamethasone (1000-fold concentration) dissolved in water or ethanol, depending on its formulation. (d) A 62.5 mM indomethacin (500-fold concentration) dissolved in ethanol. (e) A 75 mM IBMX (150-fold concentration) dissolved in ethanol. (f) A 30 mM rosiglitazone (60,000-fold concentration) dissolved in dimethyl sulfoxide. (g) Prepare the beige adipocyte differentiation medium by supplementing the base medium (see Subheading 2.5, item 2) with 5 μg/mL insulin, 1 nM T3, 5 μM dexamethasone, 125 μM indomethacin, 0.5 mM IBMX, and 0.5 μM rosiglitazone. 5. Beige adipocyte maintenance medium: This solution is prepared in an identical manner as beige adipocyte differentiation

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medium but only contains insulin and T3. Use this solution within 2 weeks of its preparation. 6. Osteoblast differentiation medium: In advance, prepare and aliquot the following stock solutions and store frozen at 20 °C until required. (a) 1 M stock solution of β-glycerol phosphate (100-fold concentration) dissolved in PBS. (b) 0.5 M stock solution ascorbic acid (2500-fold concentration) dissolved in PBS. (c) 1 mM dexamethasone stock solution (10,000-fold concentration) (see Note 2). (d) Prepare the osteoblast differentiation media as MEM α, 10%FBS, 1× antibiotic solution (from 100× stock of penicillin/streptomycin), 10 mM β-Glycerol Phosphate, 0.2 mM ascorbic acid, and 0.1 μM dexamethasone. 7. Freezing medium: The freezing medium consists of 15% FBS, 75% DMEM/F12, and 10% dimethyl sulfoxide. Use this solution within 2 weeks of its preparation. 8. Oil Red O staining solution: Weigh out 0.7 grams of Oil Red O. Dissolve in 200 mL isopropanol. Filter through a 0.45 μm filter. Store at room temperature as stock solution. To stain cells, make a 60% Oil Red O solution in distilled water by adding 30 mL of the Oil Red O stock to 20 mL of distilled water. Mix and let stand at room temperature for 30 min before use and filter again if particulates are present. Use this solution within 24 h of its preparation.

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Methods

3.1 Obtaining the Adipose Tissue

1. Ensure the ASC isolation procedure is initiated as soon as possible after the tissue is extracted. This is especially important to consider if a large number of samples are being obtained. 2. Warm the collagenase solution and stromal media in the 37 °C water bath. 3. Prepare a labeled sterile 50 mL tube containing sterile PBS with 2× antibiotic solution and record the weight. Place the tubes with PBS on ice. 4. Place bench protector down in biosafety hood. 5. Sacrifice animal using anesthesia or CO2 asphyxiation as approved by the AVMA guidelines on euthanasia. Saturate the fur with 70% ethanol and place the animal ventral side up. 6. Using sterile tissue forceps, pull up on the skin and cut the skin vertically toward the head.

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7. Using sterile scissors and forceps, open the abdominal cavity and remove the desired fat pads. When isolating the inguinal fat pad, be sure to remove lymph nodes. When isolating perigonadal fat pads, remove any gonadal tissue. 8. Place the tissue in the preweighed labeled sterile 50 mL tube containing enough sterile PBS with 2× antibiotic solution to cover the tissue. Maintain the tube/tissue on ice until all samples are harvested. 3.2 Processing the Adipose Tissue

1. Working under a laminar flow hood, use sterile forceps to transfer the tissue to the bottom of a sterile 35 mm tissue culture dish containing enough ice-cold sterile HBSS or PBS to rinse tissue well. 2. Transfer the rinsed tissue to the lid of the sterile 35 mm tissue culture dish on ice and mince the adipose tissue into small pieces using sterile scissors. 3. Place the minced tissue in sterile 50 mL centrifuge tubes. If digesting less than 2.0 grams adipose tissue, use a minimum of 10 mL collagenase solution to ensure complete digestion. If digesting ≥2 grams tissue, add 5 mL collagenase solution/ gram adipose tissue to each centrifuge tube (see Note 3). 4. Cap and wipe the centrifuge tubes with 70% ethanol, dry, then wrap with parafilm before placing the tubes in a 37 °C shaking water bath or incubator. Shake at 100 rpm for up to 60 min. The tissue should appear smooth on visual inspection. If a shaking water bath or incubator is unavailable, vigorously shake the bottles every 5–10 min in a 37 °C water bath.

3.3 Isolating the Stromal Vascular Fraction (SVF)

1. After the collagenase digestion, filter solution through a sterile 100 μm filter into a sterile 50 mL conical tube. Centrifuge the tubes at 450 × g at room temperature for 5 min. The stromal vascular fraction will be apparent as the dark red cells pelleted on the bottom. Carefully aspirate the oil on top and the primary adipocytes, which will appear as a yellow layer of floating cells (see Notes 4 and 5). Leave a small amount of the brown collagenase solution so that the pelleted stromal vascular fraction is not disturbed. 2. Resuspend the cells in 2 mL ACK red cell lysis buffer with gentle vortexing or gentle trituration. Incubate at room temperature for 3–5 min. 3. Add 10 mL sterile HBSS or PBS (CaCl2 and MgCl2 free) to neutralize the ACK red cell lysis buffer. 4. Centrifuge the resuspended cells at 450 × g for 5 min and carefully aspirate the remaining collagenase solution. When aspirating, the tip of the pipette should aspirate from the top

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so that the oil is removed as much as possible. The stromal cell pellet should be tightly packed at the bottom of the tube. 5. If isolating the stromal vascular fraction cells for primary cell culture: (a) Add 2–3 mL of stromal media per gram adipose tissue (weight when harvested) to each centrifuge tube and resuspend the cells again with gentle vortexing or gentle trituration. (b) Plate the SVF obtained from 0.5 to 1.0 gram adipose tissue in a 35 mm tissue culture dish or a single well of a 6-well plate. (c) 24 h after plating, aspirate the media from the plate, rinse the cells with PBS prewarmed to 37 °C and aspirate. Check the cells using the inverted microscope; if excess unattached cells remain after the first PBS rinse, repeat the rinse and aspirate. (d) Add fresh stromal media. (e) The media is changed every 2–3 days until the cells are 80–90% confluent. The cells may begin to detach if the media is removed completely; leave a small amount of media on the cells with each media change. At 80–90% confluence, the cells are either passaged, stored as frozen stocks as passage one (P1) or differentiated. 6. If isolating the stromal vascular fraction cells for flow cytometry: (a) The cells must be kept ice cold from this point onward. (b) Add 1 mL PBS (CaCl2 and MgCl2 free) and resuspend the cells with a transfer pipette. (c) Filter the resuspended cells into a 5 mL polystyrene round bottom tube with a 40 μm cell strainer cap. (d) Use an aliquot of the cells diluted with trypan blue (a 1: 2 or 1:4 dilution is usually convenient) to count the cells using a hemocytometer. (e) See Subheading 3.7 for flow cytometry staining steps. 3.4 Differentiation of the ASCs to White Adipocytes

When the cells reach about 90% confluence, the murine ASCs are ready to differentiate (see Note 6). Exchange the stromal media for the differentiation media and leave on the cells for 3 days. On day 3 post-induction, replace the differentiation media with the adipocyte maintenance media. The adipocyte maintenance media is changed every 2–3 days and mature white adipocytes should be formed by day 8–12 post-induction (Fig. 1).

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Fig. 1 Oil Red O staining of neutral lipid accumulation in murine adipose-derived stromal cells. ASCs were obtained from the inguinal adipose tissue of 4-week-old wild-type male mice. The ASCs were at passage number 2 and adipogenesis media was added after the cells reached 80–90% confluence. Neutral lipid staining was carried out in the uninduced and induced ASCs at day 8 post-induction 3.5 Differentiation of the ASCs to Beige Adipocytes

When the ASCs isolated from inguinal adipose tissue reach about 95% confluence, exchange the stromal media for the beige adipocyte differentiation media (see Subheading 2.5, item 4) and leave on the cells for 2 days. On day 2 post-induction, replace the differentiation media with the beige adipocyte maintenance media supplemented with rosiglitazone (0.5 μM) and incubate the cells for 2 days. After that, the beige adipocyte maintenance media containing rosiglitazone (1 μM) is changed every 2 days and mature beige adipocytes should be formed by day 6–7 post-induction (Fig. 2).

3.6 Differentiation of the ASCs to Osteoblasts

When ASCs reach 80–90% confluence, exchange stromal media for osteoblast differentiation media (see Subheading 2.5, item 6). Replace media every 48-h and maintain culture to intended endpoints (see Note 7). ASCs differentiated towards osteoblasts will begin forming mineralized plaques after 10–20 days [12, 13].

3.7 Flow Cytometry Staining of the SVF Cells

For a more detailed description of FACS analysis of adipose tissue immune cell populations, including antibody selection and gating strategies, see Grant et al. [14] and Cho et al. [15]. 1. Keep everything ice cold. Prechill centrifuge to 4 °C. 2. Place 1–2 × 106 SVF cells in a 5 mL polypropylene tube. 3. Centrifuge the cells at 450 × g for 5 min at 4 °C. 4. Carefully pour off the supernatant (do not tap the tube). A small amount of supernatant will remain, and the pellet will be clearly visible. Disperse the cell pellet by adding 100 μL flow cytometry staining buffer followed by gentle trituration.

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Fig. 2 Quantitative real-time PCR shows the upregulation of beige adipocyteselective genes. ASCs were obtained from the inguinal adipose tissue of 5-week-old BL6 male mice. The ASCs were at passage number 2 and beige adipocyte differentiation media was added to induce differentiation after the cells reached 95% confluence. RNA was isolated from ASCs prior to induction (n = 2) and at day 7 post-induction (n = 6) and analyzed for expression of adipogenic marker genes (Fabp4, Pparg) and beige-selective genes (Ucp1, Dio2, Elovl3, Cidea, and Cox8b). Relative expression of mRNAs was determined after normalization to cyclophilin

Immediately add 2 μL Fc block and mix gently. If staining a large number of samples, a master mix of 100 μL staining buffer +2 μL Fc block per sample can be prepared and 102 μL of the master mix transferred to each sample tube. After Fc block is added, incubate samples on ice for 15 min. 5. Make a master mix containing all of the antibodies that will be used. The optimal concentration of each antibody should be determined for the stromal cell population via a titration assay prior to the experiment. Add 2 μL of each antibody per sample to the master mix. Staining buffer can be added to the master mix to allow a larger volume to be dispensed to each sample tube (see step 6 below). 6. Add the antibody master mix to each tube at a total volume equivalent to 2 μL of each antibody plus amount of staining buffer added per sample (i.e., add 30 μL master mix if using 5 antibodies and 20 μL sample buffer). Mix the samples well by vortexing. Place on ice and cover with foil (or place in dark refrigerator) to protect from light for 30 min (monoclonal antibodies) or 45 min (polyclonal antibodies). 7. Add 1 mL ice cold CaCl2 and MgCl2-free PBS and centrifuge at 450 × g for 5 min at 4 °C. Pour off supernatant without tapping tube. A small amount of supernatant will remain. Disperse the pellet by adding 2 mL ice cold CaCl2 and MgCl2-free PBS followed by gentle vortexing or trituration. 8. Centrifuge at 450 × g for 5 min at 4 °C, then remove supernatant.

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Fig. 3 Representative dot blots of murine stromal vascular fraction (SVF) cells. SVF cells were isolated from the epididymal adipose tissue of a 5-month-old wild-type mouse maintained on a high fat diet and analyzed on a BD Fortessa flow cytometer. (a) Total ungated SVF population. (b) Leukocytes were identified from the ungated population by CD45 positivity (boxed region). (c) Macrophages were identified (boxed region) from the CD45+ region by positive staining for CD64. (d) The CD64+ macrophages were divided into M1-like and M2-like populations based on CD11b and CD11c staining. M1-like macrophages (CD11b and CD11c double positive) appear in the upper right quadrant. M2-like macrophages (CD11b positive and CD11c negative) appear in the lower right quadrant. Monoclonal antibodies used were anti-CD45-SB600, anti-CD64-PE, anti-CD11b-BB515, and anti-CD11c-BV711

9. Add 100 μL 1% paraformaldehyde and gently vortex. If cells are to be sorted live (FACS) for downstream applications, add 100 μL CaCl2 and MgCl2-free PBS instead of 1% paraformaldehyde. 10. Paraformaldehyde-fixed samples may be stored in the dark at 4 °C until analyzed; live cells must be sorted via FACS as soon as possible after staining (Fig. 3).

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3.8 Harvesting the Plated Cells

1. Remove media from tissue culture plate and rinse with 1.5–2 mL prewarmed sterile PBS if using a T75 flask (75 cm2 growth area). As a rule of thumb, use 0.025 mL PBS/cm2 growth area of the plate. 2. Aspirate the PBS and add a minimum 0.025 mL/cm2 growth area of 0.25% Trypsin/EDTA solution. For example, if working in a 6-well plate format (9.5 cm2 growth area), add 0.250 ml 0.5% Trypsin/EDTA solution. If working in a 100 mm plate format (55 cm2 growth area), use at least 1.375 mL 0.25%Trypsin/EDTA solution. 3. Incubate in 37 °C incubator for 2–5 min. Do not allow the cells to remain in contact with the Trypsin/EDTA solution longer than is necessary to detach the cells. When more than 90% of the cells have detached (verify using the inverted microscope), add a sufficient volume of stromal media to completely resuspend the cells. 4. Transfer the resuspended cells to a sterile 15 mL tube and centrifuge at 450 × g for 5 min at room temperature. Aspirate the media and resuspend the pelleted cells in 1–2 mL of stromal media. Use an aliquot of the cells and dilute with Trypan blue (a 1:2 or 1:4 dilution is usually convenient) to count the cells using a hemocytometer.

3.9 Replating or Cryopreserving the Plated Cells

Replating the cells: Suspend the cells in stromal media and replate at 5000 cells/cm2 for further passages or experiments (see Note 8). Cryopreservation of the cells: Centrifuge the cells once more to obtain a cell pellet. Resuspend the pelleted cells in room temperature freezing media at a concentration of 1–2 × 106 cells per ml. Dispense 1 mL aliquots of the resuspended cells to sterile cryovials. Place the vials in the freezing apparatus (Nalgene Cryo freezing container) and place at -80 °C overnight. Transfer to long-term storage in liquid nitrogen the next day.

3.10 Cell Fixation and Oil Red O Detection of Neutral Lipids

1. Cell fixation: To fix plated cells for Oil Red O staining, the media is removed, and the cells are rinsed in PBS and the PBS is aspirated before adding an amount of 10% buffered formalin sufficient to completely cover the cells. Incubate the cells in the formalin for at least 1 h at room temperature or overnight at 4 ° C. The cells can remain in the formalin for a longer time, but this is not recommended unless it is necessary to collect multiple plates over several days before staining. 2. Oil Red O staining: After fixing the cells, add sufficient Oil Red O to each plate or well to completely cover the cells. Gently rock at room temperature for 1 h, and then rinse at least 3 times with distilled water. At this point, the cells can be imaged using

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a microscope/camera (see Note 9). If desired, the amount of Oil Red O staining is quantified by removing the distilled water and adding 100% isopropanol. Add 0.375 mL of 100% isopropanol/cm2 (6-well plate is 9.5 cm2/well; 12-well plate is 4.0 cm2/well, 24-well plate is 2 cm2/well). As a blank, add an equal amount of isopropanol to an empty well. Elution of the Oil Red O should be complete within 10–15 min at room temperature. Determine the absorbance of the mixture at 500 nm (or 540 nm if using selected UV filters) and subtract the blank measurement from the sample absorbance.

4

Notes 1. If fungal contamination is a concern, solutions of antibiotics combined with Amphotericin as an antimycotic agent are also commercially available. 2. Dexamethasone is available in a water-soluble and waterinsoluble form from Sigma. For the water-insoluble form (Sigma), the 1 mM stock is made by adding 0.39 mg dexamethasone to 1 mL 100% ethanol. For the water-soluble form (Sigma), there is 69 mg dexamethasone/100 mg total weight. To obtain a 1 mM stock solution using the water-soluble form, add 6.0 mg dexamethasone mixture to 1 mL distilled water. 3. This amount of collagenase solution/gram tissue is a good starting point for determining how much collagenase solution is needed to efficiently digest the adipose tissue within 1 h. Another approach is to add a volume of collagenase solution that is equal to the volume of minced adipose tissue. 4. The tissue can be filtered at this step to remove any remaining undigested pieces. Use a 100 μm filter (BD-Falcon) that is available for filtering small volumes. However, if it is desirable to collect the mature adipocytes at a later step, use a nylon filter (sterile) screen with a pore size of 250 μm. 5. As has been reported [16], it is also possible to separate small and large mature adipocytes for further analysis by using a 75 μm pore size filter at this step. Rather than aspirating the oil and yellow layer containing the adipocytes, carefully remove the oil and yellow layer and filter (BD-Falcon has filters of various pore sizes that work well for small volumes). The flow-through will contain the smaller adipocytes and the solution remaining on top of the filter will contain adipocytes greater than 75 μm in size. 6. ASCs obtained from young mice (3–6 weeks of age) or lean mice will typically differentiate more readily than ASCs obtained from older mice or obese animals. The yield of SVF

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cells or ASCs may also be affected by the age or body weight of the animal. With experience, this can be factored into the number of fat pads needed to obtain the desired number of cells for each application. In addition, ASCs from inguinal adipose tissue will differentiate more readily than ASCs from gonadal adipose tissue. 7. ASCs differentiated toward osteoblasts can be probed for alkaline phosphatase as an early (4–6 days post-induction) indicator of osteoblast differentiation. Calcium staining (Alizarin red or Von Kossa) can be used to visualize mineralized plaques, a marker of mature osteoblast differentiation (14–28 days). 8. Differentiation of the ASCs occurs in a higher percentage of the total number of cells when the cells are plated in smaller wells (i.e., the cell will better differentiate in a 12-well plate than a 6-well plate) although both are originally seeded at the same concentration of cells per cm2. 9. Oil Red O stain will initially be specific to neutral lipid droplets but may disperse over time. For best results, add enough distilled water to cover the plate/wells after third rinse and image immediately.

Acknowledgments This work used the Cell Biology and Bioimaging Core facility at Pennington Biomedical Research Center that is supported in part by COBRE (NIH P20 GM135002) and NORC (NIH 5P30DK072476-17) center grants from the National Institutes of Health. Dr. Scott is supported by NIH T32AT004094. References 1. Cawthorn WP, Scheller EL, MacDougald OA (2012) Adipose tissue stem cells meet preadipocyte commitment: going back to the future. J Lipid Res 53(2):227–246. https://doi.org/ 10.1194/jlr.R021089 2. Bourin P, Bunnell BA, Casteilla L et al (2013) Stromal cells from the adipose tissue-derived stromal vascular fraction and culture expanded adipose tissue-derived stromal/stem cells: a joint statement of the International Federation for Adipose Therapeutics and Science (IFATS) and the International Society for Cellular Therapy (ISCT). Cytotherapy 15(6):641–648. https://doi.org/10.1016/j.jcyt.2013.02.006 3. Kelly KA, Tanaka S, Baron R et al (1998) Murine bone marrow stromally derived BMS2 adipocytes support differentiation and function

of osteoclast-like cells in vitro. Endocrinology 139(4):2092–2101 4. Thompson DL, Lum KD, Nygaard SC et al (1998) The derivation and characterization of stromal cell lines from the bone marrow of p53-/- mice: new insights into osteoblast and adipocyte differentiation. J Bone Miner Res 13(2):195–204 5. Safford KM, Hicok KC, Safford SD et al (2002) Neurogenic differentiation of murine and human adipose-derived stromal cells. Biochem Biophys Res Commun 294(2):371–379 6. Zheng B, Cao B, Li G et al (2006) Mouse adipose-derived stem cells undergo multilineage differentiation in vitro but primarily osteogenic and chondrogenic differentiation in vivo. Tissue Eng 12(7):1891–1901. https://doi. org/10.1089/ten.2006.12.1891

Isolating Murine Adipose-Derived Stromal/Stem Cells 7. Wu J, Bostrom P, Sparks LM et al (2012) Beige adipocytes are a distinct type of thermogenic fat cell in mouse and human. Cell 150(2): 366–376. https://doi.org/10.1016/j.cell. 2012.05.016 8. Ohno H, Shinoda K, Spiegelman BM et al (2012) PPARgamma agonists induce a whiteto-brown fat conversion through stabilization of PRDM16 protein. Cell Metab 15(3): 395–404. https://doi.org/10.1016/j.cmet. 2012.01.019 9. Reilly SM, Saltiel AR (2017) Adapting to obesity with adipose tissue inflammation. Nat Rev Endocrinol 13(11):633–643. https://doi. org/10.1038/nrendo.2017.90 10. Ghaben AL, Scherer PE (2019) Adipogenesis and metabolic health. Nat Rev Mol Cell Biol 20:242. https://doi.org/10.1038/s41580018-0093-z 11. Bagchi DP, MacDougald OA (2019) Identification and dissection of diverse mouse adipose depots. J Vis Exp 149. https://doi.org/10. 3791/59499 12. Langenbach F, Handschel J (2013) Effects of dexamethasone, ascorbic acid and beta-

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glycerophosphate on the osteogenic differentiation of stem cells in vitro. Stem Cell Res Ther 4(5):117. https://doi.org/10.1186/scrt328 13. Huang CP, Hsu KC, Wu CP et al (2022) Osteogenic differentiation from mouse adiposederived stem cells and bone marrow stem cells. Chin J Physiol 65(1):21–29. https:// doi.org/10.4103/cjp.cjp_64_21 14. Grant R, Youm YH, Ravussin A et al (2013) Quantification of adipose tissue leukocytosis in obesity. Methods Mol Biol 1040:195–209. https://doi.org/10.1007/978-1-62703-5231_15 15. Cho KW, Morris DL, Lumeng CN (2014) Flow cytometry analyses of adipose tissue macrophages. Methods Enzymol 537:297– 314. https://doi.org/10.1016/B978-0-12411619-1.00016-1 16. Bluher M, Patti ME, Gesta S et al (2004) Intrinsic heterogeneity in adipose tissue of fat-specific insulin receptor knock-out mice is associated with differences in patterns of gene expression. J Biol Chem 279(30): 31891–31901. https://doi.org/10.1074/jbc. M404569200

Chapter 7 Isolation of Murine Adipose-Derived Stromal/Stem Cells Using an Explant Culture Method Ziyi Mei, Yuejia Li, Jeffrey M. Gimble, and Jie Li Abstract Adipose tissue provides a valuable cell source for tissue engineering, regenerative medicine, and adipose tissue biology studies. The most widely used adipose-derived stromal/stem cells (ASCs) isolation protocol involves enzymatic digestion with collagenase. However, the yield of the method often proves to be poor if not impossible for collection of sufficient stromal vascular fraction (SVF) for expansion when the sample size is small, for instance when only newborn mice are available for cell culture. Here, we describe an efficient protocol for the isolation and expansion of ASCs using explant culture as an alternative. Briefly, adipose tissue was minced after removing excess liquid. Then, the minced tissue was placed in culture dishes or flasks. The cells will migrate out of tissue and adhere to the culture surface after one or more days. Key words Murine, Adipose-derived stromal/stem cells (ASCs), Collagenase, Explant, Mesenchymal stromal/stem cells (MSCs)

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Introduction Two methods have been widely adopted to isolate stem/stromal cells from mammal tissue: enzymatic dissociation of tissues and explant culture. ASCs with similar biological properties could be acquired from either methods using adipose tissue, which has proven to be an abundant source of stromal/ stem cells [1]. The majority of methods isolating ASCs from mammalian adipose tissue use an approximately one-hour type I collagenase digestion. However, the digestion process is time consuming, expensive, infrastructure-demanding, and in most western countries, collagenase digestion and culture is a highly regulated cellular operation [2]. Compared to explant culture method, digestion techniques can be more labor-intensive and less cost-effective. Explant culture can be defined as an in vitro technique that puts small fragments of tissue on a growth surface for adherence and eventual outgrowth of cells. In particular, explant culture may be preferable when only

Jeffrey M. Gimble et al. (eds.), Adipose-Derived Stem Cells: Methods and Protocols, Methods in Molecular Biology, vol. 2783, https://doi.org/10.1007/978-1-0716-3762-3_7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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small fragments of tissue are available. The process of explant culture is more moderate for ASCs to transfer from an in vivo to an in vitro environment than using digestion culture and may simplify the process of satisfying the minimal manipulation criterion in U.S. Food and Drug Administration (FDA) guidelines. While the current protocol focuses on murine adipose tissue(s), similar isolation methods can be adopted in human or other mammalian adipose tissues as well [3–5]. In addition to traditional, manual tissue digestion, various closed systems have been designed and developed to perform the isolation process in a high quality, safe, and sterile manner. Several mechanical schemes were proposed, including vibration, prolonged centrifugation, blade cutting, emulsification, and so on [6]. One study found that directly plating the lipoaspirate can yield a similar ASC population compared to collagenase digestion methodology [7]. Research has also shown that the fluid portion of the liposuction aspirate contains a substantial amount of ASCs [8]. Shaking the lipoaspirates in ammonium-chloride-potassium lysing buffer following centrifugation generates a mesenchymal stromal/stem cell (MSC) population [9]. Vigorously washing the floating lipoaspirate can release a fraction of ASCs, which may be useful when processing large volumes of lipoaspirate [10]. These results suggest that mechanical disruption is able to release ASCs from the adipose tissue. Towards this end, a commercially available mechanical device called the “Lipogems device” is gaining acceptance using a self-contained kit to extract, manipulate, and isolate ASCs [11]. Thus, both non-enzymatic and enzymatic methods have been documented that provide techniques to isolate and extract stem/stromal cells quickly and efficiently from adipose tissue. In general, compared to enzymatic methods, explant methods require minimal use of equipment and harvest homogeneous cells that have high proliferation rates and cell viability [12]. The basic method of explant culture is to cut adipose tissue into small pieces and adhere them to the dish, so that the cells subsequently can migrate out of the tissue and adhere to the culture surface.

2

Materials

2.1

Tissue

1. Subcutaneous inguinal adipose tissue from mouse or rat (see Note 1).

2.2

Supplies

1. 60 mm diameter culture dish or 25-cm2 tissue culture (T25) flasks. 2. Autoclaved scissors and forceps. 3. Hemocytometer. 4. Freezing apparatus.

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5. Sterile plastic disposable conical centrifuge tubes. 6. Plastic disposable snap-cap centrifuge tubes: 1.5 mL. 7. Cell strainer. 2.3

Equipment

1. Phase contrast microscope. 2. Imaging software. 3. Biosafety hood. 4. CO2 incubator. 5. Centrifuge. 6. Flow cytometer.

2.4 Reagents and Buffers

1. 70% ethanol. 2. Stromal medium: Add 55 mL of 10% fetal bovine serum and 5.6 mL of 100× antibiotic (penicillin/streptomycin)/antimycotic (amphotericin) to 500 mL of α-minimal essential medium (α-MEM) or DMEM/Ham’s F-12 medium. This solution should be used within 4 weeks of its preparation. All fetal bovine serum should be pre-screened prior to purchase for its ability to support both cell proliferation and adipocyte differentiation. 3. 0.5% trypsin/EDTA solution. 4. Phosphate-buffered saline (PBS). 5. Wash solution: 99% PBS, 1% penicillin-streptomycin solution. 6. Bovine serum albumin (BSA). 7. Fluorochrome-conjugated monoclonal antibodies against stromal (CD29, CD105, CD106, and Sca1), hematopoietic (CD34 and CD45), endothelial (CD31), and related isotype control antibody.

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Methods

3.1 Isolation of the Adipose Tissue

All procedures are performed in a biosafety hood or other aseptic environment. 1. Ensure the ASC isolation procedure is initiated within 20 min of the tissue extraction, especially if a large number of animals are being euthanized for harvest on a single day. 2. Warm the PBS in water bath at 37 °C. 3. Place a bench protector down in hood. 4. Sacrifice the animal(s) using anesthesia or CO2 asphyxiation as approved by the AVMA guidelines on euthanasia.

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Fig. 1 The explant culture of the ASCs. (a) The adipose tissue fragments attached to the dish after 5–10 min. (b) ASCs outgrowth from the tissue explants after 2 days. (c) ASCs outgrowth from the tissue explants after 4 days

5. Saturate the fur with 70% ethanol and place the animal ventral side up. A dissection board may be used if desired. 6. Using sterile tissue forceps, pull up on the skin and cut the skin vertically toward the head along the midline at the abdominal level. 7. Using sterile scissors and forceps, remove the desired fat pads from the subcutaneous inguinal depots. 8. Place the tissue in a culture dish containing wash solution. 3.2 Plate the Adipose Tissue Explants

1. Wash the adipose tissue twice with PBS and absorb the excess PBS (see Note 2). 2. Transfer the adipose tissue to another dry container and mince into 1–2 mm3 pieces using sterile scissors under aseptic conditions (see Note 3). 3. Place the tissue explants in a culture dish or flask with a 5 mm space between the adjacent fragments (see Note 4 and Fig. 1). 4. After adhesion (5–10 min, depends on the liquid content in the minced tissue), gently add fresh stromal media to the petridish/flask without disturbing the explants (see Note 5). 5. Change the media every 2–3 days (see Note 6). 6. Remove the tissue explants at day 5 or day 6 (see Note 7). 7. At 80–90% confluence, the cells are either passaged, stored as frozen stocks at passage zero (P0), or differentiated using an experimental protocol (see Note 8).

3.3 Characterization of the Cells

1. For harvesting viable ASCs, wash the flask or plate twice with PBS containing 2% FBS. 2. Add 0.5% trypsin/EDTA solution to the flask or plate and incubate for 5–10 min in an incubator. 3. After 90% of the cells have detached when observed under the microscope, add an equal volume of media to terminate the reaction.

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4. Wash the cells with PBS containing 2% FBS three times. 5. Transfer the medium containing the suspended cells to a sterile 15 mL tube. 6. Centrifuge at 300 × g for 5 min, resuspend in stromal media and count the cells. 7. Aliquot between 2.5 and 5 × 105 cells per conical tube. 8. Add the appropriate primary antibodies and incubate for 30 min in the dark at room temperature according to the manufacturer’s instructions. 9. Wash the cells with 1 mL PBS with 1% BSA and pellet the cells at 300 × g for 3 min at room temperature. Repeat this step three times. 10. Resuspend the final pellet in 500 μL PBS and filter it with cell strainer prior to flow cytometry analysis.

4

Notes 1. The age, sex, body mass index (BMI), obesity, and the subsequent chronic outcomes, cell culture conditions such as medium composition, and culture surfaces serve as impactful factors influencing the proliferation, differentiation, and paracrine secretion potential of ASCs [13]. 2. In order to attach the explants to the dish quicker, try to remove as much liquid as possible from the adipose tissue following the PBS wash. 3. We prefer smaller tissue fragments than other groups [3] because central necrosis will happen due to low nutrient and oxygen exchange. 4. For reasons that are not quite clear, seeding explants at too high of a density in the culture dish or flask will inhibit the growth of cells from the explant. Based on our experience, two fat pads from an adult mouse for five to six 60-mm diameter culture dishes is optimal. As reviewed in the literature, numerous substrates such as collagen, fibronectin, and basement membrane proteins provide a physical scaffold for cellular components and releasing growth factors that enhance ASCs growth [14]. 5. It is critical that the tissue fragments tightly adhere to the culture dish or plate to obtain ASCs consistently and efficiently. As described by other independent studies and investigators, ASCs can only migrate from the adherent tissue fragments and seed as adherent cells from explants that are on the plastic but not from floating tissue fragments [15]. 6. Cell outgrowth from the explants usually occurs during day 1–3. If the outgrowth of cells is not observed after 4–5 days, the explant culture failed and should be tried again.

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7. After about 4–7 days, adipogenic differentiation of the ASCs will occur around the explants. Certain adipogenic factors secreted by the explants such as miR-450a-5p [18] and matrix metalloproteinases [19] from exosomes induce the differentiation process [16, 17]. 8. The explant culture method yields about 2.4 × 104 cells per mg tissue after 7 days in culture [1]. References 1. Jing W, Xiao J, Xiong Z et al (2011) Explant culture: an efficient method to isolate adiposederived stromal cells for tissue engineering. Artif Organs 35:105–112 2. Raposio E, Ciliberti R (2017) Clinical use of adipose-derived stem cells: European legislative issues. Ann Med Surg 24:61–64 3. Priya N, Sarcar S, Majumdar AS, SundarRaj S (2014) Explant culture: a simple, reproducible, efficient and economic technique for isolation of mesenchymal stromal cells from human adipose tissue and lipoaspirate. J Tissue Eng Regen Med 8:706–716 4. Zeng G, Lai K, Li J et al (2013) A rapid and efficient method for primary culture of human adipose-derived stem cells. Organogenesis 9: 287–295 5. Gittel C, Brehm W, Burk J, Juelke H, Staszyk C, Ribitsch I (2013) Isolation of equine multipotent mesenchymal stromal cells by enzymatic tissue digestion or explant technique: comparison of cellular properties. BMC Vet Res 9:221 6. Kim BS, Chen SH, Vasella M et al (2022) In vivo evaluation of mechanically processed stromal vascular fraction in a chamber vascularized by an arteriovenous shunt. Pharmaceutics 14: 417 7. Busser H, De Bruyn C, Urbain F et al (2014) Isolation of adipose-derived stromal cells without enzymatic treatment: expansion, phenotypical, and functional characterization. Stem Cells Dev 23:2390–2400 8. Yoshimura K, Shigeura T, Matsumoto D et al (2006) Characterization of freshly isolated and cultured cells derived from the fatty and fluid portions of liposuction aspirates. J Cell Physiol 208:64–76 9. Baptista LS, do Amaral RJ, Carias RB, Aniceto M, Claudio-da-Silva C, Borojevic R (2009) An alternative method for the isolation of mesenchymal stromal cells derived from lipoaspirate samples. Cytotherapy 11:706–715

10. Shah FS, Wu X, Dietrich M, Rood J, Gimble JM (2013) A non-enzymatic method for isolating human adipose tissue-derived stromal stem cells. Cytotherapy 15:979–985 11. Bianchi F, Maioli M, Leonardi E et al (2013) A new nonenzymatic method and device to obtain a fat tissue derivative highly enriched in pericyte-like elements by mild mechanical forces from human lipoaspirates. Cell Transplant 22:2063–2077 12. Mushahary D, Spittler A, Kasper C, Weber V, Charwat V (2018) Isolation, cultivation, and characterization of human mesenchymal stem cells. Cytometry A 93:19–31 13. Caˆmara DAD, Shibli JA, Mu¨ller EA et al (2020) Adipose tissue-derived stem cells: the biologic basis and future directions for tissue engineering. Materials (Basel) 13:3210 14. Hendijani F (2017) Explant culture: an advantageous method for isolation of mesenchymal stem cells from human tissues. Cell Prolif 50: e12334 15. Mori Y, Ohshimo J, Shimazu T et al (2015) Improved explant method to isolate umbilical cord-derived mesenchymal stem cells and their immunosuppressive properties. Tissue Eng Part C Methods 21:367–372 16. Li J, Qiao X, Yu M et al (2014) Secretory factors from rat adipose tissue explants promote adipogenesis and angiogenesis. Artif Organs 38:E33–E45 17. Dai M, Zhang Y, Yu M, Tian W (2016) Therapeutic applications of conditioned medium from adipose tissue. Cell Prolif 49:561–567 18. Zhang Y, Yu M, Dai M et al (2017) miR-450a5p within rat adipose tissue exosome-like vesicles promotes adipogenic differentiation by targeting WISP2. J Cell Sci 130:1158–1168 19. Dai M, Yu M, Zhang Y, Tian W (2017) Exosome-like vesicles derived from adipose tissue provide biochemical cues for adipose tissue regeneration. Tissue Eng Part A 23:1221– 1230

Chapter 8 Canine Adult Adipose Tissue-Derived Multipotent Stromal Cell Isolation, Characterization, and Differentiation Takashi Taguchi and Mandi J. Lopez Abstract Adult mesenchymal stromal/stem cells (MSCs) are a standard component of de novo tissue generation to treat and study injury, disease, and degeneration. Canine patients constitute a major component of veterinary practice, and dogs share numerous pathologic conditions with humans. The relative abundance of adipose-derived stromal/stem cells (ASCs) in various canine adipose tissue depots is well described. Refined isolation, characterization, and differentiation techniques contribute to the collective knowledge of ASC phenotypes and subpopulations for specific tissue targets. Continued efforts to advance the knowledge of canine ASC behavior in vivo are critical to harnessing the full potential of primary cell isolates. This chapter contains a description of techniques to isolate, characterize, and differentiate canine ASCs. Key words Dog, Fat, Tissue, Bioengineering, Regenerative medicine

1

Introduction Adipose tissue is an established source of canine adult multipotent stromal cells. Current information confirms the expansion potential, plasticity, and revitalization from cryopreservation of canine adult adipose derived stromal/stem cells (ASCs) [1–3]. Comparisons to canine MSCs from other tissues confirm similar cell expansion rates and comparable plasticity [1, 2, 4]. Additionally, canine ASCs are capable of dedifferentiation and transdifferentiation into ectodermal and endodermal cell lineages [5, 6]. Recent studies confirm immunomodulatory properties, which support their potential for engraftment without destructive immune system activation [7, 8]. Collectively, canine adipose tissue has tremendous potential as a viable and accessible tissue source of ASCs intended for both therapeutic and investigative purposes [9–11]. Undifferentiated cells from different tissue harvest sites vary in capacity to produce tissue-specific extracellular matrix [12]. Differences are attributed to epigenetic influences as well as cell

Jeffrey M. Gimble et al. (eds.), Adipose-Derived Stem Cells: Methods and Protocols, Methods in Molecular Biology, vol. 2783, https://doi.org/10.1007/978-1-0716-3762-3_8, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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phenotype and maturity, among other factors [13]. Primary cell isolates contain heterogeneous cell populations and vary widely in in vitro behaviors [14, 15]. The minimum criteria for MSC identity include protein and genetic confirmation of differentiation, histochemical staining, surface marker expression, and plastic affinity [16, 17]. Standardization of isolation, identification, and culture methods are crucial to developing effective bioengineering protocols. This chapter includes current protocols for canine ASC isolation, identification, 2- and 3-D culture, and differentiation into distinct tissue lineages, including ligament neotissue.

2

Materials

2.1 Adipose Tissue Collection and ASC Isolation

1. Scalpel handles (#3, 4) and scalpel blades (#10, 20). 2. Forceps (Brown-Adson). 3. 70% ethyl alcohol (EtOH). 4. 50 mL centrifuge tubes. 5. Petri dishes. 6. Phosphate-buffered saline (PBS). 7. 1% bovine serum albumin (BSA). Dissolved in double-distilled water. Filter sterilize with a 0.22 μm filter. Warm to 37 °C. 8. 0.1% Collagenase: Collagenase type-1 and 1% BSA dissolved in double-distilled water. Filter sterilize with a 0.22 μm filter. Warm to 37 °C. 9. Red cell lysis buffer: 155 mM NH4Cl, 10 mM KHCO3, 0.1 mM EDTA, pH 7.3. 10. Complete stromal medium: Dulbecco’s modified Eagle’s medium (DMEM)-Ham’s F12 medium supplemented with 1% antibiotic/antimycotic solution, 20% characterized fetal bovine serum (FBS; see Note 1). 11. Trypan blue.

2.2 Cryopreservation of ASCs

1. Cryopreservation medium: 80% FBS, 10% DMEM/F-12, 10% dimethylsulfoxide (DMSO). 2. Mr. Frosty™ or CoolCell® freezing container (see Note 2). 3. Cryovials.

2.3

Cell Culture

1. Complete stromal medium: DMEM-Ham’s F12 medium supplemented with 1% antibiotic/antimycotic solution, 20% characterized FBS. 2. Tissue culture flasks (T-25 or T-75).

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Cell Doubling

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1. PBS. 2. 0.05% Trypsin (1:250): 0.05% porcine trypsin in Hank’s balanced salt solution (HBSS) with 0.2 g/L ethylenediaminetetraacetic acid (EDTA). Aliquots can be stored in microtubes at -20 °C. 3. Trypan blue. 4. Hemocytometer. 5. Complete stromal medium: DMEM-Ham’s F12 medium supplemented with 1% antibiotic/antimycotic solution and 20% characterized FBS. 6. 12-well culture plates.

2.5 Adipogenic Differentiation

1. Complete stromal medium: DMEM-Ham’s F12 medium supplemented with 1% antibiotic/antimycotic solution, 20% characterized FBS. 2. Adipogenic induction medium: DMEM-Ham’s F-12 medium supplemented with 3% FBS, 1% antibiotic/antimycotic solution, 33 μM biotin, 17 μM pantothenate, 1 μM dexamethasone, 100 μM indomethacin, 1 μM insulin, 0.5 mM isobutylmethylxanthine (IBMX), and 5 μM rosiglitazone (TZD). 3. 4% formalin in PBS. 4. 0.3% Oil red O in isopropanol. Filter solution with a 0.22 μm syringe filter and store at room temperature protected from light. 5. Double-distilled water.

2.6 Osteogenic Differentiation

1. Complete stromal medium: DMEM-Ham’s F12 medium supplemented with 1% antibiotic/antimycotic solution, 20% characterized FBS. 2. Osteogenic induction medium: DMEM-Ham’s F12 supplemented with 10% FBS, 1% antibiotic/antimycotic solution, 10 nM dexamethasone, 10 mM β-glycerophosphate, 50 μg/ mL sodium 2-phosphate ascorbate. 3. 70% EtOH. 4. 150 mM NaCl in double-distilled water. 5. 2% Alizarin red solution. Dissolve alizarin red (2 g/100 mL) in double-distilled water. Adjust pH to 4.1–4.3 with dilute NaOH. Filter solution through 0.2-μm syringe filter. The solution can be stored at room temperature protected from light.

2.7 Chondrogenic Differentiation

1. Complete stromal medium: DMEM-Ham’s F12 medium supplemented with 1% antibiotic/antimycotic solution, 20% characterized FBS.

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2. Chondrogenic induction medium: DMEM-Ham’s F12 medium supplemented with 3% FBS, 1% antibiotic/antimycotic solution, 50 μg/mL ascorbate phosphate, 100 nM dexamethasone, 1% insulin-transferrin-selenium (ITS), 10 ng/mL recombinant human transforming growth factor-β3 (rTGF-β 3), 2 mM sodium pyruvate, and 40 μg/mL proline. 3. 10% formalin. 4. Paraffin. 5. 1% Alcian blue. Dissolve alcian blue (1 g/100 mL) in 3% glacial acetic acid. Adjust pH to 2.5 using acetic acid. 6. 0.1% Nuclear fast red. Dissolve nuclear fast red (1 g/L) and aluminum sulfate (50 g/L) in distilled water. Heat with stirring until nearly boiling, then leave overnight to cool. 2.8 Tenogenic Differentiation in 2-Dimensional Culture

1. Basal medium: α-Minimum Essential Medium (MEM) supplemented with 2% FBS.

2.9 Tenogenic Differentiation in 3-Dimensional Culture

1. Bioreactor consists of a chamber and frame (Fig. 1). A frame with crossbars on each end fits within the bioreactor chamber. Each chamber has a removable cap with a centrally located port at the highest end and a port in the center of the lowest end of the chamber base.

2. Tenogenic differentiation medium: α-MEM supplemented with 2% FBS, 1000 ng/mL bone morphogenetic protein (BMP)-12.

Fig. 1 The custom perfusion bioreactor system consisted of two bioreactor chambers, a medium reservoir with a port for gas exchange, a peristaltic pump and an external computer to control medium flow direction and rate (a). Each bioreactor chamber (b) has a removable internal frame (c) to accommodate templates. The templates are constructed from #2-0 PDS II®, configured as a figure 8 (d) and wrapped in COLI secured with a polyglactin 910 finger trap (e). Templates are tensioned and secured to each frame (f). A canine ASC/COLI construct after 21 days of culture in ligamentogenic medium (g). (The image is reprinted from Taguchi et al. [20], with permission from The Sheridan Press)

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2. 10-mL medium reservoir for gas exchange. 3. Peristaltic pump (ISM404b; Ismatec) connected to the bioreactor and medium reservoir via tubing (Tygon; Saint-Gobain Performance Plastics; inner diameter, 4.8 mm). 4. Peristaltic pump fluid flow controller software program (LabView). 5. Polydioxanone sulfate suture (#2-0). 6. Bovine collagen type I (COLI) sponge. 7. Polyglactin 910 suture (#2-0). 8. PBS containing 10 mg/mL fibronectin. 9. Tenogenic medium: Low-glucose DMEM supplemented with 5% FBS, 1% antibiotic/antimycotic solution, 1 ng/mL basic fibroblast growth factor (bFGF), 5 ng/mL TGF-β, and 10 ng/ mL platelet-derived growth factor (PDGF). 2.10 Neurogenic Differentiation

1. Predifferentiation neural induction medium (mitogenically stimulated; STIM1): DMEM Glutamax supplemented with 20 ng/mL epidermal growth factor (EGF), 20 ng/mL bFGF, 1× B27 supplement. 2. Neural induction medium a (NIMa): DMEM Glutamax supplemented with 2× B27 supplement, 1× N2 supplement, 1% FBS, 0.1% penicillin/streptomycin, and 10 nM retinoic acid (RA).

2.11 InsulinProducing Cell Differentiation

1. Serum Free Medium (SFM)-1: DMEM supplemented with 1% ITS, 1% BSA, 4 nM Activin A, and 1 nM sodium butyrate. 2. SFM-1.2: DMEM supplemented with 1% BSA, 4 nM Chir99021, and 1 nM sodium butyrate. 3. SFM-2.3.1: DMEM supplemented with 1% BSA, 1% ITS, 0.3 mM taurine, 20 ng/mL bFGF, and 50 ng/mL EGF. 4. SFM-2.3.2: DMEM supplemented with 1% BSA, 1% ITS, 0.3 mM taurine, 2 μM RA, 10 mM nicotinamide, 25 μL N-[N-(3, 5-difluorophenacetyl)-l-alanyl]-s-phenylglycinetbutyl ester (DAPT), 1 μM dorsomorphin, and 10 μM SB431542. 5. 2% alginate solution. 6. 100 mM CaCl2. 7. Krebs–Ringer–Hepes (KRH) with CaCl2 buffer. 8. 30% pluronic F127. 9. SFM-3.2: DMEM supplemented 1% BSA, 1% ITS, 3 mM taurine, 1 mM nicotinamide, 100 nM glucagon-like peptide (GLP)-1, 1% non-essential amino acid, 10 μM SB431542, 10 μL forskolin, and 10 μM LY294002.

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2.12 Papain Digestion

1. 25 mg/mL Papain in double-distilled water. 2. Solution A: 10 mM di-sodium EDTA and 0.4 M sodium acetate in double-distilled water. 3. Solution B: 200 mM L-cysteine in double-distilled water. 4. Solution C: Add 1 mL of solution B to 9 mL of solution A just before use (1:10 dilution). 5. Solution D: Add 200 μL of papain suspension to 10 mL solution C.

2.13 Hydroxyproline Assay

1. Double-distilled water. 2. Clear, flat-bottom 96-well ELISA plate. 3. 12 N HCl. Dilute 1:2 in double-distilled water for 6 N working solution. 4. 17% NaCl dissolved in double-distilled water. 5. Acetate/citrate buffer: 120 g sodium acetate trihydrate, 12 mL acetic acid, 50 g citric acid monohydrate, and 34 g NaOH in 1 L double-distilled water (pH 6.0). Store at 4 °C. 6. Oxidant solution: 0.178 g chloramine-T in 15 mL of isopropanol and 10 mL of double-distilled water; add 25 mL acetate/ citrate buffer. Make fresh (see Note 3). 7. Ehrlich’s reagent: 1 g dimethylaminobenzaldehyde in 20 mL isopropanol; combine with 6.6 mL perchloric acid and 15.6 mL double-distilled water. Make fresh (see Note 3). 8. 1 mg/mL Trans-4-hydroxy-L-proline in double-distilled water. Store at 4 °C.

2.14 PicoGreen Double Stranded DNA Assay

1. 20× TE buffer: 200 mM Tris–HCl, 20 mM EDTA, pH 7.5. Dilute 1:20 in sterile, distilled, DNase-free water for a 1× TE working solution. 2. PicoGreen® dsDNA reagent: Dilute 1:200 in 1× TE. Store at 4 °C protected from light. Make fresh just before use. 3. Black, flat-bottom 96-well microtiter plate. 4. Bacteriophage lambda DNA stock: 100 μg/mL in TE. Dilute 1:50 in 1× TE to 2 μg/mL. Make serial dilutions from 1 ng/ mL to 1 μg/mL for the standard curve. Be sure to include a blank sample of TE buffer alone. Store at 4 °C protected from light. Make fresh just before use.

2.15 Lowry Total Protein Assay

1. Biuret reagent: 0.5 mL 1% cupric sulfate, 0.5 mL 2% sodium potassium tartrate, and 50 mL 2% sodium carbonate in 0.1 N NaOH. 2. 1 N Folin & Ciocalteu’s reagent. 3. 1 mg/mL BSA in distilled water.

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1. 1,9-dimethylmethylene blue (DMMB) assay buffer: Dissolve 21 mg of DMMB in 5 mL of absolute EtOH. In a separate beaker, dissolve 2 g of sodium formate in 980 mL of distilled water. Adjust the pH to 3.5 with formic acid. Combine the solutions and bring the total volume to 1 L with doubledistilled water. The DMMB assay buffer may be stored at 4 °C. 2. Shark chondroitin-6-sulfate C: 1 mg/mL in double-distilled water. Prepare standards, usually 100, 50, 40, 30, 20, 10, 5, and 0 μg/mL. Standards can be stored at -25 °C.

2.17 Transmission Electron Microscopy (TEM) Sample Preparation

1. PBS. 2. T75 culture flasks. 3. 2% paraformaldehyde. 4. 0.05% Trypsin—0.5 mM EDTA. Store long-term at -20 °C. 5. 5% sucrose in 0.1 M sodium cacodylate (CAC) buffer. 6. 1.25% Glutaraldehyde in CAC buffer (pH 7.4). 7. 1% Osmium tetroxide in CAC buffer. 8. 70%, 80%, 95%, and 100% ethanol/distilled water solutions. 9. EPONtm epoxy resin.

2.18 Immunocytochemistry

1. Complete stromal medium: DMEM/F-12 supplemented with 1% antibiotic/antimycotic solution, 20% FBS. 2. 4-well chamber slides. 3. 10% formalin. 4. TBS buffer: 100 mM Tris–HCl, 138 mM NaCl, 27 mM KCl, pH 7.4. 5. SDS/TBS antigen retrieval buffer: 1% SDS in TBS buffer. 6. Goat serum (1:10 in TBS buffer). 7. Labeled antibodies. 8. 1 mg/L 4′,6-diamidino-2-phenylindole (DAPI) solution.

2.19 Protein Isolation for Western Blot

1. T75 culture flasks. 2. PBS. 3. RIPA buffer: 150 mM NaCl, 1.0% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 50 mM Tris–HCl, and 1 mM protease inhibitor. 4. Cell lifter. 5. Pierce™ bicinchoninic acid (BCA) protein assay kit. 6. 2 mg/mL BSA in 0.9% saline and 0.05% sodium azide.

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2.20 Relative Cell Number Quantification in 3-D Construct

1. PBS. 2. Resazurin reduction solution. 3. Black, clear, flat bottom 96-well plate. 4. Microplate reader.

2.21 Peripheral Blood Mononuclear Cell (PBMC) Isolation

1. Histopaque 1.111: 33.73 mL water (cell culture grade) and 500 mL of 1.119 g/mL Histopaque for a final specific gravity of 1.111. 2. Cell culture grade water. 3. 1.077 g/mL Ficoll-Paque. 4. Peripheral blood from an allogeneic donor in a vacutainer heparin tube. 5. Tyrode’s/HEPES (TH) buffer: 12 mM NaHCO3, 138 mM NaCl, 2.9 mM KCl, 10 mM HEPES, and 1 M EDTA. 6. PBS. 7. Basal medium: DMEM/low-glucose supplemented with 10% heat-inactivated FBS and 1% penicillin/streptomycin.

2.22 Modified Leukocyte Reaction (MLR)

1. 500 μg/mL Concanavalin A in cell culture grade water. 2. Bromodeoxyuridine (BrdU) mixture: 7.75 μL of BrdU in 250 μL of basal medium. 3. APC BrdU Flow Kits (BD Biosciences). 4. Anti-canine CD3 antibody conjugated to Alexa Flour®488 (clone CA17.2A12). 5. Flow cytometer.

3

Methods Characterization of ASC isolates is critical to assess the potential for targeted application. In vitro expansion rates determine the culture time required to produce specific cell quantities. Confirmation of multilineage differentiation is one of the cornerstones of multipotent stromal cell evaluation. Compositional analysis of tissuespecific protein levels normalized to DNA content helps quantify lineage differentiation capacities among cell isolates. Determination of the presence and absence of cell surface proteins is necessary to distinguish among MSC phenotypes. Cell ultrastructure and immunomodulation capabilities are additional measures of cell capabilities. Examples of standard approaches to these contemporary measures are provided as a basic platform that can be customized according to specific interests.

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3.1 Adipose Tissue Harvest and ASC Isolation [1, 2]

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1. Harvest the adipose tissue (subcutaneous, intra-articular, perigonadal, etc.) using aseptic technique. 2. Place the tissue in a preweighed, sterile petri dish to determine the tissue weight. 3. Rinse the tissue with sterile PBS. 4. Mince the tissue using a sterile scalpel and blade until the tissue is no longer fibrous. 5. Place the tissue in a 50 mL tube with an equal volume of prewarmed PBS and agitate vigorously for 30 s. 6. Allow the sample to separate into phases for 3 min. 7. Remove the infranatant (lower, aqueous phase, see Note 4) and any solid pieces at the bottom of the tube. 8. Wash the remaining tissue with PBS. Specifically, suspend the tissue in PBS, agitate for 45 s, and remove the infranatant after the adipose tissue floats to the top. Continue this process until the infranatant is clear. 9. After removing the last infranatant wash, add a 0.1% collagenase type I solution equal to the adipose tissue volume and agitate at about 75 rpm for 90 min at 37 °C or until the solution becomes homogeneous. Alternatively, maintain the solution at 37 °C for 90 min, vortexing every 10–15 min. 10. Filter the mixture with a 100 μm cell strainer, and then centrifuge (260 × g, 5 min) the solution to form the stromal vascular fraction (SVF) pellet. 11. Carefully remove the supernatant, consisting of lipids, adipocytes, and collagenase, without disturbing the SVF pellet. It is okay to leave some fluid to avoid disrupting the pellet. 12. Resuspend the cells in an equal volume of 1% BSA (see Note 5). 13. Centrifuge (260 × g, 5 min), and remove the supernatant. When aspirating, the tip of the pipette should remain at the top of the fluid to avoid disturbing the pellet. 14. Resuspend the pellet in 1 mL of complete stromal medium. 15. Centrifuge (260 × g, 5 min). 16. Carefully aspirate and discard the supernatant. 17. Resuspend the pellet in an equal volume of red lysis buffer and maintain it at room temperature for 5 min. 18. Centrifuge the cell suspension (260 × g, 5 min). 19. Resuspend the pellet in complete stromal medium. 20. Remove 10 μL and combine it with an equal volume of trypan blue.

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21. Quantify the number of cells/mL with a hemocytometer (see Note 6) [18]. 22. Plate the cells in the appropriate volume of complete stromal medium for the intended cell concentration. 3.2 Cryopreservation of ASCs

1. Centrifuge the ASC suspension of known concentration (260 × g, 5 min) to form a cell pellet. 2. Discard the supernatant and resuspend cells in freezing medium at 1–2 × 106 cells/mL. 3. Add 1 mL aliquots to the cryovials. Gently vortex the cell suspension or agitate by pipetting to keep the cells in suspension. 4. Store the vials in a freezing container at -80° C for 24 h, then transfer the vials to liquid nitrogen for long-term storage.

3.3

Cell Culture

1. Remove the medium from the flasks and rinse the attached cells with PBS. 2. Add enough trypsin-EDTA to cover the surface of the flask (3–4 mL) and incubate at 37 °C for 5 min. 3. Use an inverted light microscope to confirm that the cells are detached. Add an equal volume of the complete stromal medium to the flask. The FBS in the medium will stop the enzymatic action of the trypsin. 4. Place the cell suspension in a 15 mL tube and centrifuge (260 × g, 5 min). 5. Discard the supernatant and resuspend the cell pellet in a known volume of the complete stromal medium. 6. Remove 10 μL and combine it with an equal volume of trypan blue. 7. Determine the number of cells/mL with a hemocytometer (see Note 6). 8. Seed the cells at the desired density, typically 3–5 × 103 cells/ cm2. 9. Incubate the flasks at 37 °C in 5% CO2. 10. Change the medium every 2–3 days until the cells have reached 70–80% confluence, and then repeat the cell passage procedure.

3.4

Cell Doubling

1. Seed the cells in multi-well plates with two or three wells for each interval to be included in the analysis. For the first quantification, allow sufficient time for the cells to attach, but not double (12–18 h). Subsequent evaluations can be performed at standardized intervals according to the study design. 2. Repeat steps 1–7 in Subheading 3.3.

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3. Calculate the cell doublings (CD) and doubling time (DT) using the following formulae: CD = lnðN f =N i Þ= lnð2Þ and DT = CT=CD where CT = culture time, Nf = final cell number and Ni = initial cell number. 3.5 Adipogenic Differentiation

1. Detach the cells with trypsin-EDTA and quantify the cell number. 2. Seed the cells at the desired density, typically 3–5 × 103 cell/ cm2, and culture in the complete stromal medium under standard conditions (37 °C, 5% CO2) for 7 days with media changes every 2–3 days until the cells are 70–80% confluent. 3. Remove the medium and add the adipogenic induction medium. 4. Culture the cells in the adipogenic induction medium for 21 more days with media changes every 2–3 days. 5. At the end of the culture period, remove the adipogenic induction medium and fix the cells with 4% formalin for 20 min at room temperature. 6. Remove the formalin and rinse the cells three times with distilled water. 7. Stain the cells with Oil Red O at room temperature for 20 min. 8. Rinse the cells with distilled water three times to remove the residual dye.

3.6 Osteogenic Differentiation

1. Detach the cells with trypsin-EDTA and quantify the cell concentration. 2. Seed the cells at the desired density, typically 3–5 × 103 cell/ cm2, and culture in the complete stromal medium under standard conditions (37 °C, 5% CO2) for 7 days with media changes every 2–3 days until the cells are 70–80% confluent. 3. Remove the medium and culture cells in osteogenic differentiation medium for 14 more days. Change the medium every 2–3 days. 4. At the end of the culture period, remove the osteogenic induction medium, wash with 150 mM NaCl solution three times, and fix the cells with 70% EtOH for 1 h at 4 °C. 5. Stain the cells with alizarin red at room temperature for 10 min. 6. Rinse the cells with distilled water five times to remove the residual dye.

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3.7 Chondrogenic Differentiation

1. Detach the cells with trypsin-EDTA and quantify the cell number. Aliquots of 2.5 × 105 cells are required for chondrogenic pellet culture. 2. Centrifuge (250 × g, 5 min) the cell aliquot suspended in the complete stromal medium at room temperature in a 1.5 mL microcentrifuge tube to form a pellet. 3. Remove the supernatant and add 1 mL of the chondrogenic differentiation medium to the tube and culture under standard culture conditions (37 °C, 5% CO2) with media changes every 2–3 days. 4. The tube should remain upright and the cap loose to allow gas exchange. 5. The pellet should be ready for harvest after 14–21 days of culture. 6. After the desired culture period, remove the chondrogenic differentiation medium and fix the pellet in 10% formalin overnight at 4° C. 7. Embed the pellet in paraffin for sectioning (3–5 μm) and staining with alcian blue and nuclear fast red.

3.8 Tenogenic Differentiation in 2Dimensional Culture [19]

1. Seed the cells on cultureware at a density of 1 × 104 cells/cm2 and incubate in the basal medium for 1 day. 2. Replace the medium with the tenogenic differentiation medium and culture for up to 14 days. 3. Change the medium every 2–3 days.

3.9 Tenogenic Differentiation in 3Dimensional Culture [20]

1. Prepare the core structure with a monofilament absorbable polydioxanone sulfate suture coiled into a 3-cm-long, 1-cm-wide loop consisting of 5 suture strands (Fig. 1). 2. With the ends held in position by hemostats, wrap the core loop with a COLI sponge rectangle measuring 3 × 2 × 0.3 cm3. 3. Secure the COLI sponge with a braided absorbable polyglactin 910 suture in a finger trap pattern. 4. Affix one polyglactin 910 suture tag to the lowermost bar of the bioreactor frame and the other to the uppermost bar. 5. Saturate the COLI with a PBS solution containing 10 mg/mL fibronectin. 6. Place the bioreactor frame into a bioreactor chamber and apply the top lid. 7. Add 24 mL of tenogenic medium to the bioreactor chamber through the stopcocks on the top and equilibrate with the fluid flow for approximately 60 min in an incubator (37 °C, 5% CO2, 90% humidity).

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8. Stop the fluid flow and add the ASC aliquots (3.3 × 106 cells/ cm3) through the top port of the reach bioreactor. 9. Resume the fluid flow after an hour and maintain a constant fluid flow rate (1 mL/min). The bioreactors should remain within the incubator for the culture duration. 10. During culture, the fluid flow should reverse when it reaches the lowest end of the top frame bar within a bioreactor so that the constructs remain immersed throughout the perfusion. 11. Replace the tenogenic medium every 7 days through the reservoir. 3.10 Neurogenic Differentiation [6]

1. Culture cells in the basal medium on cultureware at 1 × 106 cells/cm2 for 24 h. 2. Replace the medium with the predifferentiation STIM1 and incubate for 24 h. 3. Replace the medium with the NIMa. 4. Incubate the cells with the NIMa for 9 days (Fig. 2).

3.11 InsulinProducing Cell Differentiation [5]

1. Seed 1 × 106 ASCs onto each well of the 24-well polystyrene culture plate and incubate with the SFM-1.2 for 24 h in an incubator (37 °C, 5% CO2, 90% humidity). Cells will form detached 3D clusters. 2. Replace the medium with SFM-1.1 and incubate for 2 days by carefully aspirating the medium without disturbing clusters. 3. Differentiate the cells into pancreatic endocrine progenitors by culturing in the SFM-2.3.1 for 2 days, followed by culture with the SFM-2.3.2 for 3 days. 4. Harvest the clusters by aspirating the medium containing the clusters and resuspending the mixture in 2% alginate solution. 5. Place the alginate mixture into a polystyrene syringe connected to a 22 G needle and eject it into 100 mM CaCl2 while stirring with a magnetic bar at the lowest speed that prevents beads from coalescing. 6. Collect the alginate beads by aspirating them into a syringe. 7. Resuspend the alginate beads in KRH containing CaCl2 buffer and mix by stirring. 8. After removing the buffer by aspirating it with a syringe, add the refrigerated solution of 30% pluronic F127 to the clusters. After 3 min incubation at room temperature, carefully aspirate the supernatant from the alginate beads with a syringe. 9. Incubate the alginate beads with the clusters in the SFM-3.2 on a 24-well polystyrene culture plate for 5 days. Observe cells for differentiation into insulin-producing cells.

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Fig. 2 Expression of neural markers TUBB3 (a, c), NF-H (d, f), and GFAP (g, i) in canine ASCs exposed to differentiation medium NIMa for 9 days. Before NIMa was added, cells were grown in pre-differentiation medium STIM1 for 24 h. Nuclei were counterstained with DAPI (blue). Scale bars are 20 μm. (The image is reprinted from Mihevc et al. [6], with permission from Springer Nature)

3.12 Papain Digestion

1. Add enough solution D to the tissue samples to give about 10 mL of solution per 1 mg of the sample. 2. Incubate at 60 °C overnight. Return the samples to the incubator if they are not homogeneous after vortexing. Repeat until the samples are at least 70% homogeneous (see Note 7). 3. Store the papain-digested samples at –80 °C.

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3.13 Hydroxyproline Total Collagen Assay [21]

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1. Dilute the samples in an equal volume of 6 N HCl and incubate overnight or for 18 h at 110 °C (see Notes 8 and 9). 2. Prepare a series of 1:2 serial dilutions, typically at 0–20 μg/mL, of the trans-4-hydroxy-L-proline in distilled water for the standard curve. 3. Mix 250 μL of distilled water and 17% NaCl to each sample. 4. Add 500 μL of the oxidant solution to the samples and standards and incubate at room temperature for 5 min. 5. After incubation, add 500 μL of Ehrlich’s reagent to each sample and standard and incubate at 60 °C for 12 min. 6. Cool the samples and standards on ice for 4 min. 7. Pipet 100 μL of each sample and standard onto a microtiter plate. 8. Read the absorbance at 550 nm and determine the hydroxyproline concentration from the standard curve.

3.14 PicoGreen Double Stranded DNA Assay [22]

1. Prepare the bacteriophage lambda DNA standard curve samples in 1× TE buffer, typically 1 ng/mL–1 μg/mL. 2. Dilute the samples in the 1× TE buffer so that they are within the standard curve. 3. Add 100 μL of the samples and standards to a black, flatbottom, 96-well microtiter plate. 4. Add 100 μL of the 1× PicoGreen® dsDNA reagent and incubate for 2–5 min in the dark. 5. Read the fluorescence of each well at an excitation wavelength of 480 nm and an emission wavelength of 520 nm. Determine the DNA concentration from the standard curve.

3.15 Lowry Total Protein Assay [23]

1. Prepare a series of serial dilutions of BSA, usually 0–40 μg/ 100 μL. 2. Transfer 100 μL of each sample and standard to a flat-bottom microtiter plate. 3. Dilute the samples with distilled water so that they are within the standard curve. 4. Add 200 μL of the Biuret reagent to each well and mix thoroughly by pipetting several times. 5. Allow the plate to incubate for 10–15 min at room temperature. 6. Add 20 μL/well of the 1.0 N Folin and Ciocalteu’s reagent. Mix thoroughly with repeated pipetting. 7. Allow the color to develop for 30 min at room temperature. 8. Immediately read the plate at 650 nm absorbance.

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3.16 1,9-Dimethylmethylene Blue (DMMB) Total Sulfated Glycosaminoglycan Assay [24]

1. Prepare a series of serial dilutions of chondroitin-6-sμLfate in double-distilled water, usually 0–100 μg/mL. 2. Add 40 μL of each sample and standard to a flat-bottom microtiter plate. 3. Add 250 μL of the DMMB assay buffer to each sample and standard. 4. Immediately read the plate at 530 nm (see Note 10).

3.17 Transmission Electron Microscopy (TEM) Sample Preparation

1. Remove the culture medium and rinse the cells (minimum 1 × 104 cells) with PBS. 2. Add 3–4 mL of trypsin-EDTA (enough to cover the surface) and incubate at 37° C for 5 min. 3. Decant the solution into a 15 mL tube and centrifuge (260 × g, 5 min) at room temperature. 4. Resuspend the cells in 5 mL PBS and centrifuge (260 × g, 5 min) at room temperature. 5. Resuspend the cells in 2 mL 2% paraformaldehyde in a 5 mL tube and centrifuge (260 × g, 10 min). 6. Decant the solution. 7. Incubate the pellet in 1.25% glutaraldehyde at room temperature for 24 hrs. 8. Wash the pellet with 5% sucrose in 0.1 M CAC buffer. Be careful not to disrupt the pellet at this and succeeding steps. 9. Incubate the pellet in 1% osmium tetroxide for 2 h at room temperature. 10. Dehydrate the pellet in a series of ethanol-distilled water solutions by soaking in each concentration for 30 min (low to high) at room temperature. 11. Embed the pellet in Epon and prepare ultrathin sections for transmission electron microscopy.

3.18 Immunocytochemistry

1. Culture the cells in the complete stromal medium on 4-well chamber slides for 3 days or until about 50% confluent. 2. Remove the medium and rinse the cells with PBS. 3. Fix the cells with 10% formalin (200 μL/well) for 24 h at room temperature. 4. Remove the formalin and add the SDS/TBS antigen retrieval buffer (200 μL/well) for 5 min at room temperature. 5. Remove the buffer and incubate with goat serum (200 μL/ well) overnight at 4 °C. 6. Add the desired antibodies (e.g., CD34-PE 1:200 in TBS buffer) to the goat serum in each well and incubate for an additional 2 h at room temperature.

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7. Remove the solution from each well and wash the cells with the TBS buffer. 8. Add 200 μL/well of the DAPI solution and incubate at room temperature for 30 min in the darkness. 9. Use an inverted fluorescent microscope to view and image the cells. 3.19 Protein Isolation for Western Blot

1. Remove the medium when the cells are 80–90% confluent. 2. Place the flasks on ice and wash the cells with ice-cold PBS. 3. Remove the PBS, add the ice-cold RIPA buffer (enough to cover the flask surface), and detach the cells using the cell lifter. 4. Pour the cell suspension into a precooled 1.5 mL microfuge tube and maintain it at 4° C for 30 min, vortexing every 5–10 min. 5. Centrifuge the cell suspension (12 × 103 g, 20 min) at 4° C. 6. Aspirate the supernatant and place it into a new 1.5 mL tube. Discard the pellet. Store the protein samples at –80 °C until they are needed for analysis. 7. Prepare the BCA working reagent by mixing BCA reagent A and reagent B 50:1. 8. Prepare a series of 1:2 serial BSA dilutions in double-distilled water, usually 0–2.0 mg/mL. 9. Pipette 25 μL of each standard and sample into separate wells of a 96-well plate. 10. Dilute the samples in double-distilled water if necessary. 11. Add 200 μL of the BCA working reagent to each well and mix it well by pipetting. 12. Incubate the plate at 37° C for 30 min. 13. Allow the plate to cool to room temperature and measure the absorbance at 562 nm.

3.20 Relative Cell Number Quantification in 3-D Construct

1. Place a construct biopsy of a known volume in 100 μL of PBS with 10% resazurin reduction solution in a well of a 96-well plate for 4 h at 37 °C. 2. Collect the solution from each well and transfer it to a black 96-well plate. 3. Measure the fluorescence at an excitation wavelength of 530 nm and an emission wavelength of 590 nm. The fluorescence intensity is an indirect measurement of the relative cell quantity.

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3.21 PBMCs Isolation

1. Add 5 mL of the Histopaque 1.111 to a 50 mL conical tube. 2. Prepare a 1:6 mixture of 1.5 mL water (cell culture grade) and 9 mL of Ficoll-Paque in a 15 mL conical tube for a final specific gravity of 1.066. 3. Overlay the Ficoll-Paque-water solution (10.5 mL) on the Histopaque in the 50 mL conical tube (see Note 11). 4. Mix 2 mL of peripheral blood and 6 mL of the TH buffer mixture in a 15 mL conical tube. 5. Overlay the peripheral blood mixture on top of the FicollPaque/Histopaque layers. 6. Centrifuge the 50 mL tube (400 × g, 20 min) with no deceleration (no braking). 7. Slowly aspirate the supernatant (serum layer) using a 5–10 mL serological pipette. Leave a thin layer over the PBMC layer directly below to avoid disturbing it. 8. Aspirate the PBMCs from beneath the serum layer with a 5–10 mL serological pipette and transfer them to a new 50 mL conical tube. 9. Add 15 mL of PBS and mix well by inverting it several times. 10. Centrifuge at (400 × g, 10 min) with maximum acceleration and deceleration. 11. Decant the supernatant, resuspend the PBMCs in 30 mL of PBS, and centrifuge (400 × g, 10 min) at room temperature. 12. Decant the supernatant, resuspend the cells in 500 μL of the basal medium and maintain them on ice until use.

3.22

MLR [7]

1. Add 12 × 106 ASCs in 6 mL of the basal medium to a T25 flask and maintain on ice until irradiation. 2. Irradiate the flask with 10 Gy using a linear accelerator. 3. Prepare four treatment groups as described: (1) PBMCs; (2) PBMCs with ConA; (3) ASCs; and (4) PBMCs and ASCs with ConA. 4. Add ASCs (2 × 105 cells), PBMCs (1 × 106 cells), or both in basal medium to each well of a 24-well plate for a total volume of 750 μL (more than 3 replicates per treatment group). 5. Add 7.5 μL of 500 μg/mL ConA to each well of treatment groups 2 and 4 to give a volume ratio of 1:100 ConA: cell solution. 6. After 3 days of culture (37 °C, 5% CO2, 90% humidity), add 7.5 μL of the BrdU mixture to each well. 7. After 24 h, transfer the PBMCs from each well with a pipette into a flow cytometry tube.

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Fig. 3 Flow cytometry gating strategy in MLR. PBMCs were selected using forward scatter and side scatter, excluding small debris (left column). Within PBMCs, CD3+ T cells were selected (middle column), within which percentages of BrdU+ T cells were selected (right column). Top row: PBMCs alone; middle row: PBMCs activated by ConA (PBMC + ConA); bottom row: PBMCs activated by ConA and co-cultured with ASCs (PBMC + ConA + ASCs). (The image is reprinted from Taguchi et al. [7], with permission from Mary Ann Liebert Inc)

8. Add the anti-canine CD3 and anti-BrdU-APC antibodies to each tube at the manufacturer-recommended concentrations. 9. Divide the replicates of PBMCs stimulated with ConA into 3 analysis groups (n ≥ 3 per group): (1) BrdU only; (2) CD3 only; and (3) No antibodies (Unstained) to establish positive and negative staining (Fig. 3). 10. Analyze with the flow cytometer. The ratio of BrdU-positive cells to CD3-positive cells represents the fraction of activated T cells (Fig. 3).

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Notes 1. Complete stromal media can be made with 10% FBS instead of 20% FBS. We used 20% FBS in our initial cell culture and continued its use to remain constant throughout the experiments. 2. Mr. Frosty freezing container can be used up to five times before the isopropanol needs to be changed. 3. The oxidant solution and Ehrlich’s reagent can be used up to 2 days after initial use if stored at 4 °C. 4. We use a small gauge, 5-inch needle to remove the infranatant after each wash step. This makes it easy to remove the liquid without losing some of the sample. 5. Transferring the solution to a new centrifuge tube helps to reduce the amount of free lipids in the sample. 6. Mix 10 μL of cell suspension with an equal volume of trypan blue. Count both viable (unstained with trypan blue) and non-viable (stained with trypan blue) cells under a microscope using a hemocytometer. Obtain cell concentration based on the appropriate formula [18], and multiply by dilution factor (two times dilution) to compute the concentration of cell suspension. 7. If pieces of tissue still remain after repeated incubation at 60 ° C, centrifuge the sample and collect the supernatant. 8. Use of screw cap tubes helps prevent the tubes from popping open when exposed to high temperatures. 9. The sample may have a charred appearance after incubating at 110 °C. Mix sample well to dissolve. 10. Read the plate immediately to prevent precipitation of the sample and inaccurate results. 11. Tilt the 50 mL conical tube while adding layers and avoid mixing them by adding different components along the wall of the tube slowly and not directly into the solution already in the tube.

Acknowledgments The authors thank previous and current Laboratory for Equine and Comparative Orthopedic Research members at Louisiana State University, School of Veterinary Medicine for sharing their expertise.

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References 1. Spencer ND, Chun R, Vidal MA, et al. In vitro expansion and differentiation of fresh and revitalized adult canine bone marrow-derived and adipose tissue-derived stromal cells. Vet J. 2012;191(2):231–239 2. Zhang N, Dietrich MA, Lopez MJ (2013) Canine intra-articular multipotent stromal cells (MSC) from adipose tissue have the highest in vitro expansion rates, multipotentiality, and MSC immunophenotypes. Vet Surg 42(2): 137–146 3. Ho YK, Loke KM, Woo JY et al (2022) Cryopreservation does not change the performance and characteristics of allogenic mesenchymal stem cells highly over-expressing a cytoplasmic therapeutic transgene for cancer treatment. Stem Cell Res Ther 13(1):519 4. Takemitsu H, Zhao D, Yamamoto I et al (2012) Comparison of bone marrow and adipose tissue-derived canine mesenchymal stem cells. BMC Vet Res 8:150 5. Dang Le Q, Rodprasert W, Kuncorojakti S et al (2022) In vitro generation of transplantable insulin-producing cells from canine adiposederived mesenchymal stem cells. Sci Rep 12(1):9127 6. Prpar Mihevc S, Grgich K et al (2020) Neural differentiation of canine mesenchymal stem cells/multipotent mesenchymal stromal cells. BMC Vet Res 16(1):282 7. Taguchi T, Borjesson DL, Osmond C et al (2019) Influence of donor’s age on immunomodulatory properties of canine adipose tissuederived mesenchymal stem cells. Stem Cells Dev 28(23):1562–1571 8. Russell KA, Chow NH, Dukoff D et al (2016) Characterization and immunomodulatory effects of canine adipose tissue- and bone marrow-derived mesenchymal stromal cells. PLoS One 11(12):e0167442 9. Ostrander EA, Galibert F, Patterson DF (2000) Canine genetics comes of age. Trends Genet 16(3):117–124 10. Peer BA, Bhat AR, Shabir U et al (2022) Comparative evaluation of fracture healing potential of differentiated and undifferentiated Guinea pig and canine bone marrow-derived mesenchymal stem cells in a Guinea pig model. Tissue Cell 76:101768 11. Nokhbatolfoghahaei H, Bastami F, FarzadMohajeri S et al (2022) Prefabrication technique by preserving a muscular pedicle from

masseter muscle as an in vivo bioreactor for reconstruction of mandibular critical-sized bone defects in canine models. J Biomed Mater Res B Appl Biomater 110(7): 1675–1686 12. Xie L, Zhang N, Marsano A et al (2013) In vitro mesenchymal trilineage differentiation and extracellular matrix production by adipose and bone marrow derived adult equine multipotent stromal cells on a collagen scaffold. Stem Cell Rev Rep 9(6):858–872 13. Eilertsen KJ, Floyd Z, Gimble JM (2008) The epigenetics of adult (somatic) stem cells. Crit Rev Eukaryot Gene Expr 18(3):189–206 14. de Mattos CA, Alves AL, Golim MA et al (2009) Isolation and immunophenotypic characterization of mesenchymal stem cells derived from equine species adipose tissue. Vet Immunol Immunopathol 132(2–4):303–306 15. Delfi I, Wood CR, Johnson LDV et al (2020) An in vitro comparison of the neurotrophic and angiogenic activity of human and canine adipose-derived mesenchymal stem cells (MSCs): translating MSC-based therapies for spinal cord injury. Biomolecules 10(9):1301 16. Bourin P, Bunnell BA, Casteilla L et al (2013) Stromal cells from the adipose tissue-derived stromal vascular fraction and culture expanded adipose tissue-derived stromal/stem cells: a joint statement of the International Federation for Adipose Therapeutics and Science (IFATS) and the International Society for Cellular Therapy (ISCT). Cytotherapy 15(6):641–648 17. Dominici M, Le Blanc K, Mueller I et al (2006) Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8(4):315–317 18. Green MR, Sambrook J. Estimation of cell number by hemocytometry counting. Cold Spring Harb Protoc. 2019;2019(11). https:// doi.org/10.1101/pdb.prot097980 19. Shen H, Gelberman RH, Silva MJ et al (2013) BMP12 induces tenogenic differentiation of adipose-derived stromal cells. PLoS One 8(10):e77613 20. Taguchi T, Zhang N, Angibeau D et al (2021) Evaluation of canine adipose-derived multipotent stromal cell differentiation to ligamentoblasts on tensioned collagen type I templates in a custom bioreactor culture system. Am J Vet Res 82(11):924–934

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21. Stegemann H, Stalder K (1967) Determination of hydroxyproline. Clin Chim Acta 18(2):267–273 22. Labarca C, Paigen K (1980) A simple, rapid, and sensitive DNA assay procedure. Anal Biochem 102(2):344–352

23. Lowry OH, Rosebrough NJ, Farr AL et al (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193(1):265–275 24. Ratcliffe A, Billingham ME, Saed-Nejad F et al (1992) Increased release of matrix components from articular cartilage in experimental canine osteoarthritis. J Orthop Res 10(3):350–358

Chapter 9 Feline Adult Adipose Tissue-Derived Multipotent Stromal Cell Isolation and Differentiation Catherine Takawira, Wei Duan, Takashi Taguchi, and Mandi J. Lopez Abstract Cats are among the most popular household pets. However, compared to other species, there is little information specific to feline adult mesenchymal stromal/stem cells. Despite the phylogenetic distance between domesticated cats, Felis silvestris catus, and humans, they share some similar health challenges like kidney disease, asthma, and diabetes. Investigative efforts have been focused on adult adipose-derived stromal/stem cell (ASC) therapies to address feline illnesses, including de novo pancreatic tissue generation for diabetes treatment. Given the relatively small size of domestic cats, optimized cell isolation from small quantities of adipose tissue is important in the development of feline ASC-based therapies. Additionally, there are unique features of feline ASC culture conditions and characterization. This chapter contains a few of the novel aspects of feline ASC isolation, culture, preservation, and differentiation. Key words Feline, Cat, Adipose, Adult mesenchymal stromal/stem cell, Stem cell

1

Introduction Feline mesenchymal stromal/stem cells (MSCs) isolated from adipose tissue exhibit similar attributes to those from other species including in vitro culture expansion, multipotentiality, and immunophenotypes [1]. It is well established, however, that culture conditions, including those for lineage induction, vary among species [2], and there are differences between fresh and cryopreserved MSCs [3]. Hence, isolation and culture conditions as well as lineage differentiation must be customized for cell collection and characterization. Previously reported therapeutic targets for feline adipose tissue-derived multipotent stromal cell (ASC) therapies include chronic kidney disease, asthma, and diabetes, among others (Table 1) [4–13]. In general, therapeutic doses of MSCs range from 105 to 108 cells/kg. Isolation methods that yield therapeutic numbers of ASCs from small quantities of adipose tissue within an acceptable time period and low passage number are reported [2, 4,

Jeffrey M. Gimble et al. (eds.), Adipose-Derived Stem Cells: Methods and Protocols, Methods in Molecular Biology, vol. 2783, https://doi.org/10.1007/978-1-0716-3762-3_9, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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Table 1 Feline ASC trials Treatment condition Cell source

ASC dose

Administration

Treatment effect

Chronic kidney disease (CKD) [18]

Autologous

1–4 × 10

1 intrarenal injection

Modest improvement in glomerular filtration rate and mild decrease in serum creatinine

CKD4

Allogeneic

2 or 4 × 106 cryopreserved 4 × 106 ASCs from cryopreserved adipose

3 intravenous injections every 2 weeks

Low dose of ASCs was well tolerated, and serum creatinine improved. Adverse side effects associated with higher doses

Chronic allergic feline asthma [6]

Allogeneic

3.6–2.5 × 106 cryopreserved

6 intravenous infusions bimonthly

Improved airway structure evaluated with computed tomography 8 months after treatment

Feline chronic gingivostomatitis [7]

Allogeneic 20 × 106 and autologous

6

2 intravenous No positive clinical injections of response was observed 20 million fresh, in three cats and a mild allogeneic or response was noted in autologous ASCs two cats

Multiple sclerosis [8] Allogeneic

10 × 106

1 injection directly He SVF co-treated group showed into the foramen remyelination and magnum, regeneration capacity, 14 days post-MS along with reduced induction apoptosis and axonal degeneration

Inflammatory bowel Allogeneic disease [9]

2 × 106

2 intravenous injections over 2 weeks

Treatment with MSCs reduced Purina fecal and feline chronic enteropathy activity index score over 6 months

Chronic, non-responsive gingivostomatitis [10]

Autologous and allogeneic

20 × 106

2 IV transfusions, 1 month apart

Decrease in circulating globulin concentration and histological evidence of decreased CD3+ T cells and CD20+ B cells in the oral mucosa

Chronic kidney disease [11]

Autologous

3 × 106 cells were 2 intra-arterial renal Procedure deemed safe and feasible. Effect of infusion on day sought with MSC infusion on the 2 and day 14 doses between health of kidneys over 1.5 and 6 × 106 time is still unclear cells (continued)

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Table 1 (continued) Treatment condition Cell source

ASC dose

Administration

Treatment effect

Severe refractory feline chronic gingivostomatitis [12]

20 × 106

2 intravenous transfusions, 1 month apart

Allogeneic treatment delayed clinical and histologic resolution. Mechanisms of action differ between autologous and allogeneic ASCs

2 × 106

2 injections, 2 months apart

Substantial clinical improvement of ocular signs and resolution of the corneal and conjunctiva lesions

Autologous and allogeneic

Eosinophilic keratitis Allogeneic [13]

14, 15]. Diabetes mellitus is a highly prevalent feline endocrinopathy [16, 17]. De novo generation of functional pancreatic cell clusters from adipose-derived stromal/stem cells may provide implantable, viable tissue grafts that restore natural glucose regulation without the limitations and concerns of other cell origins, including allogeneic pancreatic islets harvested post-mortem [15, 16]. The information in this chapter surrounds isolation, preservation, and characterization of ASCs from feline white adipose tissue harvested during elective procedures as well as de novo generation of functional pancreatic cell clusters using a novel threestage culture process.

2

Materials

2.1 Adipose Tissue Collection and ASC Isolation from Adipose Tissue Surrounding Excised Reproductive Organs

1. 15 mL Conical tubes. 2. Phosphate-buffered saline (PBS). 3. 1% Chlorhexidine in PBS. 4. Scalpel handle (#3) and blades (#10). 5. Tissue forceps. 6. 3 cm Sterile petri dish. 7. 1 oz/30 mL Graduated, glass bottle beakers. 8. Stir bars (1 cm) and stir plate. 9. 0.3% Collagenase solution: 0.3% (375 units/mL) type I collagenase (Worthington Biochemical Corporation) and 1% bovine serum albumin (BSA) dissolved in Krebs-Ringer bicarbonate

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buffer warmed to 37 °C. Filter to sterilize with 0.22 μm filter. Make fresh the day of intended use. 10. 70 and 100 μm Cell strainers. 11. Red cell lysis buffer: 155 mM NH4Cl, 10 mM KHCO3, 0.1 mM EDTA, and pH 7.3. 12. Stromal medium: Dulbecco’s modified Eagle’s medium F-12 (DMEM-F12) supplemented with 10% fetal bovine serum (FBS) and 1% antibiotic/antimycotic (AA) solution. 13. T25 culture flasks. 2.2

Cell Expansion

1. T75 culture flasks. 2. PBS. 3. 0.05% Trypsin-ethylenediaminetetraacetic acid (EDTA). 4. 10% FBS in PBS. 5. 15 mL Conical tubes. 6. Stromal medium: DMEM-F12 supplemented with 10% FBS and 1% AA solution. 7. Trypan blue. 8. Hemocytometer.

2.3

Cryopreservation

1. T75 cell culture flasks. 2. PBS. 3. 0.05% Trypsin-EDTA. 4. 10% FBS in PBS. 5. 15 mL Conical tubes. 6. DMEM-F12. 7. Trypan blue. 8. Hemocytometer. 9. Cryopreservation medium: 80% FBS, 10% DMEM-F12, and 10% dimethyl sulfoxide (DMSO). 10. 10.1.5 mL Cryovials. 11. CoolCell® freezing container.

2.4

Flow Cytometry

1. T75 cell culture flasks. 2. PBS. 3. 0.05% Trypsin-EDTA. 4. 10% FBS in PBS. 5. 15 mL Conical tubes. 6. Trypan blue. 7. Hemocytometer.

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Table 2 Fluorescence-activated cell sorting antibodies Antibody

label

Target species

Antibody

Manufacturer

Cat No.

CD9

N/A

Cat

CD9

Serotec

MCA1345

CD29

N/A

Human

CD29

BD Biosciences

610468

CD44

N/A

Cat

CD44

VMRD

BAG40A

CD90

N/A

Human

CD90

eBiosciences

14–0909–80

CD105

PE

Human

CD105

eBiosciences

12–1057–41

IgG

FITC

Mouse

IgG

Sigma-Aldrich

F9006

8. 5 mL Round-bottom polystyrene tubes. 9. Antibodies (Table 2). 10. 10% Formalin in dH2O. 11. 1% FBS in PBS. 12. Stromal medium. 13. 6-well culture plates. 14. 1% BSA 15. 0.2% Sodium azide in PBS. 2.5

Cell Doubling

1. 96-well culture plates. 2. AlamarBlue®. 3. Stromal medium: DMEM-F12 supplemented with 10% FBS and 1% AA.

2.6 Adipogenic Differentiation (Fig. 1)

1. 24-well cell culture plates. 2. Stromal medium: DMEM-F12 supplemented with 10% FBS and 1% AA. 3. PBS. 4. Adipogenic induction medium: Minimum essential medium-α (MEM-α), 10% rabbit serum, 10% FBS, 10 nM dexamethasone, 5 μg/mL insulin, 50 μM 5, 8, 11, 14-eicosatetraynoic acid (ETYA), and 100 μM indomethacin. 5. 4% Paraformaldehyde (PFA) in PBS. 6. 0.3% Oil red O in isopropanol. Stock solution is reconstituted with 20 mL 100% isopropanol. For working solution, combine 3 parts of Oil Red O stock solution with 2 parts of water. Filter solution with a 0.22 μm syringe filter and store at room temperature protected from light. 7. Double-distilled water.

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Fig. 1 Photomicrographs of feline ASCs following culture in stromal medium (a), with calcium deposition alizarin red staining after culture in osteogenic medium (b), and with oil red O lipid staining after culture in adipogenic medium (c). Light photomicrographs of cell clusters from feline ASCs in polystyrene culture ware after 3-stage culture in stromal (d) or pancreatic induction (e) medium. Cells attached to the polystyrene surface are evident with clusters cultured in stromal medium (arrow). Dithizone staining of zinc [red, (f, g)] and viability staining (h, i) of viable (green) and non-viable (red) cells in clusters from feline ASCs after 3-stage culture in stromal (f, h) or pancreatic induction (g, i) medium. Scale bars = 200 μm (a, d, e), 50 μm (b, c), and 100 μm (f–i). (Image used with permission [5])

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1. 24-well cell culture plates. 2. Stromal medium: DMEM-F12 supplemented with 10% FBS and 1% AA. 3. PBS. 4. Osteogenic induction medium: DMEM F-12, 10% FBS, 100 nM dexamethasone, and 0.25 mM L-ascorbic acid. 5. Osteogenic maintenance medium: Osteogenic medium with 10 nM β-Glycerophosphate. 6. 70% Ethanol (EtOH) in dH2O (ice cold). 7. 2% Alizarin red in double-distilled H20 (pH 4.1–4.3). Dissolve 2 g Alzarin red in ddH2O. Adjust pH with NaOH or HCl. Filter solution with a 0.22 μm syringe filter and store at room temperature protected from light. 8. 150 mM NaCl solution.

2.8 Chondrogenic Differentiation

1. 6-well cell culture plates. 2. Stromal medium: DMEM-F12 supplemented with 10% FBS and 1% AA solution. 3. 15 mL centrifuge tubes. 4. Chondrogenesis medium: 1 g/L low glucose DMEM-F12, 3% FBS, 1% AA, 50 μg/mL ascorbate phosphate, 100 nM dexamethasone, 40 μg/mL proline, 1 mM sodium pyruvate, 1% insulin-transferrin-selenium (ITS), 10 ng/mL bone morphogenetic protein-6 (BMP-6), and 10 ng/mL transforming growth factor-β1 (TGF-β1). 5. 10% Formalin in dH2O. 6. 1% Alcian blue. Dissolve 1 g/100 mL of Alcian blue in 3% acetic acid. Adjust to pH to 2.5 using acetic acid. 7. 0.1% Nuclear fast red. Dissolve 1 g/L nuclear fast red and 50 g/L aluminum sulfate in distilled water. Heat with stirring until nearly boiling, then leave overnight to cool.

2.9 Neurogenic Differentiation (Fig. 1)

1. 24-well cell culture plates. 2. Stromal medium: DMEM-F12 supplemented with 10% FBS and 1% AA. 3. PBS. 4. Neurogenic induction medium: 4.5 g/L high glucose DMEM supplemented with 20% FBS and 1 mM β-mercaptoethanol. 5. Neurogenic maintenance medium: 4.5 g/L high glucose DMEM supplemented with 2% FBS, 2% DMSO, and 200 μM butylated hydroxyanisol. 6. 10% Formalin in dH2O. 7. 1 mg/mL Cresyl violet. Dissolve 1 mg/mL of cresyl violet in water.

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2.10 Functional Pancreatic Cell Cluster Generation

1. 24-well cell culture plates. 2. Serum free medium (SFM) 1: DMEM F-12, 1% BSA, 1 × insulin-transferrin-selenium (ITS), 1 mM sodium butyrate, 50 μM 2-mercapethanol, 1% N-2 supplement, 2% B-27 supplement, 5 μg/mL laminin, 50 ng/mL recombinant human hepatocyte growth factor (HFG), and 20 ng/mL basic fibroblast growth factor (bFGF). 3. SFM 2: DMEM F-12, 1% BSA, 1 × ITS, 0.3 mM taurine, 5 μg/ mL laminin, 20 ng/mL bFGF, 1% N-2 supplement, 2% B-27 supplement, 50 ng/mL HGF, and 1 mM nicotinamide. 4. SFM 3: DMEM F-12, 1.5% BSA, 1.5 × ITS, 3 mM taurine, 100 nM glucagon-like peptide 1, 1 mM nicotinamide, 1 × nonessential amino acids, 10 nM pentagastrin, 1% N-2 supplement, 1% B-27 supplement, 50 ng/mL HGF, 20 ng/mL bFGF, 5 μg/mL laminin, 20 ng/mL betacellulin, and 10 nM extendin-4. 5. Test tube rocker. 6. Dithizone powder. 7. 24-well laminin-coated plate. 8. 24-well ultralow attachment plate.

2.11 Colony Forming Unit Assays

1. T75 culture flasks. 2. Stromal medium: DMEM-F12 supplemented with 1% AA and 10% FBS. 3. PBS. 4. 96-well cell culture plates. 5. Adipogenic induction medium. 6. Osteogenic induction medium. 7. Osteogenic maintenance medium. 8. 4% PFA. 9. 70% EtOH. 10. 0.3% Oil red O. 11. 2% Alizarin red. 12. 0.1% Toluidine blue in dH2O. 0.1 g Toluidine blue dissolved in 100 mL distilled water.

2.12

mRNA Isolation

1. 6-well cell culture plates. 2. RNeasy® Plus Mini Kit (Qiagen). 3. Nuclease-free water. 4. Polymerase chain reaction (PCR)-grade 0.5 mL tubes. 5. 70% EtOH in nuclease-free water.

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Fig. 2 Medium insulin levels (LS mean ± SEM) following incubation of differentiated clusters with KRB buffer containing distinct glucose concentrations (a). Fold change (LS mean ± SEM) in medium insulin levels relative to 5.5 mM glucose following incubation of differentiated clusters with KRB buffer containing distinct glucose concentrations with and without theophylline (b), and intracellular insulin levels (LS mean ± SEM) following cluster incubation in KRB buffer containing 55 mM glucose with (black bar) and without (white bar) theophylline (c). Columns with distinct superscripts are significantly different between theophylline treatments within glucose concentrations, and those with different asterisk (*) numbers are significantly different among glucose concentrations within theophylline treatments ( p < 0.05). (Image used with permission [5]) 2.13 Dithizone Staining

1. Dithizone powder: Dissolve 0.2 g dithizone powder in 6 mLs of DMSO. Bring total solution volume to 40 mL by adding PBS. Filter solution. 2. DMSO. 3. PBS. 4. 15 mL Conical tube. 5. Stromal media. 6. 24-well culture plate. 7. Inverted light microscope.

2.14 GlucoseStimulated Insulin Secretion (Fig. 2)

1. Krebs-Ringer bicarbonate buffer. Suspend 9.5 g in 900 mL sterile water. Add 1.260 g sodium bicarbonate and stir until dissolved. 2. 15 mL Conical tubes. 3. Glucose. 4. 1.5 Microcentrifuge tubes. 5. Feline insulin ELISA kit (MyBioSource).

3

Methods

3.1 ASC Collection and Isolation from Minimal Adipose Tissue ( G′ in the LVE region, the viscous forces are dominant, and the sample is more fluid-like (c). (Created with BioRender.com)

Fig. 3 Typical workflow for material preparation, rheological evaluation, and gelation analysis of adiposederived hydrogels. (Created with BioRender.com)

2

Materials When handling any biologic material, ensure all instruments and experimental areas are thoroughly sterilized and cleaned during and after use of a rheometer. Before running any experiments, ensure the material(s) being evaluated are compatible with the rheometer and all proper, if any, safety protocols are being followed. Safety equipment, e.g., gloves and eye protection, should be used during all steps. 1. Micro-pipettes and tips. 2. Scoopula or spatula. 3. Diluent: Media (DMEM), phosphate buffer saline, or deionized water.

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4. Matrigel™ (VWR/Corning). 5. XGel (see Note 1). 6. 1.5 mL Eppendorf tubes. 7. Rheometer system (TA Discovery HR-2). 8. Operating software (TRIOS, version 5.1.1). 9. Analysis software (Excel or Prism, version 10.1.2). 10. Temperature control system (TA Advanced Peltier Plate). 11. Rheometer geometries (25 mm or 8 mm parallel). 12. Kimwipes. 13. 70% Ethanol.

3

Methods

3.1 Hydrogel Preparation

1. Handle and prepare all hydrogel materials following the manufacturers’ protocols. For example, the Matrigel™ solution was aliquoted into 1 mL Eppendorf tubes and stored at 4 °C until use. 2. Transport the materials on ice during experiments to ensure none prematurely crosslink.

3.2 Rheometer Setup: Preparing and Using the Rheometer Instrument (see Note 2)

1. Turn on the rheometer air supply prior to using and maintain throughout the experimental procedure (Fig. 4a) (see Note 3). 2. Turn on the power supply and Peltier plate temperature circulator (Fig. 4b). 3. Once the instrument powers on, connect the instrument to the instrument software. 4. Unscrew the safety cap that protects the spindle by holding the spindle from the top and carefully unscrewing the safety cap (Fig. 4c). 5. Attach the geometry, or probe, with the embossed line aligned with the marker on the bottom of the rheometer head (see Note 4). 6. After the geometry is loaded and lowered to ~1 cm above the stage, select “zero the gap” and the instrument will then determine the relationship between the geometry head and the stage distance. This process is comparable to zeroing a scale prior to weighing a sample. 7. Calibrate the geometry to the software. This includes geometry inertia, friction, rotational mapping, and gap compensation (see Note 5). It is important to ensure geometries are calibrated at least monthly to ensure accurate data collection (see Note 6).

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Fig. 4 Turn on procedure for rheometer, Peltier circulator, and loading of 25 mm flat geometry

8. Create the procedural setup using the instrument software for each test. This includes critical information on the sample, operator, project information or notes, where to save the file, information on the geometry being used, and the experimental procedure specifications. Figure 5 presents a typical experimental setup for a frequency sweep of a hydrogel that was cured on the stage (see Note 7). 3.3 Rheometer Setup: Sample Loading and Trimming the Gap

1. Raise the gap height and load the sample onto the stage, after completing the instrument setup, calibration, and procedural setup. Load excess sample to ensure the head is filled. 2. Set the gap height to ~5% over the final height and remove excess material as shown in Fig. 6. The excess material can be removed using Kimwipes or a spatula depending on the viscosity.

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Fig. 5 Screen captures from the rheometer software (Trios, version 5.1.1) showing sample information, geometry settings, and the procedural setup for an example frequency sweep

Fig. 6 Sample loading steps for rheometry analysis. Proper sample loading, trimming, and compression ensures proper plate coverage and accurate data collection

3. Set the gap to the final height and begin the procedure. These settings are pre-set based on the geometry dimensions but can be edited and saved based on the material or procedure being used (Fig. 5). 3.4 Rheological Characterization

The characterization of hydrogel mechanical properties can be challenging, sometimes requiring preliminary experiments to determine ideal operating parameters. For more common hydrogels (e.g., Matrigel™, collagen type I, or agarose), there are already published experimental data that can be referenced or used as a comparison [25]. These methods focus on three common

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procedures that provide critical information for adipose-derived hydrogels (see Note 8). For all procedures presented below, the plate temperature was stabilized at 4 °C before ~350 μL of liquid was pipetted onto the stage. The geometry was lowered to 550 μm, excess sample was removed, and then set to a final gap of 500 μm before running the procedure (see Note 9). Between procedures, the geometry was lifted to the loading gap and the stage and geometry were gently cleaned with 70% ethanol. Once the stage and geometry were clean, the next sample was loaded, and the process was repeated. 3.5 Strain/Shear Sweep of Pre-gel Solutions

Pre-gel viscosity is useful for adipose-derived hydrogels that might be injected or extruded prior to crosslinking. A viscosity that is too low will result in the solution escaping the target site or not holding shape, while too high of a viscosity will be difficult to inject or extrude, potentially clogging the needle. A flow ramp study determines viscosity by rotating the geometry in one direction with an increasing strain/shear rate. 1. Load the pre-gel sample (see Subheading 3.2, step 2). 2. Set the shear rate from 1 to 100 (s-1) and Peltier plate temperature to 4 °C. The Peltier plate temperature prevents premature crosslinking or gelation (see Note 10). 3. Raise the geometry to the loading gap and clean the stage with 70% ethanol after the procedure is complete.

3.6 Temperature Sweep of AdiposeDerived Hydrogels

Determining the sol-gel characteristics of any hydrogel is a critical step in collecting accurate mechanical properties. Without knowing if the hydrogel is fully crosslinked, all subsequent studies could be conducted on a partially crosslinking hydrogel and not truly representing the native network. Typical temperature sensitive hydrogels (e.g., Matrigel™, collagen type I, and agarose) will gel or crosslink within an hour. If the evaluated hydrogel requires a longer gelation or crosslinking period, additional steps must be taken (see Note 11). 1. Load the pre-gel sample (see Subheading 3.2, step 2). 2. Set the stage to 4 °C to prevent premature crosslinking or gelation. 3. Set the mechanical starting parameters to 1% strain and 1 Hz frequency to determine the sol-gel transition. 4. Modify the temperature settings based on the specific hydrogel. For example, a temperature ramp of 1 °C min-1 up to 37 °C, and for 1800 s post-soak at 37 °C is a typical setup for ECM-based hydrogels (see Note 12). 5. Raise the geometry to the loading gap and clean the stage with 70% ethanol after the procedure is complete.

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3.7 Strain/Shear Sweep of Fully Crosslinking AdiposeDerived Hydrogels

A strain/shear sweep can also be applied to fully crosslinked hydrogels to determine the linear viscoelastic region (LVE), or the region where samples can be evaluated without permanently damaging the microstructure (Fig. 2b, c). In this region, both moduli plateau and minimally change for a region of strain or shear rate. The resulting storage and loss modulus profiles provide the upper limit of the shear/strain rate that should be used. This is particularly useful when profiling or defining testable parameters for specific hydrogels. 1. Load the pre-gel sample (see Subheading 3.2, step 2) and crosslink the pre-gel sample on the stage following the crosslinking protocol. For example, Matrigel™ is crosslinked at 37 °C for at least 30 min. 2. Set the shear rate from 1 to 100 (s-1) with the Peltier plate temperature set to 37 °C. 3. Raise the geometry to the loading gap and clean the stage with 70% ethanol after the procedure is complete.

3.8 Frequency Sweep of Fully Crosslinked AdiposeDerived Hydrogels

Frequency sweeps are used to determine the time-dependent response from the hydrogel microstructure when evaluated in the LVE. Low frequencies simulate the hydrogel response to slow or minimal motion over a longer period, where high frequencies simulate fast motion over a short period. This procedure must be performed within the LVE region using the shear/strain determined from previous procedures (see Note 13). 1. Load the pre-gel sample (see Subheading 3.2, step 2) and crosslink the pre-gel sample on the stage following the crosslinking protocol. 2. Set the frequency range from 0.1 to 100 Hz, at 1% strain, with the Peltier plate temperature set to 37 °C. 3. Raise the geometry to the loading gap and clean the stage with 70% ethanol after the procedure is complete.

3.9

Data Processing

After completing the experimental procedures, the data can be plotted and analyzed within the rheometer software or exported as .csv files and plotted using external software. The data collected will depend on the experimental procedure, but can include storage and loss moduli, tan δ, viscosity, temperature, time, and other experimental parameters. When collecting mechanical data, it is recommended to perform the experiments in triplicate to ensure data collecting is consistent across different hydrogels. This will ensure the adipose-derived hydrogel is homogenous and/or the experimental setup and procedures are consistently evaluating the native microstructure.

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Fig. 7 Example plot of operating/handling viscosity of pre-crosslinked adiposederived hydrogel made using Trios rheology software (version 5.1.1). Sample was evaluated using a flow ramp from 1 to 100 s-1 with a plate temperature maintained at 4 °C to prevent premature crosslinking 3.10 Plotting Handling Viscosity of Pre-gel Solutions

1. Export the data from the experimental procedures as .csv files and open in Excel. 2. Plot the viscosity of the pre-gel solutions against the shear rate using a graphing or plotting software (see Note 14) (Fig. 7). 3. Plot other data including stress, torque, or velocity using the graphing or plotting software. 4. Review data to ensure there are no jumps or breaks in the data collection. Jumps or discontinuity would likely be due to poor contact with the geometry or the sample containing solid particles or aggregates (see Note 15).

3.11 Plotting Gelation and Sol-gel Transition

1. Export the data from the experimental procedures as .csv files and open in Excel. 2. Plot the storage and loss modulus against time or temperature using a graphing or plotting software (see Note 16) (Fig. 8). 3. Adjust the viscosity axis to logarithmic scale if the data collection was logarithmic. 4. Plot the other data including stress, viscosity, torque, or velocity using a graphing or plotting software. 5. Review the moduli profiles to ensure the data was accurately collected (see Note 17).

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Fig. 8 Example plot of solution to gelation transition of an adipose-derived hydrogel made using Trios rheology software (version 5.1.1). Sample was evaluated using a temperature ramp of 1 °C/min, from 4 to 37 °C, and maintained for an additional 1800 s (post-soak) with a 1% strain and 1 Hz frequency 3.12 Plotting Moduli of Fully Crosslinked Hydrogels

1. Export the data from the experimental procedures as .csv files and open in Excel. 2. Plot the storage and loss moduli against frequency using a graphing or plotting software (see Note 16) (Fig. 9). 3. Adjust the axes to a logarithmic scale if the data collection was logarithmic. 4. Plot the other data including stress, viscosity, torque, or velocity using a graphing or plotting software. 5. Review the moduli to ensure the storage modulus is located above the loss modulus and both lines are nearly parallel. This exemplifies a well-crosslinked adipose-derived hydrogel (see Note 18).

3.13 Instrument Shutdown

Once all procedures and data collection are complete are complete, shutdown the instrument using the reversal of the startup procedure. 1. Raise the geometry and thoroughly clean the stage using 70% ethanol after completing the final procedure. 2. Close the rheometer program. 3. Unscrew the geometry from the spindle by holding the spindle from the top and carefully holding the geometry. Once the geometry is removed, clean and dry the geometry using 70% ethanol and place back into the corresponding container.

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Fig. 9 Example plot of storage (solid) and loss (open) modulus for adiposederived hydrogels and resulting Tan δ, using Trios rheology software (version 5.1.1). Sample was evaluated by frequency sweeping from 0.1 to 100 Hz at 1% strain. Tan δ was calculated using the software and is the ratio of loss to storage modulus

4. Screw the safety cap back to the spindle by holding the spindle from the top and carefully screwing the safety cap. This needs to only be hand tight and over tightening can cause damage. 5. Turn off the power for the circulator and power supply. 6. Close the gas cylinder or air supply.

4

Notes 1. Xgel is a thermo-responsive hydrogel sourced from decellularized human tissue [30]. It was developed and optimized by Obatala Sciences, Inc. as an alternative to murine-derived and/or synthetic hydrogels. 2. This chapter is designed following the use of a TA rheometer and the Trios software. If different models or manufacturers are used, some of the steps may look different. However, the general workflow and operation should remain the same or similar. Especially with the types of experiments outlined and used to profile new adipose-derived hydrogels. 3. Depending on the instrument, a minimum PSI is required, e.g., 30 PSI for this protocol. Going above or below could damage the bearings of the spindle. The instrument should have safeguards warning if the air supply pressure is not in the proper operating window.

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4. A larger geometry is ideal when evaluating liquids as they reduce noise from torque and displacement of the geometry [29]. This should be monitored during an experimental procedure and will also be shown with the resulting data exhibiting variation. The geometry can then be screwed onto the spindle and the software will either identify the geometry automatically or you will need to select the geometry you loaded manually. 5. The calibration settings are defined as: (a) Geometry inertia—the inertia for each geometry will vary and is important to ensure accuracy for high frequency oscillations or low viscosity fluids. (b) Friction—the friction from the bearings used for the geometry are minimized, but still need to be accounted for. (c) Rotational mapping—bearings can have small variations in the torque behavior when completing one revolution that can cause inaccuracies. This calibration combines the absolute angular position with the microprocessor control of the motor to map the small variations. (d) Gap compensation—only needs to be calibrated when using temperature-controlled sweep and should be performed for the entire temperature range being used. 6. Calibration of geometries and instruments should occur at least monthly to ensure accurate data collection. This should be done following the manufacturer’s recommendations and protocol. 7. For consistent data collection ensure the stage temperature is maintained at a specified temperature. Labs can fluctuate in temperature that could then alter experimental replicates. For example, all experiments not using a set temperature should be conducted at 25 °C, or a relevant temperature. Large changes in humidity would also alter hydrogel performance, monitor and minimize if possible. 8. The methods presented here are only a few options and there are other applications of a rheometer that can evaluate compressive mechanical properties or other important material parameters [9, 31]. 9. To have a better understanding of the hydrogel network, all experiments should be performed with the hydrogels crosslinked on the stage, if possible. Hydrogels that are crosslinked, placed onto the stage, and then slightly compressed with geometry will provide data on a slightly deformed hydrogel network rather than the native network. When possible, it is better to perform the crosslinking after the pre-gel liquid is loaded onto the stage and making good contact with the geometry.

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10. Preliminary flow studies should use a wide range of shear values to ensure identification of the materials response. If the range is too narrow, it may be difficult to determine if the material is shear thinning or thickening, for example. Smaller or larger shear values can be included, but this range is typical for most hydrogel materials. 11. If a sample is not crosslinking on stage or requires a longer crosslinking duration, then evaporation could be a problem. It is possible to crosslink using the specified method and then transfer to the rheometer stage for evaluation. This will depend on the sample and how easy it is to handle and move. Additionally, it is possible to crosslink a large quantity and then using a biopsy punch create a hydrogel with a defined diameter that would fit the geometry being used. In this case, using a smaller geometry is appropriate, e.g., 8 mm, for evaluation. This is also applicable to tissues (decellularized or native), cellladen hydrogels, or other in vivo applications. (a) Other instruments also offer solvent traps or accessories to minimize evaporation for long-term studies. 12. A low temperature ramp slowly crosslinks the hydrogel to reduce evaporation. A long post-soak time will give the hydrogel enough time to fully crosslink and can be further extended to ensure the entire sol-gel profile is captured. 13. During the experiments, e.g., frequency sweep, you can monitor the success of the samples based on the displacement and torque plots. Both plots should follow a smooth, sin/cosine plot and initially should remain in phase. Once the frequency increases high enough, the displacement will lag compared to the torque and the plots will remain smooth, but out of phase. If the plots are not smooth: (a) Sample may be in good contact with geometry. (b) Interrogation settings (e.g., frequency) may need to be optimized. (c) Sample may have been damaged during experimental interrogation. 14. If the data collection was logarithmic, then adjust the viscosity axis to logarithmic scale. 15. Most adipose-derived hydrogels should exhibit shear thinning, or a gradual decrease in viscosity with an increase in shear or strain. 16. You can also overlay multiple variables on the X-axis to show the tan δ and temperature showing the entire gelation process (Fig. 8).

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17. The sol-gel transition profile will depend on the hydrogel, but a fully crosslinked hydrogel will result in a profile similar to Fig. 8. Where, the storage modulus remains above the loss modulus with both increasing until eventually approaching a plateau. Additional time should result in both moduli reaching a plateau where the final biomechanical data could be observed. When observing the sol-gel transition, it can be difficult depending on the crosslinking method. With thermo-responsive hydrogels, other factors can make smooth and accurate data collection difficult, e.g., evaporation or uneven gelation. 18. It is normal for both to slightly increase as the frequency increases. Additionally, at higher frequencies the sample may fail resulting in the storage modulus rapidly decreasing below the loss modulus. References 1. Sivashanmugam A, Arun Kumar R, Vishnu Priya M et al (2015) An overview of injectable polymeric hydrogels for tissue engineering. Eur Polym J 72:543–565 2. Bashir S, Hina M, Iqbal J et al (2020) Fundamental concepts of hydrogels: synthesis, properties, and their applications. Polymers (Basel) 12(11):2702 3. Huskin G, Chen J, Davis T et al (2023) Tissueengineered 3D in vitro disease models for highthroughput drug screening. Tissue Eng Regen Med:1–16 4. Lin X, Wang JL, Wu XY et al (2022) Marinederived hydrogels for biomedical applications. Adv Funct Mater 33:2211323 5. Hoshiba T (2019) Decellularized extracellular matrix for cancer research. Mater (Basel) 12(8):1311 6. Fernandez-Perez J, Ahearne M (2019) The impact of decellularization methods on extracellular matrix derived hydrogels. Sci Rep 9(1): 14933 7. Kokai LE, Sivak WN, Schilling BK et al (2020) Clinical evaluation of an off-the-shelf allogeneic adipose matrix for soft tissue reconstruction. Plast Reconstr Surg Glob Open 8(1): e2574 8. Belgodere JA, Zamin SA, Kalinoski RM et al (2019) Modulating mechanical properties of collagen-lignin composites. ACS Appl Bio Mater 2(8):3562–3572 9. Balaji S, Short WD, Padon BW et al (2023) Injectable antioxidant and oxygen-releasing lignin composites to promote wound healing.

ACS Appl Mater Interfaces 15(15): 18639–18652 10. Yoon H, Seo JK, Park TE (2023) Microphysiological system recapitulating the pathophysiology of adipose tissue in obesity. Acta Biomater 159:188–200 11. O’Halloran NA, Dolan EB, Kerin MJ et al (2018) Hydrogels in adipose tissue engineering-potential application in postmastectomy breast regeneration. J Tissue Eng Regen Med 12(12):2234–2247 12. Hayes DJ, Gimble JM (2022) Developing a clinical grade human adipose decellularized biomaterial. Biomat Biosyst 7:100053 13. Alkhouli N, Mansfield J, Green E et al (2013) The mechanical properties of human adipose tissues and their relationships to the structure and composition of the extracellular matrix. Am J Physiol Endocrinol Metab 305(12): E1427–E1E35 14. Divoux A, Cle´ment K (2011) Architecture and the extracellular matrix: the still unappreciated components of the adipose tissue. Obes Rev 12(5):e494–e503 15. Oyen ML (2013) Mechanical characterisation of hydrogel materials. Int Mater Rev 59(1): 44–59 16. Kloxin AM, Kloxin CJ, Bowman CN et al (2010) Mechanical properties of cellularly responsive hydrogels and their experimental determination. Adv Mater 22(31):3484–3494 17. Anseth KS, Bowman CN, Brannon-Peppas L (1996) Mechanical properties of hydrogels and their experimental determination. Biomaterials 17(17):1647–1657

Biomechanical Characterization of Adipose-Derived Hydrogels 18. Peppas NA, Huang Y, Torres-Lugo M et al (2000) Physicochemical foundations and structural design of hydrogels in medicine and biology. Annu Rev Biomed Eng 2(1):9–29 19. Akcay G, Luttge R (2021) Stiff-to-soft transition from glass to 3D hydrogel substrates in neuronal cell culture. Micromachines (Basel) 12(2):165 20. Eyckmans J, Boudou T, Yu X et al (2011) A hitchhiker’s guide to mechanobiology. Dev Cell 21(1):35–47 21. Lee S, Stanton AE, Tong X et al (2019) Hydrogels with enhanced protein conjugation efficiency reveal stiffness-induced YAP localization in stem cells depends on biochemical cues. Biomaterials 202:26–34 22. Baroli B (2007) Hydrogels for tissue engineering and delivery of tissue-inducing substances. J Pharm Sci 96(9):2197–2223 23. Callister WD, Rethwisch DG (2007) Materials science and engineering: an introduction. Wiley, New York 24. Tabor D (2000) The hardness of metals. Oxford University Press, Oxford

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25. Zuidema JM, Rivet CJ, Gilbert RJ et al (2014) A protocol for rheological characterization of hydrogels for tissue engineering strategies. J Biomed Mater Res B Appl Biomater 102(5): 1063–1073 26. Morrison FA (2001) Understanding rheology. Oxford University Press, Oxford 27. Ferry JD (1980) Viscoelastic properties of polymers. Wiley, New York 28. Mezger TG (2014) Applied rheology, 4th edn. Vincentz Network, Hanover 29. Mezger T (2020) The rheology handbook: for users of rotational and oscillatory rheometers. European Coatings 30. Belgodere JA, Lassiter HR, Robinson JT et al (2023) Biomechanical and biological characterization of XGel, a human-derived hydrogel for stem cell expansion and tissue engineering. Adv Biol:2200332 31. Belgodere JA, Son D, Jeon B et al (2021) Attenuating fibrotic markers of patient-derived dermal fibroblasts by Thiolated lignin composites. ACS Biomater Sci Eng 7(6):2212–2218

Chapter 13 Combination of Adipose-Derived Stromal/Stem Cells and Decellularized Adipose Tissue Hydrogel for Osteogenic Applications Kainat Ahmed, Haadia Tauseef, and Omair A. Mohiuddin Abstract Adipose-derived stromal/stem cells (ASCs) and decellularized adipose tissue (DAT) are adipose tissue products obtained from individuals undergoing fat removal procedures like liposuction, lipectomy, or breast reduction. DAT hydrogel is prepared by removing the cells from the adipose tissue and digesting it to form a liquid material that forms a gel at physiological temperature. ASCs seeded on DAT have displayed osteogenic potential in vitro and in animal models of bone defects. Herein, we describe the methods for preparing DAT hydrogel, ASC seeding in DAT hydrogel, osteogenic differentiation of ASCs, creation of critical-sized femur defect model in mice, its treatment with ASC-DAT hydrogel, and analyses. Key words Adipose-derived stromal/stem cells, Decellularized adipose tissue hydrogel, Osteogenic differentiation, Critical-sized femur defect model, Histology, Micro-computed tomography

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Introduction Bones possess efficient regenerative capability; therefore, most bone defects heal naturally. However, large bone defects, also referred to as “critical-sized bone defects,” fail to heal, thus requiring external medical intervention [1–4]. Presently, the standard of care for such conditions is bone grafting, which could be autologous or allogeneic in source [1]. Autologous tissue is generally unavailable in sufficient quantity for efficient bone regeneration. Additionally, the quality of autologous bone grafts is diminished in older individuals, postmenopausal women, or individuals with underlying health conditions such as diabetes and osteoporosis. Allogeneic bone grafts are available more abundantly; however, they present the risk of eliciting a host immune response and transmission of infectious diseases [2]. Therefore, to fulfill the demand for effective and safe bone grafts, there is a need to develop

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bone engineering strategies using materials that support bone formation and are minimally immunogenic. The ability of adipose stem cells (ASCs) to undergo osteogenic differentiation is well established [5, 6]. In addition to their ability to differentiate into osteoblasts in vitro, they have also displayed potent regenerative effects in preclinical models of bone defect [2, 7]. Scaffolds are an integral component of tissue engineering; they provide a three-dimensional (3D) space for the cells to attach, proliferate, and differentiate. Moreover, they allow for targeted delivery and retention of cells at the site of tissue injury. Several synthetic and natural scaffolds, including polylactic acid co-glycolic acid (PLGA), hydroxyapatite, collagen I, decellularized bone, and decellularized adipose tissue (DAT), have been shown to be compatible with ASCs and have displayed regenerative effects in criticalsized bone defect preclinical models [2, 3, 7–11]. DAT is prepared by the removal of cells from adipose tissue while preserving the extracellular matrix (ECM) [12, 13]. Both ASCs and DAT are obtained from adipose tissue, which is routinely discarded as medical waste following liposuction and lipectomy [5, 8]. Therefore, adipose tissue is an immense source of stem cells and biomaterial, which can be employed in combination for bone tissue engineering. Previous studies have shown that DAT-derived hydrogel is compatible with ASCs and supports their osteogenic differentiation. ASC-DAT hydrogel composites enhance bone regeneration in mouse models of critical-sized femur defects [2, 8]. This chapter will present the methods utilized for (1) the preparation of DAT hydrogel; (2) ASC seeding in the DAT hydrogel, their osteogenic differentiation, and analysis; and (3) the creation of mouse femur defect model, the application of ASC-DAT hydrogel to the defect site, and analysis of bone regeneration.

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Materials

2.1 Decellularized Adipose Tissue Preparation

1. Deionized (DI) water. 2. 1× phosphate-buffered saline (PBS). Dilute 10 mL of 10× PBS in DI water to make 100 mL 1× PBS. 3. 1 M sodium chloride (NaCl) solution. Dissolve 58.5 g of NaCl in 1 L of DI water. 4. 0.5 M NaCl solution. Dilute 500 mL of 1 M NaCl in DI water to make 1 L of 0.5 M NaCl. 5. 0.25% trypsin. 6. 100% isopropyl alcohol (IPA). 7. 1% Triton ×-100. Dilute 1 mL of Triton ×-100 in DI water to make 100 mL of 1% Triton ×-100.

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2.2 DAT Pre-gel Formation

1. DAT digestion medium. Dissolve 10 mg pepsin powder in 1 mL of 0.05 M hydrochloric acid (HCl).

2.3 Neutralization of DAT Pre-gel Solution and DAT Hydrogel Formation

1. 1 M sodium hydroxide (NaOH) solution. Dissolve 40 g of NaOH in 1 L of DI water.

2.4 In Vitro AdiposeDerived Stromal/Stem Cell Seeding and Osteogenic Differentiation in DAT Hydrogel

1. Human ASCs.

2.5 Evaluation of In Vitro Osteogenic Differentiation

2. 10× PBS.

2. Stromal media; DMEM/F12, 10% fetal bovine serum (FBS), 1% L-glutamine, and 1% penicillin/streptomycin. 3. Osteogenic differentiation media; stromal media, 10 nM dexamethasone, 20 mM β-glycerophosphate, and 50 μM Lascorbic acid. 1. RNA extraction kit. 2. 4% paraformaldehyde. 3. 30% sucrose solution. 4. Optimal cutting temperature (OCT) compound. 5. Von Kossa stain kit.

2.6 Critical-Sized Femur Defect Model and ASC-DAT Hydrogel Implantation

1. Stainless steel tubes, 22G and 19G.

2.7 Evaluation of Femur Defect Regeneration

1. Hematoxylin.

2. Diamond-coated rotary cutter wheel. 3. Isoflurane. 4. Scalpel blades.

2. Eosin. 3. Masson’s trichrome stain. 4. Safranin O. 5. Fast green. 6. Paraffin. 7. Embedding cassettes and molds.

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Methods The preparation of adipose tissue-derived hydrogel is divided into four steps: decellularizing adipose tissue, digestion of DAT, pre-gel formation, and gel formation.

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3.1 Decellularized Adipose Tissue (DAT) Preparation

1. Freeze the adipose tissue sample at -80 °C for 1 h and thaw at room temperature for 1 h; repeat this step two more times. 2. Place the samples in appropriately sized containers depending on the amount of tissue being decellularized (Fig. 1a) and add the wash solutions for decellularization in the sequence as stated below (see Note 1 for the volume of wash solutions). 3. Wash the samples in PBS several times to remove maximum debris and blood (see Note 2). 4. After that, unless otherwise stated, perform the following washes on an orbital shaker at room temperature. 5. DI water for 24 h. Replace with fresh DI water after 12 h. 6. 0.5 M NaCl solution for 4 h. 7. 1× PBS for 30 min. 8. 1 M NaCl solution for 4 h. 9. DI water overnight. 10. 0.25% trypsin for 2 h at 37 °C (see Note 3). 11. DI water for 1 h. 12. Repeat 100% IPA washes until the tissue loses its yellow appearance and turns white in color (see Note 4). 13. DI water for 1 h. 14. 1% Triton ×-100 for 48 h. Replace with fresh Triton ×-100 solution after 24 h. 15. 1× PBS for 24 h. Replace with fresh PBS (1×) after 12 h. 16. DI water for 24 h. Replace with fresh DI water after 12 h. This completes the preparation of DAT. 17. Remove the DAT from DI water and freeze it at -80 °C overnight (see Note 5). 18. Lyophilize the DAT until the complete removal of moisture from the tissue (see Note 6). 19. Powder the DAT using a bladed grinder. Cool the tissue with liquid nitrogen before and during the grinding process; this keeps the tissue hard and facilitates the grinding process. 20. Disinfect the DAT powder by spreading it evenly on a sterile surface and exposing it to ultraviolet (UV) radiation for 30 min (see Note 7). 21. Store the DAT powder in a sterile container at 4 °C until further use (Fig. 1b). 22. All subsequent steps must be performed under sterile conditions using a biosafety cabinet. All solutions used hereafter must be sterilized by filtration through a 0.22 μm filter.

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Fig. 1 Decellularized adipose tissue (DAT) hydrogel preparation. (a) Native adipose tissue, (b) DAT powder, (c) DAT pre-gel solution, and (d) DAT hydrogels (black arrow) in a 6-well plate with media 3.2 DAT Pre-gel Formation

1. Transfer the DAT powder to a fresh sterile tube and add the DAT digestion medium (10 mg pepsin per 1 mL of 0.05 M HCl) to the tube. For 50 mg of DAT powder, add 1 mL of digestion medium (see Note 8). 2. The digestion of DAT powder must be carried out at room temperature. 3. Let the tube stand vertically for 12 h to allow the DAT powder to soak in the digestion solution. Do not shake the tube during this period. 4. Place the tube horizontally on an orbital shaker and shake for 36 h. 5. Intermittently shake the solution on vortex mixer for 15 s every 2 h (see Note 9). 6. After 48 h of digestion, a viscous pre-gel solution of DAT will be formed (Fig. 1c). Store this solution at 4 °C.

3.3 Neutralization of DAT Pre-gel Solution and DAT Hydrogel Formation

1. Use ice-cold solutions for this procedure. 2. To neutralize 1 mL of pre-gel solution, add 50 μL of 1 M NaOH and 120 μL of 10× PBS (see Notes 10 and 11). 3. Mix gently on a vortex mixer. 4. Store this solution at 4 °C until further use. At this point, it is still a pre-gel that can be stored long term. To solidify it to a hydrogel-like form, proceed to the next step. 5. Incubate the neutralized DAT pre-gel solution at 37 °C for 30 min in a humidified incubator to form DAT hydrogel (Fig. 1d).

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3.4 In Vitro ASC Seeding and Osteogenic Differentiation in DAT Hydrogel

1. Count the ASCs and centrifuge the cell suspension at 200 g for 5 min. 2. Aspirate all the stromal media from the centrifuged cell suspension while carefully preserving the cell pellet at the bottom of the tube. 3. Resuspend the cell pellet in the neutralized DAT pre-gel solution at two million cells per mL (see Note 12). Mix gently to avoid the introduction of air bubbles in the hydrogel (see Note 13). 4. Pipette 200 μL of ASC-DAT pre-gel suspension at the center of each well in a 6-well plate (see Note 14). 5. Transfer the plate to a humidified cell culture incubator for 30 min to allow the pre-gels to solidify. A dome-shaped ASC-DAT hydrogel will be formed, which sticks to the cell culture plate. 6. After forming the ASC-DAT hydrogel, add 2 mL of osteogenic differentiation media to each well and place the plates back in the incubator. 7. Culture the ASC-hydrogel constructs for 28 days, with the media replaced every 2–3 days before analysis.

3.5 Evaluation of In Vitro Osteogenic Differentiation

1. For the analysis of osteogenic differentiation, perform quantitative reverse transcriptase polymerase chain reaction (qRT-PCR)-based analysis of the expression of osteogenic marker genes. 2. Harvest the ASC-DAT hydrogel constructs at day 28 by sliding a spatula under the hydrogel and lifting it off the plate and transfer the hydrogel to a cell lysis buffer (see Note 15). 3. Pulverize the hydrogel using a tissue homogenizer to release the cells. Alternatively, the hydrogel can be cryomilled by freezing it in liquid nitrogen and subsequently crushing it using a mortar and pestle. 4. Extract RNA from the samples using the manufacturer’s instructions for specific RNA extraction kits. 5. Amplify the osteogenic marker genes using qRT-PCR to confirm the osteogenic differentiation of ASCs in DAT hydrogel. Common osteogenic differentiation markers include collagen 1, Runt-related transcription factor 2 (RUNX2), alkaline phosphatase, osteopontin, osteonectin, osteocalcin, and osterix [6]. 6. To further analyze the osteogenic differentiation, perform histochemical assessment of the hydrogel frozen sections. 7. Fix the ASC-DAT hydrogel in 4% paraformaldehyde for 24 h. 8. Rinse the hydrogel with PBS and then submerge it in 30% sucrose solution at 4 °C overnight (see Note 16).

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Fig. 2 Von Kossa-stained section of adipos-derived stromal/stem cells–decellularized adipose tissue (ASC-DAT) hydrogel cultured in osteogenic differentiation media for 28 days. Arrows indicate calcium deposits, while cells (pink) and nuclei (red) are visible in the background. Scale bar = 100 μm

9. Rinse the hydrogel with OCT compound three times for 30–45 min (see Note 17). 10. Fill a cryomold up to three-fourth with fresh OCT medium. Transfer the ASC-DAT hydrogel to the mold and carefully place it in the center. Add more OCT on the top to cover the hydrogel. 11. Immediately transfer the mold to -80 °C for 24 h before sectioning the sample. 12. Prepare 7–10 μm sections of the samples using a cryotome. 13. Stain the sections with Von Kossa staining kit. The calcium deposits will be observed in gray/brown color, whereas the nuclei and cytoplasm of cells will appear red and pink, respectively (Fig. 2). 3.6 Critical-Sized Femur Defect Model and ASC-DAT Hydrogel Implantation

We will be discussing the creation of critical-sized femur defects in mice, the application of ASC-DAT hydrogel to the defect site, and the analysis of bone regeneration. The surgical procedure has been adapted from the methods initially described by Clough et al. [3, 14]. 1. Begin with the fabrication of intramedullary pins [2, 14]. Cut a 9 mm segment of 22G tube and a 3 mm segment of 19G tube using a diamond-coated rotary cutter. Apply glue in the middle of the 9 mm 22G tube and slide the 3 mm 19G tube. Measure the dimensions to make sure that there is 3 mm of 22G pin on each side of the central 3 mm 19G collar (Fig. 3a). Let the pin

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Fig. 3 Critical-sized femur defect model and analysis of bone regeneration. (a) Intramedullary pin. (b–e) Creation of femur defect, positioning of intramedullary pin, adipose-derived stromal/stem cells–decellularized adipose tissue (ASC-DAT) hydrogel implantation, and suturing to close the muscle and skin. (f) X-ray image of regenerated bone at week 6 post surgery. Histochemical analysis of regenerated bone using hematoxylin and eosin (H&E) (g), Masson’s trichrome (h), and safranin O (i) staining

dry, then clean it to remove any excess glue and disinfect it by placing it in 70% ethanol for 1 h. 2. Before the femur defect surgery and implantation of ASC-DAT hydrogel, the institutional animal care and use committee must approve the following surgical procedure. 3. Induce anesthesia in the mice using 3% isoflurane. After 2–3 min, check the adequacy of anesthesia by gently pressing its paw. Then shift the animal to the surgery table and supply 2% isoflurane through the nose cone to maintain anesthesia (see Note 18). 4. Apply eye ointment to both eyes to protect them from dryness. 5. Inject buprenorphine subcutaneously (0.1 mg/kg) for pain management.

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6. Turn the animal on its side with the left hind limb facing upward. Remove hair from the surgical site using clippers from the knee to the hip to expose the entire thigh and hip area. Disinfect the surgical site using 70% ethanol. 7. Locate the femur by touching the thigh and then, using a scalpel blade, make a longitudinal incision along the femur from the knee to the hip joint. 8. Blunt dissect under the skin to expose the thigh muscle. Locate where the septa of both muscles meet (it is a white line on the muscle, and the femur lies underneath it). 9. Make an incision on the white line to expose the femur and elevate the femur shaft by sliding a blunt periosteal elevator under it (see Note 19). 10. Remove the soft tissue on the bone using a scalpel blade (see Note 20). 11. Locate the center of the bone and cut a 3 mm segment using a diamond-coated rotary cutter wheel. Irrigate the bone with cold PBS during the cutting process. 12. Ream the medullary cavity on both sides using a 22G pin to clear the bone shaft before the implantation of the intramedullary pin. 13. Insert the prefabricated intramedullary pin (with a 3 mm collar in the middle) into the medullary cavity on both sides (Fig. 3b). 14. Move the leg to make sure that the pin is fit snug and moves with the bone. 15. The muscle surrounding the bone creates a pocket for the implantation of DAT hydrogel. 16. Place the ASC-DAT hydrogel in the pocket and wind it around the bone using forceps (Fig. 3c). 17. Bring the muscle back to position and stitch it using 5/0 absorbable suture (Fig. 3d). Then reposition the skin and stitch using 5/0 nylon sutures (Fig. 3e). 18. Transfer the mice to individual cages after the surgery. Warm them with a warming blanket or heating lamp until they recover from anesthesia. 19. Each cage should have an igloo-type nest and water and food on the cage floor for the initial few days post surgery. 20. Inject buprenorphine every 12 h for the next 48 h after surgery (see Note 21). 21. Finally, euthanize the animals at the desired time point and extract the femur by cutting at the hip joint and the knee. Preserve the femur in 10% formalin until further analysis.

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3.7 Evaluation of Femur Defect Regeneration

1. For the analysis of bone regeneration by micro-computed tomography (micro-CT), wrap the femur in a gauze moistened by PBS to prevent drying during the micro-CT imaging. Position the femur vertically and scan at 100 kV, 100 μA, 8 μm resolution, copper/aluminum filter, 360° scan with 0.2° rotation step, 1553 ms exposure time, frame averaging (10), random movement (10), and flat field and geometrical correction “ON” (see Note 22). The 3 mm central collar of the intramedullary pin guides the region of new bone formation and is designated as the region of interest (ROI) for analyses. Determine the bone volume, bone area, trabecular thickness, and other bone parameters within this ROI (Fig. 3f) [2]. 2. For the histochemical assessment of bone regeneration, partially decalcify the bone using Immunocal™ for 24 h (see Note 23). Embed the bone horizontally in paraffin and make 5 μm sections along the length of the bone above the intramedullary pin (see Note 24) [2]. 3. Stain the tissue sections using hematoxylin and eosin (H&E), Masson’s trichrome (MT), and safranin O (SO) counterstained with fast green. H&E is used to observe the overall regenerated tissue area and its gross histological features (Fig. 3g). MT is used to identify the mineralized (blue) versus nonmineralized (red) regions in the regenerated tissue (Fig. 3h). SO is used to identify the cartilaginous region (red) versus the rest of the tissue (green) in the regenerated tissue (Fig. 3i).

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Notes 1. Add all wash solutions at a volume of 2 mL per 1 g of adipose tissue except trypsin. 2. The number of PBS washes will vary each time depending on the amount of blood and clots in the tissues. Continue with the washing until the wash solution appears clear of blood and debris. 3. Add trypsin at a volume of 1 mL per 1 g of adipose tissue. 4. The number of IPA washes will vary each time depending on the amount of oil in the tissue. Fresh IPA should be replaced with the IPA concentrated by the oil removed from the tissue every 2–4 h. Complete removal of oil from the tissue can take between 24 and 48 h. 5. Alternatively, the DAT can be frozen in liquid nitrogen for 20–30 min before lyophilization. 6. Generally, 48–72 h of lyophilization is sufficient, but it can take longer if working with a large amount of tissue. Complete

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dehydration is important as it makes the subsequent crushing of the tissue more efficient. 7. Alternatively, the DAT powder can be disinfected by exposure to gamma radiation for more effective sterilization. If access to UV or gamma is not available, then the entire process must be performed under a sterile biosafety cabinet from the beginning and all wash solutions must be sterilized by filtration through a 0.22 μm filter. 8. In the present method, 1 mg pepsin per 5 mg adipose tissue has been used. However, several other studies have shown that 1 mg of pepsin adequately digests up to 10 mg of adipose tissue [15, 16]. 9. A propeller mixer-type device can also be used to agitate the solution and enhance the digestion of DAT, provided that the sterility of the system is ensured. 10. Theoretically, equally concentrated solutions of HCl and NaOH neutralize each other at a ratio of 1:1. Therefore, 50 μL of 1 M NaOH is required to neutralize 1 mL pre-gel solution (prepared with 0.05 M HCl). However, the exact neutralization point should be determined for each batch of HCl and NaOH prepared due to slight batch-to-batch variability. 11. To make the hydrogel isotonic, 120 μL of 10× PBS is added to 1 mL of pre-gel containing 50 μL of 1 M NaOH, which makes an approximately 1× solution of PBS in a total pre-gel volume of 1.170 mL. 12. At a concentration of two million cells per mL of DAT hydrogel, the ASCs are at a high confluence level, which is appropriate for differentiation [8]. 13. The neutralized DAT pre-gel is a viscous solution. Use sterile scissors to cut the end of a 1000 μL pipette tip to make pipetting easier. 14. The ASC-pre-gel volume can be adjusted depending on the type of plate used in the experiment. In a 6-well plate, a 200 μL ASC-DAT hydrogel dome is easily submerged in 2 mL of media, allowing adequate media–hydrogel contact. 15. Any cell lysis buffer provided with an RNA extraction kit may be used. The volume of buffer to be used will depend on the manufacturer’s instructions for the specific product. Alternatively, the construct can be transferred to Trizol™ for RNA extraction. If the RNA extraction is not intended immediately, then snap-freeze the ASC-DAT hydrogel with liquid nitrogen and store it at -80 °C. 16. Sucrose solution works as a cryoprotectant. The hydrogel can be left in 30% sucrose solution for up to 1 week at 4 °C.

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17. An additional recommended.

overnight

OCT

compound

wash

is

18. The concentration of isoflurane must be adjusted depending on the weight of the animal. The concentration presented in the method is optimal for 25–35 g mice. 19. A spatula can be used for this purpose. Refer to Clough et al. [14] for further details. 20. Carefully remove all soft tissue from the bone to avoid hindrance while cutting the bone. 21. If the animal displays signs of distress, continue the buprenorphine injections for another 24 h. 22. Due to the presence of stainless steel pin, a high voltage is required to visualize the bone. The parameters were optimized for mouse femurs containing a 9 mm stainless steel pin. 23. Immunocal™ is a formic acid-based decalcifier; alternatively, EDTA can be used for decalcification to better preserve the tissue for immunohistochemical analysis. The endpoint of decalcification can be determined by touching the bone to see if the tissue has softened and become flexible. 24. While embedding, press the bone down as the paraffin cools to flatten it so that the entire length of the bone can be obtained in the sections. While sectioning, if you have consumed all the tissue and reached the intramedullary pin, the paraffin can be melted and the other side of the bone can be positioned at the bottom to get more tissue sections. The microtome blade will be damaged if it hits the intramedullary pin; therefore, observe caution while sectioning.

Acknowledgments This work was conducted at the Tulane University’s School of Medicine (New Orleans, LA, USA) and funded by the Musculoskeletal Transplant Foundation. References 1. Venkataiah VS, Yahata Y, Kitagawa A et al (2021) Clinical applications of cell-scaffold constructs for bone regeneration therapy. Cell 10(10):2687 2. Mohiuddin OA, Campbell B, Poche JN et al (2019) Decellularized adipose tissue hydrogel promotes bone regeneration in critical-sized mouse femoral defect model. Front Bioeng Biotechnol 7:211

3. Clough BH, McCarley MR, Krause U et al (2015) Bone regeneration with osteogenically enhanced mesenchymal stem cells and their extracellular matrix proteins. J Bone Miner Res 30(1):83–94 4. Schemitsch EH (2017) Size matters: defining critical in bone defect size! J Orthop Trauma 31(Suppl 5):S20–S22. https://doi.org/10. 1097/bot.0000000000000978

ASC and Decellularized Adipose Hydrogel Osteogenic Combination 5. Rahman G, Frazier TP, Gimble JM et al (2022) The emerging use of ASC/scaffold composites for the regeneration of Osteochondral defects. Front Bioeng Biotechnol 10:893992 6. Harrison MAA, Al-Ghadban SI, O’Donnell BT et al (2022) Chapter 3—establishing the adipose stem cell identity: characterization assays and functional properties. In: Kokai L, Marra K, Rubin JP (eds) Scientific principles of adipose stem cells. Academic Press, pp 23–56 7. Strong AL, Hunter RS, Jones RB et al (2016) Obesity inhibits the osteogenic differentiation of human adipose-derived stem cells. J Transl Med 14(1):27. https://doi.org/10.1186/ s12967-016-0776-1 8. Mohiuddin OA, O’Donnell BT, Poche JN et al (2019) Human adipose-derived hydrogel characterization based on in vitro ASC biocompatibility and differentiation. Stem Cells Int 27(9276398):1 9. Munmun F, Mohiuddin OA, Hoang VT et al (2022) The role of MEK1/2 and MEK5 in melatonin-mediated actions on osteoblastogenesis, osteoclastogenesis, bone microarchitecture, biomechanics, and bone formation. J Pineal Res 73(2):28 10. Mohiuddin OA, Motherwell JM, Rogers E et al (2020) Characterization and proteomic analysis of decellularized adipose tissue hydrogels derived from lean and overweight/obese human donors. Adv Biosyst 4(10):11

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11. Roddy E, DeBaun MR, Daoud-Gray A et al (2018) Treatment of critical-sized bone defects: clinical and tissue engineering perspectives. Eur J Orthop Surg Traumatol 28(3): 351–362. https://doi.org/10.1007/s00590017-2063-0 12. Mohiuddin OA, Campbell B, Poche JN et al (2019) Decellularized adipose tissue: biochemical composition, in vivo analysis and potential clinical applications. Adv Exp Med Biol. https://doi.org/10.1007/5584_2019_371 13. Flynn LE (2010) The use of decellularized adipose tissue to provide an inductive microenvironment for the adipogenic differentiation of human adipose-derived stem cells. Biomaterials 31(17):4715–4724. https://doi.org/10. 1016/j.biomaterials.2010.02.046 14. Clough BH, McCarley MR, Gregory CA (2015) A simple critical-sized femoral defect model in mice. J Vis Exp 97:e52368 15. Young DA, Ibrahim DO, Hu D et al (2011) Injectable hydrogel scaffold from decellularized human lipoaspirate. Acta Biomater 7(3): 1040–1049. https://doi.org/10.1016/j. actbio.2010.09.035 16. Zhao Y, Fan J, Bai S (2019) Biocompatibility of injectable hydrogel from decellularized human adipose tissue in vitro and in vivo. J Biomed Mater Res B Appl Biomater 107(5): 1684–1694. https://doi.org/10.1002/jbm. b.34261

Chapter 14 Preparation of Decellularized Amniotic Membrane and Adipose-Derived Stromal/Stem Cell Seeding Haadia Tauseef, Kainat Ahmed, Asmat Salim, and Omair A. Mohiuddin Abstract Amniotic membrane, being part of the placenta, is discarded as medical waste after childbirth. It can be decellularized to convert it into an acellular material while retaining the extracellular matrix. Such amniotic membrane grafts support stem cell adhesion, growth, and proliferation. These properties make it a useful candidate to be used as a bio-scaffold in regenerative medicine. This chapter describes a method for the decellularization of the amniotic membrane. Furthermore, the method for seeding adipose-derived stem cells on the decellularized amniotic membrane is described. Key words Adipose-derived stromal/stem cells, Decellularized human amniotic membrane, DNA extraction, Histology, Cell seeding, Silicone sheet

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Introduction The human amniotic membrane (HAM) is a potent bio-scaffold for tissue engineering because of its abundance of extracellular proteins, cytokines, and growth factors [1]. After childbirth, HAM is disposed of as medical waste, thus making it a readily available, economical, and abundant biomaterial source. It has displayed biological properties such as being angiogenic, anti-inflammatory, antifibrotic, antimicrobial, and immune privileged [2]. Fresh HAM has been used for regenerative applications in the eye and skin for several decades [3–7]. HAM can also be processed to remove the cells to minimize the immunogenic and inflammatory responses that lead to implant rejection [8]. Decellularized HAM (dHAM) has been re-seeded with several cell types, including embryonic stem cells, tissue-specific stem cells, mesenchymal stem cells (MSCs), and induced pluripotent stem cells [9]. The dHAM seeded with adipose-derived stromal/stem cells (ASCs) can be beneficial in regenerative therapies, especially for the regeneration of skin [10]. ASCs are isolated from adipose tissue,

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which is also a readily available source of human tissue that can be collected as medical waste after lipoaspiration procedures in significant amounts. Among the several adipose tissue depots in the body, subcutaneous adipose tissue is the most clinically significant source of multipotent ASCs, which can be obtained from the belly, thigh, and arm by subcutaneous lipoaspiration [11, 12]. The therapeutic potential of HAM in combination with ASCs has been evaluated in preclinical models of osteoarthritis, periodontitis, myocardial infarction, and burn wounds [13–16]. This chapter presents the methods used for the (1) preparation of dHAM, (2) characterization of dHAM, (3) ASC seeding on the dHAM, and (4) analysis of dHAM seeded with ASCs.

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Materials Use sterile deionized (DI) water to prepare all solutions. Moreover, all solutions can be stored at room temperature unless otherwise stated.

2.1 Decellularization of HAM

1. 0.25% trypsin/EDTA solution: To be stored at -20 °C. Thaw it at 37 °C before use. 2. Phosphate-buffered saline (PBS) solution (pH 7.4): Weigh and add 8 g of sodium chloride, 0.2 g of potassium chloride, 1.44 g of sodium phosphate dibasic, and 0.245 g of potassium phosphate monobasic to 800 mL DI water. Adjust the pH to 7.4 and add DI water until the volume is 1 L. 3. DI water.

2.2 Characterization of dHAM by DNA Extraction

1. DNA extraction kit.

2.3 Characterization of dHAM by Histochemical Analysis

1. Chrome Alum Gelatin solution: Add 1 g of gelatin to 100 mL DI water heated at 65 °C and stir until completely dissolved. Further, cool down the solution at room temperature and add 0.1 g of chromium potassium sulfate. Filter the solution using filter paper.

2. NanoDrop™. 3. 1% agarose gel. Dissolve 1 g agarose in 1 mL Tris acetateEDTA buffer.

2. 4% paraformaldehyde (PFA) solution. 3. 4′-6-Diamidino-2-phenylindole (DAPI) stain working solution: Dissolve 5 mg DAPI in 10 mL PBS to make a 0.5 mg/ mL stock solution. Prepare a 0.5 μg/mL DAPI working solution by diluting 1 μL of stock solution in 999 μL of PBS solution. 4. PBS solution.

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5. Dibutylphthalate Polystyrene Xylene (DPX) mounting media. 6. Hematoxylin stain. 7. Eosin Y stain. 8. DI water. 9. Optimal cutting temperature (OCT) medium. 2.4 In Vitro ASC Culture on dHAM

1. Silicone sheet (1 mm thickness). 2. PBS solution. 3. Decellularized HAM. 4. Stromal media: Dulbecco’s Modified Eagle Medium (DMEM), 1% L-glutamine, 1% pen strep, and 10% fetal bovine serum. 5. ASCs.

2.5 Analysis of Cell Attachment and Viability by Calcein AM Staining

1. Calcein AM stains working solution: Dissolve 1 mg of calcein AM in 1 mL of dimethylsulfoxide to make a 1 μg/μL stock solution. Prepare a 10 ng/μL calcein AM working solution by diluting 10 μL of stock solution in 990 μL of serumfree DMEM. 2. PBS solution.

2.6 Analysis of Cell Attachment by Crystal Violet Staining

3

1. PBS solution. 2. 3% crystal violet stain: Dissolve 3 g of crystal violet in 20 mL of 95% alcohol. Further, add 80 mL of 1% aqueous ammonium oxalate to the crystal violet–alcohol solution and filter.

Methods

3.1 Decellularization of HAM

1. Procure the complete placenta and fetal membranes from a healthy human donor and transfer them to a sterile container partially filled with PBS-EDTA solution for transportation. All human tissues must be obtained in accordance with the international standards for the protection of patients and their healthcare privacy and autonomy. 2. Store the placenta at -80 °C until further use. 3. The following steps must be carried out using a biosafety cabinet under sterile conditions. The instruments, containers, and vessels must be sterilized before use. All the wash steps must be performed on a shaker. 4. Carefully separate the fetal membrane (containing both the amniotic and chorionic membranes) from the placenta. Starting from the edges, move toward the umbilical cord on the fetal surface using forceps and scissors (Fig. 1a).

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Fig. 1 Gross examination of the human placenta, umbilical cord, and amniotic membrane (a), freshly separated amniotic and chorionic membranes (b), and decellularized human amniotic membrane (dHAM) (c)

5. Peel and separate the amniotic membrane, the innermost fetal membrane, from the chorionic membrane (the thicker membrane) using forceps (Fig. 1b). 6. After isolation, rinse the HAM with PBS solution until blood remnants are completely washed off (see Notes 1–4). 7. Drain the PBS solution and transfer the HAM to a 50 mL conical tube. Add 2 mL of 0.25% trypsin-EDTA per gram of HAM. Incubate it for 30 min at 37 °C [17, 18] (see Notes 5 and 6). 8. Transfer and spread the trypsin-treated HAM on a Petri plate. Gently scrape off the cells using a cell scraper from both sides of the membrane while holding it in place with the help of forceps (see Note 7). 9. Transfer the HAM to the conical tube and wash it with PBS solution to remove the remaining cellular content. Add fresh PBS solution and incubate it for 2 h at room temperature with gentle shaking. 10. Replace the PBS solution with DI water and incubate it at 4 °C overnight. 11. Drain the DI water, add PBS solution, and incubate at room temperature for 2 h, placed on a shaker. This completes the preparation of dHAM (Fig. 1c) (see Note 8). 12. For long-term preservation and stability, drain the liquid content and store the dHAM at -20 °C overnight, then spread the membranes on Petri plates using a spatula and lyophilize them using a freeze-dryer for 24 h (see Notes 9 and 10). 13. To disinfect the dHAM, expose each side to ultraviolet radiation for 15–20 min. 14. The dHAM can be cut to the required size using a scalpel blade or scissors.

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15. The lyophilized form of dHAM can be stored at 4 °C for an extended period until further use. 16. The confirmation of decellularization and characterization of dHAM is discussed in Subheading 3.2. 3.2 Characterization of dHAM by DNA Extraction

1. Chop the HAM and dHAM into fine pieces and extract the DNA using any commercial genomic DNA extraction kit. 2. Quantify the extracted DNA using a NanoDrop instrument or another spectrophotometric method. 3. Confirm the removal of cells from the dHAM by observing a lower amount of DNA in the dHAM compared to their respective HAM samples (see Note 11). 4. Prepare a 1% agarose gel and load the DNA samples obtained from HAM and dHAM in the wells. 5. Run the gel at 120 V for around 40 min. Visualize the DNA bands using a gel imaging system (see Note 12).

3.3 Characterization of dHAM by Histochemical Analysis

1. The glass slides must be coated with an adhesive substance to ensure that the tissue sections are retained on glass slides throughout the staining and washing procedures. Gelatin solution mixed with chrome alum is the most commonly utilized material for coating glass slides. Chrome-gelatin-coated slides are called “subbed slides” [19–21]. Dip the glass slides in Chrome Alum Gelatin solution for 5–10 s and drain the excess solution onto tissue paper. Repeat this step three times. Let the slides dry for a duration of 1 h at room temperature in a dustfree environment by placing them vertically on a wooden slide rack (see Notes 13 and 14). 2. Fix the HAM and dHAM samples in 4% PFA for 24 h. Rinse them with PBS for 5 min, two times. Fill a cryomold up to three-fourth with fresh optimal cutting temperature (OCT) medium. Position the membranes vertically in the OCT using forceps. Such an orientation will produce sections that will present the amniotic membrane’s epithelial and basal layers. 3. Immediately transfer the mold to -20 °C for 24 h before sectioning the membrane (see Note 15). Prepare 7–10 μm sections on gelatin-coated glass slides. 4. Immerse the slides in DI water for 2 min to wash the OCT off. Wash the sections with PBS solution three times for 5 min each and stain to observe the nuclei and extracellular matrix (ECM). 5. Pipette DAPI stain on the sections and place the slide in the dark and incubate it for 15 min at room temperature (see Notes 16–18). 6. Wash the section with PBS solution thrice for 5 min each. Remove the PBS, mount the section by adding a drop of

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Fig. 2 Histochemical analysis of human amniotic membrane (HAM) and decellularized HAM (dHAM) by staining with 4′-6-diamidino-2-phenylindole (DAPI) (a, b) and hematoxylin and eosin (H&E) (c, d). Scale bar = 100 μm

DPX mounting media, and carefully place the coverslip to avoid air bubbles’ entrapment. 7. Observe the sections under a fluorescence microscope (excitation/emission: 377/447 nm) [22]. The nuclei will appear as blue rounded structures, indicating the presence of cells (Fig. 2a, b) (see Note 19). 8. For hematoxylin and eosin (H&E) staining, first add the hematoxylin stain covering the sections for 10 min and then wash under running tap water (see Note 20). 9. Add the Eosin Y for 1 min covering the sections and wash the sections with DI water. 10. Air-dry the slide completely. 11. Put a drop of DPX mounting media on the slide and carefully place a coverslip to avoid air bubbles’ entrapment.

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12. Perform brightfield microscopy using a compound light microscope. The nuclei will appear purple, while the cytoplasm and ECM will appear pink (Fig. 2c, d) (see Note 21). 3.4 In Vitro ASC Culture on dHAM

Efficient seeding of ASCs on the dHAM cannot be achieved on regular cell culture plates since the cells escape the dHAM and attach preferentially to the plate. To overcome this problem, the precoating of wells with silicone sheets is described below, which efficiently inhibits cell adhesion to the plate and enhances the attachment of cells to the dHAM. The notes also discuss other alternatives (see Note 22). 1. Procure prefabricated 1-mm-thick silicone sheets. 2. Cut 15-mm-diameter discs from the silicone sheet to prepare nonadherent bases, which fit at the bottom of the well in a 24-well culture plate (see Note 23). 3. Autoclave the silicone discs at 121 °C for 30 min to sterilize them. 4. Place the discs at the bottom of a 24-well culture plate using forceps and press them down to ensure that they coat the surface of the wells (Fig. 3a). 5. Cut the dHAM into 15 mm2 pieces and place them in the silicone sheet-coated 24-well plate with the help of forceps (Fig. 3b) (see Note 24). 6. Add 1 mL of PBS solution in each well containing dHAMs and incubate the plate in a humidified incubator at 37 °C for 24 h (Fig. 3c) (see Note 25). 7. Remove the PBS from the wells by using a pipette. 8. Suspend the ASCs in stromal media and pipette the cell suspension (50,000 cells/200 μL) on top of the dHAM (Fig. 3d). 9. Place the plate in a cell culture incubator. 10. After 4–5 h, remove the media from the wells using a pipette. Then lift each ASC-seeded dHAM from the plate, flip and replace it in the same well. 11. Repeat step 8 for the other side of the dHAM (see Note 26). 12. Place the plate in a cell culture incubator for 1 h. After that, add 800 μL of media in each well and place the plate in the incubator. 13. Replace the media every alternate day until the end of the experiment (see Note 27).

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Fig. 3 In vitro adipose-derived stromal/stem cells (ASC) culture on decellularized human amniotic membrane (dHAM). Silicone sheet-coated wells (a), dHAM placed in coated wells (b), dHAM in phosphate-buffered saline (PBS) (c), and cell-seeded dHAM in stromal media (d) 3.5 Analysis of Cell Attachment and Viability by Calcein AM Staining

1. Remove the ASCs-seeded dHAMs from each well and place them in separate fresh wells of a 24-well plate. 2. Wash the ASCs-seeded dHAMs with PBS for 5 min, two times. 3. Add 200 μL of calcein AM stain in each well and incubate at 37 °C for 30 min (see Note 28). 4. Image the stained cells using a fluorescence microscope at the excitation/emission wavelength of 469/525 nm [22]. The viable ASCs will appear green, as shown in Fig. 4a, b.

3.6 Analysis of Cell Attachment by Crystal Violet Staining

1. Fix the ASCs-seeded dHAM in 4% PFA for 24 h. 2. Remove the ASC-seeded dHAMs from each well and place them in separate fresh wells of a 24-well plate. 3. Wash the dHAM with PBS solution twice for 5 min each. 4. Add the 3% crystal violet stain covering the dHAM for 30 min. 5. Wash the stain carefully under running tap water until the water runs clear of the stain. The attached ASCs will appear purple, as shown in Fig. 4c, d (see Note 29).

4

Notes 1. After placing the HAM, fill the container up to 80% with PBS. Leaving some free space in the container will allow efficient agitation and washing of the membrane. 2. As the PBS turns turbid, replace it with fresh PBS. Repeat this step until the PBS appears clear. 3. Place the container horizontally on the shaker to ensure efficient washing.

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Fig. 4 Mesenchymal stem cells seeded on decellularized human amniotic membrane (dHAM), stained with calcein AM (a, b) and crystal violet (c, d). Scale bar = 100 μm

4. Pipettes can be used to remove the used PBS and all subsequent wash solutions. Alternatively, for large volumes, the wash solution and HAM can be poured on a strainer; the tissue is retained on the strainer, while the wash solution can be collected in a vessel placed underneath the strainer. 5. Other studies have suggested incubation times of 90 and 120 min and a working concentration of 0.1% trypsin-EDTA [23–26]. Therefore, the incubation time and enzyme concentration may require adjustment based on the amount of tissue being processed and other specific experimental conditions. 6. For optimal enzymatic decellularization, the tubes must be placed on a shaking incubator or water bath at 37 °C. 7. The HAM should be handled gently to avoid tearing. If a cell scraper is unavailable, a spatula or another instrument with a flattened edge can scrape the cells off. 8. The dHAM can be stored in PBS solution at 4 °C for short term.

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9. Spreading of dHAM on Petri plates prevents its folding and facilitates the cutting of uniform-sized pieces and subsequent cell seeding. If moisture is observed in the dHAM after 24 h of lyophilization, repeat the step until it is completely dried. 10. When removing the dried dHAM from the plates, observe caution since it can stick to the plate. Harsh scraping or pulling may result in the tearing of the dHAM. 11. Some researchers suggest that in an efficiently decellularized tissue, the DNA amount should be reduced to below 50 ng per mg of dry tissue and the DNA fragments should not be larger than 200 bp [27]. 12. The HAM samples will present sharp bands of genomic DNA, whereas faint or no bands will be observed in the dHAM lanes [28]. 13. Coated slides can be transferred to a slide box and stored at room temperature until further use. 14. Alternatively, commercially available adhesive microscopic slides can also be utilized. 15. Tissue-embedded cryomolds can be stored for the long term at -80 °C. 16. The sections can be treated with 0.2% Triton ×-100 solution for 10 min to permeabilize them before adding the DAPI stain. This aids in efficient penetration of the stain inside the cell. 17. All DAPI staining and imaging steps must be performed in the dark. 18. Add the wash and stain solutions gently to avoid washing the sample off the slide. 19. The nuclei will be observed in the HAM, whereas very few or no nuclei will be observed in the dHAM (Fig. 2a, b). 20. Do not position the section directly under tap water. Run tap water on a part of the slide away from the section and tilt the slide to let the water flow very slowly and gently over the sections. 21. The purple nuclei will be visible in the HAM, confirming the presence of cells, whereas only the pinkish ECM will be observed in the dHAM samples (Fig. 2c, d). 22. The cell attachment surface coated with 1–2% agarose and glass coverslips has also been found to be useful in providing a nonadhesive surface [29–32]. Alternatively, ultra-low-adherent cell culture plates can also be used. 23. The size of the silicone discs must correspond to the size of the well in order to completely cover the surface and prevent cell attachment to it.

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24. Cutting the corners of dHAM is optimal to make it circular to fit the well. This ensures that a maximum number of ASCs come in contact with dHAM when the cell suspension is poured on it. 25. Soaking the dHAMs in PBS prior to cell seeding improves cell attachment. 26. Seeding on both sides yields more consistent results in terms of cell attachment and viability compared to seeding on one side only. 27. Remove or add media by touching the pipette tip against the wall of the well rather than touching the dHAM as this may damage the dHAM or disrupt cell attachment. 28. Calcein AM stain is light sensitive; therefore, the staining should be performed in the dark. To avoid interaction with light, cover the culture plate with aluminum foil. 29. The pictures presented in Fig. 4 display umbilical cord-derived MSCs seeded on dHAM. References 1. Fe´nelon M, Catros S, Meyer C et al (2021) Applications of human amniotic membrane for tissue engineering. Membranes 11(6):387 2. Deus IA, Mano JF, Custo´dio CAJAB (2020) Perinatal tissues and cells in tissue engineering and regenerative medicine. Acta Biomater 110: 1–14 3. Wee SW, Choi SU, Kim JC (2015) Deep anterior lamellar keratoplasty using irradiated acellular cornea with amniotic membrane transplantation for intractable ocular surface diseases. Korean J Ophthalmol 29(2):79–85 4. Nouri M, Ebrahimi M, Bagheri T et al (2018) Healing effects of dried and acellular human amniotic membrane and mepitelas for coverage of skin graft donor areas; a randomized clinical trial. Bull Emerg Trauma 6(3):195 5. Fairbairn NG, Randolph MA, Redmond RW et al (2014) The clinical applications of human amnion in plastic surgery. J Plast Reconstr Aesthet Surg 67(5):662–675 6. Schmiedova I, Dembickaja A, Kiselakova L et al (2021) Using of amniotic membrane derivatives for the treatment of chronic wounds. Membranes 11(12):941 7. Tabriz AG (2022) Allogeneic, xenographic, synthetic, bioengineered, and composite products for wound healing and soft tissue grafting. Healthy Blue Surg 8. Fenelon M, Etchebarne M, Siadous R et al (2020) Assessment of fresh and preserved

amniotic membrane for guided bone regeneration in mice. J Biomed Mater Res A 108(10): 2044–2056 9. Scarritt ME, Pashos NC, Bunnell BA et al (2015) A review of cellularization strategies for tissue engineering of whole organs. Front Bioeng Biotechnol 3:43 10. Taghiabadi E, Beiki B, Aghdami N et al. (2019) Cultivation of adipose-derived stromal cells on intact amniotic membrane-based scaffold for skin tissue engineering. In: Skin stem cells: methods and protocols, p 201–210 11. Tsuji W, Rubin JP, KGJW M (2014) Adiposederived stem cells: implications in tissue regeneration. World J Stem Cells 6(3):312 12. Si Z, Wang X, Sun C et al (2019) Adiposederived stem cells: sources, potency, and implications for regenerative therapies. Biomed Pharmacother 114:108765 13. Bhattacharjee M, Escobar Ivirico JL, Kan H-M et al (2022) Injectable amnion hydrogelmediated delivery of adipose-derived stem cells for osteoarthritis treatment. Proc Natl Acad Sci 119(4):e2120968119 14. Wu P-H, Chung H-Y, Wang J-H et al (2016) Amniotic membrane and adipose-derived stem cell co-culture system enhances bone regeneration in a rat periodontal defect model. J Formos Med Assoc 115(3):186–194 15. Khorramirouz R, Kameli SM, Fendereski K et al (2019) Evaluating the efficacy of tissue-

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engineered human amniotic membrane in the treatment of myocardial infarction. Regen Med 14(2):113–126 16. Sa´nchez-Sa´nchez R, BREnA-MoLInA A, MARTı´nEz-Lo´PEz V et al (2015) Generation of two biological wound dressings as a potential delivery system of human adipose-derived mesenchymal stem cells. ASAIO J 61(6):718 17. Zhang T, Yam GH-F, Riau AK et al (2013) The effect of amniotic membrane de-epithelialization method on its biological properties and ability to promote limbal epithelial cell culture. Invest Ophthalmol Vis Sci 54(4):3072–3081 18. Zhang J, Liu Z, Li Y et al (2020) FGF-2induced human amniotic mesenchymal stem cells seeded on a human acellular amniotic membrane scaffold accelerated tendon-tobone healing in a rabbit extra-articular model. Stem Cells Int 2020:1–14 19. Davis GE, Blaker SN, Engvall E et al (1987) Human amnion membrane serves as a substratum for growing axons in vitro and in vivo. Science 236(4805):1106–1109 20. Brown JJ, Papaioannou VEJD (1993) Ontogeny of hyaluronan secretion during early mouse development. Development 117(2):483–492 21. Kiernan JJMT (1999) Strategies for preventing detachment of sections from glass slides. Microscopy Today 7(6):20–24 22. Mohiuddin OA, O’Donnell BT, Poche JN et al (2019) Human adipose-derived hydrogel characterization based on in vitro ASC biocompatibility and differentiation. Stem Cells Int 2019: 1–13 23. Khalil S, El-Badri N, El-Mokhtaar M et al (2016) A cost-effective method to assemble biomimetic 3D cell culture platforms. PLoS One 11(12):e0167116 24. Motamed M, Sadr Z, Valojerdi M et al (2017) Tissue engineered human amniotic membrane

application in mouse ovarian follicular culture. Ann Biomed Eng 45:1664–1675 25. Jin CZ, Park SR, Choi BH et al (2007) Human amniotic membrane as a delivery matrix for articular cartilage repair. Tissue Eng 13(4): 693–702 26. Sangwan VS, Vemuganti GK, Singh S et al (2003) Successful reconstruction of damaged ocular outer surface in humans using limbal and conjuctival stem cell culture methods. Biosci Rep 23(4):169–174 27. Crapo PM, Gilbert TW, Badylak SFJB (2011) An overview of tissue and whole organ decellularization processes. Biomaterials 32(12): 3233–3243 28. Lakkireddy C, Vishwakarma SK, Raju N et al (2021) Fabrication of decellularized amnion and chorion scaffolds to develop bioengineered cell-laden constructs. Cell Mol Bioeng 15:1– 14 29. Oh JM, Gangadaran P, Rajendran RL et al (2022) Different expression of thyroid-specific proteins in thyroid cancer cells between 2-dimensional (2D) and 3-dimensional (3D) culture environment. Cell 11(22):3559 30. Janjic´ K, Lilaj B, Moritz A et al (2018) Formation of spheroids by dental pulp cells in the presence of hypoxia and hypoxia mimetic agents. Int Endod J 51:e146–e156 31. Anderson DE, Markway BD, Weekes KJ et al (2018) Physioxia promotes the articular chondrocyte-like phenotype in human chondroprogenitor-derived self-organized tissue. Tissue Eng A 24(3–4):264–274 32. Frontini-Lo´pez YR, Rivera L, Aldana AA et al (2023) Human adipose mesenchymal stromal cells growing into PCL-nHA electrospun scaffolds undergo hypoxia adaptive ultrastructural changes. Biotechnol J 18:2200413

Chapter 15 Robust Generation of ASC Spheroids for Use as 3D Cultures and in Bioprinted Tissue Models Martin Watzling, Hannes Horder, Petra Bauer-Kreisel, and Torsten Blunk Abstract Three-dimensional (3D) cell culture techniques have become a valuable tool to mimic the complex interactions of cells with each other and their surrounding extracellular matrix as they occur in vivo. In this respect, 3D spheroids are widely acknowledged as self-assembled cellular aggregates that can be generated from a variety of cell types without the need for exogenous material while being highly reproducible, easy to handle, and cost-effective. Furthermore, due to their capacity to be developed into microtissues, spheroids represent potential building blocks for various tissue engineering applications, including 3D bioprinting approaches for tissue model development. Adipose-derived stromal/stem cells (ASCs), due to their ease of isolation, multipotent nature, and secretory capacity, represent an attractive cell source employed in numerous tissue engineering studies and other cell-based therapy approaches. In this chapter, we describe two procedures for robust spheroid generation, namely the liquid overlay technique, either using agarose-coated 96-well plates or employing agarose-cast micromolds. Furthermore, we show, in principle, the generation of ASC spheroids with subsequent adipogenic differentiation and the spheroid generation using adipogenically differentiated ASCs, as well as the morphological characterization of generated spheroids. Key words 3D cell culture, Adipocytes, Adipose-derived stromal/stem cells, Bioprinting, Co-culture, Spheroids, Tissue engineering

1

Introduction In human tissues, cells are densely arranged in a complex threedimensional (3D) architecture and receive biochemical and mechanical signals from neighboring cells and the extracellular matrix (ECM) that determine cell function. To mimic the complex interaction of cells with each other and their surrounding ECM in vitro, 3D culture models are more conducive than conventional two-dimensional (2D) cultures. They thus can provide more physiologically relevant readouts [1–4]. 3D spheroids as self-assembled cellular aggregates represent a valuable tool in this context. They have been widely acknowledged for their ability to reconstruct cell–

Jeffrey M. Gimble et al. (eds.), Adipose-Derived Stem Cells: Methods and Protocols, Methods in Molecular Biology, vol. 2783, https://doi.org/10.1007/978-1-0716-3762-3_15, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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cell and cell–ECM interactions. Spheroids can be generated from a variety of cell types and can be formed without requiring exogenous material as a matrix or scaffolding while being highly reproducible, easy to handle, and cost-effective [5, 6]. Spheroids can be made from single cell types, but they also enable relatively easy implementation of co-cultures with direct cell–cell interaction [7]. Going one step further, due to their versatility and capacity to be developed into microtissues, spheroids represent potential building blocks for various tissue engineering applications [8–10], including 3D bioprinting approaches for tissue model development [11]. In the human body, white adipose tissue is used for energy storage, and it also represents a major endocrine organ, decisively influencing many physiological and pathophysiological processes [12–15]. It can be found, for example, as subcutaneous or visceral fat, and is relatively easily accessible. Adipose-derived stromal/stem cells (ASCs) can be conveniently isolated from lipoaspirates with high yield. Due to their multipotent nature and secretory capacity, ASCs are an attractive cell source employed in numerous tissue engineering studies and other cell-based therapy approaches [16– 20]. Culturing and delivering ASCs as multicellular spheroids have been demonstrated to increase their regenerative potential in terms of superior differentiation ability and elevated secretion of angiogenic and anti-inflammatory factors. In previous studies, we have utilized ASC spheroids for the investigation of adipogenic differentiation and generation of adipose microtissues, generation of co-cultures with direct contact to other cell types, and as building blocks in bioprinted tissue models [9, 11, 21, 22]. ASC spheroids were demonstrated to exhibit an enhanced adipogenic differentiation compared with 2D cultured cells. Spheroids required a distinctly shorter adipogenic stimulus to sustain adipogenesis, which was demonstrated based on analysis of triglyceride content and adipogenic marker gene expression. Furthermore, while undifferentiated spheroids displayed a stromal ECM pattern, in the course of adipogenesis, a dynamic shift in the ECM composition toward an adipogenic phenotype was observed, associated with enhanced expression of, for example, laminin and collagen IV, similar to native fat. Thus, culturing ASCs as spheroids can enhance their adipogenic capacity and generate adipose-like microtissues [21]. In breast cancer, ASCs, and adipocytes, as components of the mammary fat pad, come into close contact with tumor cells and are increasingly acknowledged to promote cancer development and progression [23, 24]. To mimic direct cell–cell interactions between tumor and adjacent stromal cells, we have developed a 3D co-spheroid model consisting of breast cancer cells and either ASCs or adipocytes. Direct heterotypic cell–cell contact in this model promoted the expression of a specific cytokine and one of

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its receptors (CCL5/CCR1), which, in turn, enhanced the migration of triple-negative MDA-MB-231 breast cancer cells [22]. The identification of tumor-specific markers that are upregulated upon cell–cell contact with neighboring stromal cells, as demonstrated in the 3D co-culture spheroids, may represent a promising strategy to find new targets for the diagnosis and treatment of invasive breast cancer. Besides being 3D cultures on their own, when it comes to tissue engineering applications, including advanced 3D bioprinting, spheroids can act as building blocks to generate more complex constructs. In extrusion-based bioprinting, a major challenge is to ensure homogeneous dispersion of spheroids without sedimentation in the printer cartridge and in the constructs after printing, which can be achieved by adding viscosity enhancers to the bioink. Furthermore, the diameter of the printing nozzle needs to be adjusted to the spheroid size to avoid detrimental shear stress [11]. Utilizing ASC spheroids, recently, we have developed a 3D-bioprinted breast cancer-adipose tissue model. The printability of ASC spheroids and the adipogenic differentiation within printed spheroids into adipose microtissues were demonstrated. Subsequently, a breast cancer cell compartment was printed onto the adipose tissue constructs, and in these co-cultures, a cancer cellinduced reduction of the lipid content and profibrotic remodeling of the ECM within the adipose tissues were observed [11]. Thus, 3D-printed disease models utilizing ASC spheroids can recapitulate important aspects of the complex cell–cell and cell–ECM interactions within the tumor–stroma microenvironment. In this chapter, we describe two procedures for robust spheroid generation, namely the liquid overlay technique, either using agarose-coated 96-well plates or employing agarose-cast micromolds. Furthermore, we show, in principle, the generation of ASC spheroids with subsequent adipogenic differentiation and the spheroid generation using adipogenically differentiated ASCs, as well as the morphological characterization of generated spheroids.

2

Materials

2.1 Coating of 96Well Plates

1. Agarose (for molecular biology). 2. Dulbecco’s phosphate-buffered saline (DPBS). 3. 1% penicillin–streptomycin (10,000 10,000 μg/mL streptomycin) in DPBS.

U/mL

penicillin;

4. Sterile 96-well plates (see Note 1). 5. Dulbecco’s Modified Eagle’s Medium/Ham’s F12 mixture (1: 1; DMEM/F12) with 10% fetal bovine serum (FBS) and 1%

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penicillin–streptomycin (10,000 U/mL penicillin; 10,000 μg/ mL streptomycin), referred to as basal medium. 2.2 Casting of Micromolds

1. Silicone forms (Fig. 2; available from Microtissues®, Providence, RI, USA; as described in [11, 22]). 2. 70% ethanol (EtOH) in ddH2O. 3. Agarose (for molecular biology). 4. DPBS. 5. Dulbecco’s Modified Eagle’s Medium/Ham’s F12 mixture (1: 1; DMEM/F12) with 10% FBS and 1% penicillin–streptomycin (10,000 U/mL penicillin; 10,000 μg/mL streptomycin), referred to as basal medium.

2.3 Expansion and Seeding of Cells

1. ASCs (see Note 2). 2. DPBS. 3. Trypsin-EDTA (0.25%). 4. Dulbecco’s Modified Eagle’s Medium/Ham’s F12 mixture (1: 1; DMEM/F12) with 10% FBS and 1% penicillin–streptomycin (10,000 U/mL penicillin; 10,000 μg/mL streptomycin), referred to as basal medium. 5. Dulbecco’s Modified Eagle’s Medium/Ham’s F12 mixture (1: 1; DMEM/F12) with 10% FBS, 1% penicillin–streptomycin (10,000 U/mL penicillin; 10,000 μg/mL streptomycin), and 3 ng/mL bFGF, referred to as growth medium. 6. Coated plates or cast micromolds (see Subheadings 2.1 and 2.2).

3

Methods Carry out all procedures under sterile conditions unless otherwise specified.

3.1 Expansion of ASCs

1. ASCs are thawed and seeded in growth medium in 2D conditions in conventional cell culture flasks. 2. Growth medium is exchanged every other day. 3. At ~85% confluence, cells are detached and seeded as described in Subheadings 3.3, 3.5, and 3.7.

3.2 Coating 96-Well Plates

1. Prepare 1.5% agarose solution (w/v) in DPBS (see Note 3) and heat in a microwave until agarose is fully dissolved. 2. Dispense 50 μL of agarose into each inner well while leaving the outer row of wells empty (see Fig. 1; see Note 4).

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Fig. 1 Layout of agarose-coated 96-well plate with spheroids in inner wells. Floating adipose-derived stromal/stem cell (ASC) spheroid consisting of 2500 cells (bottom right, scale bar = 200 μm)

3. Allow coated plates to dry under a laminar flow hood for 15–30 min. 4. For equilibration of agarose, add 100 μL of basal medium to each coated well and 100 μL of 1% penicillin–streptomycin in DPBS to each outer well (see Note 5). 5. Plates can be stored for up to 2 weeks and wrapped in Parafilm at 4 °C (see Note 6). 3.3 Spheroid Generation in 96-Well Plates

Spheroids generated in 96-well plates are highly homogeneous with a very narrow size distribution. However, the method is relatively labor-intensive for large experiments since only 60 spheroids can be simultaneously cultivated in one plate and, thus, multiple well plates have to be cultured when larger amounts of spheroids are required. Spheroid size is virtually not restricted in this culture technique. Furthermore, the harvesting procedure is gentler compared to harvesting from micromolds (see below). Spheroids generated in coated 96-well plates are thus best suited for analyses requiring uniform shape and size, for example, for migration/invasion assays or histochemical analyses (Fig. 1). 1. Rinse ASCs with DPBS to remove residual FBS. 2. Trypsinize cells for 5 min or until all adherent cells are detached. 3. Stop the reaction by adding at least double the amount of basal medium to the cell–trypsin mixture.

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4. Centrifuge for 10 min at 300 × g, aspirate supernatant, and add fresh basal medium. 5. Count cells and calculate cell concentration (see Note 7). 6. Dispense 50 μL of cell suspension into each coated well. 7. Add 50 μL basal medium to each coated well (see Note 8). 8. Let cells settle for 15 min. 9. Centrifuge for 5 min at 100 × g (see Note 9). 10. Cells are cultured according to experimental conditions (see Note 10). 11. Medium is exchanged according to experimental needs. For this purpose, aspirate 50 μL of medium and add 50 μL of fresh medium (see Note 8). 12. After culture, spheroids are harvested by aspirating medium together with the spheroid and collecting them (see Note 11) for downstream analyses. This is done for each well and spheroid individually. 3.4 Casting Micromolds

1. Incubate silicone forms, which are subsequently used as negatives for the casting of agarose micromolds, in 70% EtOH overnight or autoclave prior to micromold casting. 2. Dry sterile silicone forms under a laminar flow hood until all liquid is evaporated. 3. Prepare 2% agarose solution (w/v) in DPBS and heat in a microwave until agarose is fully dissolved (see Note 3). 4. Dispense 500 μL of liquid agarose solution into each silicone form. 5. Allow cast micromolds to dry for at least 15 min. 6. Carefully remove agarose micromolds from silicone forms and place them in a sterile 12-well plate, 1 micromold per well (see Note 12). 7. For equilibration of micromolds, add 2 mL of basal medium to each well containing a micromold (see Note 5). 8. After a sufficient amount of micromolds has been cast, centrifuge plates for 3 min at 500 × g to remove air pockets from micromolds. 9. Micromolds can be stored for up to 3 weeks under experimental conditions (e.g., incubator) prior to the experiment.

3.5 Spheroid Generation in Micromolds

Spheroids in micromolds can be produced in large quantities (>3000 spheroids per plate; a 50-fold increase compared to generation in 96-well plates) and are, therefore, well suited for highthroughput applications. They exhibit minor size variations, and the micromold harvesting procedure is associated with a slightly

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Fig. 2 Spheroid formation in micromolds. Silicone forms (top left) are used to cast agarose micromolds (top right, scale bar = 1 cm). Cells are seeded in micromolds, resulting in spheroids after a formation time of 48 h (bottom left, scale bar = 500 μm). Spheroids of varying size can be produced, depending on seeded cell number (bottom right). (Adapted from Horder et al. and Watzling et al. [11, 22])

higher shear stress compared to production in 96-well plates; nevertheless, this method very reproducibly yields homogeneous, round spheroids. Therefore, micromold-generated spheroids are a valuable tool for assays requiring larger sample volumes, for example, quantitative real-time polymerase chain reaction, and when building blocks for larger constructs are required, for example, for bioprinting purposes. We routinely use micromolds with a 16 × 16 array, that is, 256 slots per micromold, resulting in 256 spheroids. In a 12-well plate, 1 micromold is placed in each well, resulting in a total of 3072 spheroids per plate (see Note 13) (Fig. 2).

1. Rinse ASCs with DPBS to remove residual FBS. 2. Trypsinize cells for 5 min or until all adherent cells are detached. 3. Stop the reaction by adding at least double the amount of basal medium to the cell–trypsin mixture.

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4. Centrifuge for 10 min at 300 × g, aspirate supernatant, and add fresh basal medium. 5. Count cells and calculate cell concentration (see Note 14). 6. Dispense 190 μL of cell suspension into each well. 7. Let cells sink into slots for at least 15 min. 8. Add 2 mL of medium to each well (see Note 15). 9. Centrifuge for 5 min at 300 × g (see Note 9). 10. Cells are cultured according to experimental conditions (see Note 10). 11. Medium is exchanged according to experimental needs. For this purpose, aspirate 2 mL of medium and add 2 mL of fresh medium (see Note 15). 12. After culture, spheroids are released from micromolds by turning micromolds upside-down in their wells filled with medium and centrifuging plates gently at 50 × g for 3 min (see Note 16). 13. Spheroids are aspirated and collected (see Note 17) for downstream analyses. 3.6 Generation of ASC Spheroids and Subsequent Adipogenic Differentiation

1. Follow protocol 3.3 (up to step 9) or 3.5 (up to step 9), depending on the method of choice. 2. After seeding, cells are left to form spheroids for 48 h. 3. When ASCs have formed spheroids, they are exposed to an adipogenic induction medium (Fig. 3; for a detailed description of induction and analyses of ASC differentiation in spheroids, see [21]). 4. Exchange induction medium every other day for 14 days. 5. Harvest adipogenically differentiated spheroids (adipose microtissues) as described under Subheading 3.3 (step 12) or 3.5 (step 12), depending on the method of choice.

3.7 Spheroid Generation Using Adipogenically Differentiated ASCs

1. At 80% confluence, ASCs are exposed to an adipogenic induction medium (Fig. 3; for a detailed description of induction and analyses of ASC differentiation, see [21]). 2. Exchange induction medium every other day for 14 days. 3. Rinse confluent adipocytes with DPBS to remove residual FBS. 4. Trypsinize until all adherent cells are detached (see Note 18). 5. Stop the reaction by adding at least double the amount of basal medium to the cell–trypsin mixture. 6. Centrifuge for 10 min at 300 × g, aspirate supernatant, and add fresh basal medium. 7. Continue by following protocol 3.3 (from step 5) or 3.5 (from step 5), depending on the method of choice (see Note 19).

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Fig. 3 Generation of spheroids consisting of adipogenically differentiated adipose-derived stromal/stem cells (ASCs) (adipose microtissues). ASCs can either be adipogenically differentiated when ASCs have already formed spheroids (top) or prior to spheroid formation (bottom). (Adapted from Watzling et al. [22])

3.8 Morphological Characterization of Generated Spheroids

1. Acquire images of spheroids inside coated plates or micromolds at desired time points to analyze spheroid morphology. 2. Spheroid diameter can either be determined by employed imaging software, e.g., CellSens or ImageJ/FIJI. 3. For this purpose, open the image in ImageJ. 4. Convert to an 8-bit image (see Note 20). 5. Select the appropriate autothreshold able to detect the spheroid within the image. This threshold should not be changed for different conditions or images within a single experiment to avoid arbitrary variations. 6. Analyze particles according to previously set measurements (e.g., shape descriptors like roundness or area). Smaller particles can be excluded from particle analysis by setting size parameters in the case of cell debris. 7. Values can be extracted and further processed for graphical processing and statistical analyses (see Fig. 4).

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Notes 1. Routinely, we use U-shaped 96-well plates. However, coating of U-, V-, or R-shaped plates results in similarly shaped agarose surfaces. Nonadherent or ultra-low attachment plates are not required.

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Fig. 4 Morphological characterization of spheroids consisting of 1000 cells, either undifferentiated adiposederived stromal/stem cells (ASCs) or adipogenically differentiated ASCs (adipocytes, AD). (a) Compaction of spheroids over 48 h after seeding in micromolds. (b) Roundness of spheroids made from either ASCs or adipocytes was assessed using ImageJ (as described in Subheading 3.8). (c) Brightfield images of spheroids for morphological examination. Scale bar = 100 μm. (Adapted from Watzling et al. [22])

2. ASCs can be freshly isolated from adipose tissue or commercially acquired. Routinely, we use ASCs purchased from Lonza (Basel, Switzerland) in passages 4–6. 3. It is recommended only to prepare small volumes of agarose solution to avoid extensive dissolution times during boiling (~ 80 mL). Preparing agarose solution does not require sterile conditions before sterilization by microwaving. For this purpose, initially bring agarose solution to boil briefly and incubate for 3–5 min, mixing regularly. Then heat it again to fully dissolve it with occasional breaks for mixing. 4. The outer row of the 96-well plate would be prone to stronger evaporation of liquid than the 60 inner wells. Therefore, it is not used in the process of spheroid generation to avoid irregularities. 5. It should be considered that agarose might retain components from media (e.g., FBS components). Therefore, the equilibration medium should correspond to the medium employed in subsequent experiments. 6. If plates are used immediately or on the next day, plates should be stored in experimental conditions (e.g., incubator). 7. Per well, 50 μL of cell suspension should be dispensed. This equals one spheroid. Thus, the cell number per 50 μL should be adjusted according to the desired cell number per spheroid. For one plate with 60 coated wells, 3 mL of cell suspension is needed. It is recommended to account for possible slight

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pipetting errors by preparing a greater volume. To generate co-culture spheroids, cells should be mixed prior to seeding to ensure homogeneous cell distribution throughout the spheroid. 8. Some experimental setups require additives to the medium. When adding the final 50 μL of media, including additives, to 50 μL of cell suspension, additive concentrations should be adjusted accordingly to meet the desired final concentration in the total volume of 100 μL. 9. Depending on cell type, the ability to form spheroids varies. If cells have been shown to form coherent and round spheroids on their own, as is the case for ASCs, this step is not required but might be helpful to improve spheroid formation and shape. 10. If necessary, plates can be placed on an orbital shaker (e.g., inside an incubator), gently rotating cells at ~40 r/min to further improve spheroid formation during the initial culture. 11. ASC spheroids with up to 5000 cells can be harvested with 100 μL pipette tips without damage. For larger spheroids, 1000 μL pipette tips are recommended. (This may vary for other cell types and resulting spheroid size.) Harvest is additionally facilitated by placing plates on an illuminated surface to locate spheroids more easily. 12. Cell culture-treated plates and suspension culture plates are both suited for micromold culture since the seeded cells only come into contact with the agarose micromolds (not with the underlying plate material). 13. In the micromolds with a 16 × 16 array, spheroid size is limited due to the spatial capacity of the slots to ~10,000 cells per slot, depending on cell type. For experiments in which spheroids with higher cell numbers are required, different silicone forms with 9 × 9 arrays with more spacious slots are also available, following identical protocols for casting agarose micromolds and subsequent spheroid generation. 14. Per micromold, 190 μL of cell suspension should be dispensed. This leads to the generation of 256 spheroids (when the molds with the 16 × 16 arrays are used). The cell number per 190 μL should be adjusted according to the desired cell number per spheroid. For one plate with 12 micromolds (3072 spheroids), 2280 μL (12 × 190 μL) of the cell suspension is needed. It is recommended to account for possible slight pipetting errors by preparing a greater volume. To generate co-culture spheroids, cells should be mixed prior to seeding to ensure homogeneous cell distribution throughout the spheroid. 15. Some experimental setups require additives to the medium. When adding the 2 mL of medium, including additives, to

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micromolds containing 190 μL of cell suspension after cells have settled into slots, additive concentrations should be adjusted accordingly to meet the desired final concentration in the total volume of 2190 μL. 16. Centrifuging might damage spheroids that are less cohesive than ASC spheroids. If the shape is relevant for downstream analyses (e.g., migration/invasion assays or histological procedures), it is recommended to turn micromolds upside-down and gently tap the plate to allow spheroids to fall from micromolds into the medium-filled well or gently flush spheroids out using a 1000 μL pipette, instead of centrifuging. 17. Due to the large amount of spheroids harvested together, it is recommended to use a 1000 μL pipette to avoid shear stress and damage. 18. Detachment of adipocytes may take longer than for ASCs. To avoid loss of cells, it is recommended to facilitate detachment by gently patting the culture flask and increasing incubation times of trypsin until a sufficient number of cells are detached. 19. Due to adipocyte morphology, a gentler approach is advised, for example, by reducing resuspension to only necessary steps and avoiding too vigorous pipetting. Thereby, damage to intracellular lipid droplets can be minimized. 20. Depending on the quality of the image or imaging equipment, appropriate filters can be selected to facilitate analyses (e.g., Gaussian blur).

Acknowledgments This work was supported by the Bayerische Forschungsstiftung (BFS, Bavarian Research Foundation), grant number AZ-136518, FORTiTher (subproject TP2WP2), and the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation), project number 326998133, TRR 225 (subproject C02). References 1. Stricker S, Knaus P, Simon HG (2017) Putting cells into context. Front Cell Dev Biol 5:32. https://doi.org/10.3389/fcell.2017.00032 2. Ravi M, Paramesh V, Kaviya SR et al (2015) 3D cell culture systems: advantages and applications. J Cell Physiol 230(1):16–26. https:// doi.org/10.1002/jcp.24683 3. Bissell MJ (2017) Goodbye flat biology—time for the 3rd and the 4th dimensions. J Cell Sci 130(1):3–5. https://doi.org/10.1242/jcs. 200550

4. Jensen C, Teng Y (2020) Is it time to start transitioning from 2D to 3D cell culture? Front Mol Biosci 7:33. https://doi.org/10. 3389/fmolb.2020.00033 5. Hoarau-Vechot J, Rafii A, Touboul C et al (2018) Halfway between 2D and animal models: are 3D cultures the ideal tool to study cancer-microenvironment interactions? Int J Mol Sci 19(1):181. https://doi.org/10. 3390/ijms19010181

Robust Generation of ASC Spheroids 6. Achilli T-M, Meyer J, Morgan JR (2012) Advances in the formation, use and understanding of multi-cellular spheroids. Expert Opin Biol Ther 12(10):1347–1360. https:// doi.org/10.1517/14712598.2012.707181 7. Xin X, Yang H, Zhang F et al (2019) 3D cell coculture tumor model: a promising approach for future cancer drug discovery. Process Biochem 78:148–160. https://doi.org/10.1016/ j.procbio.2018.12.028 8. Moldovan NI, Hibino N, Nakayama K (2017) Principles of the Kenzan method for robotic cell spheroid-based three-dimensional bioprinting. Tissue Eng Part B Rev 23(3): 237–244. https://doi.org/10.1089/ten. TEB.2016.0322 9. McMaster R, Hoefner C, Hrynevich A et al (2019) Tailored melt electrowritten scaffolds for the generation of sheet-like tissue constructs from multicellular spheroids. Adv Healthc Mater 8(7):e1801326. https://doi. org/10.1002/adhm.201801326 10. Mekhileri NV, Lim KS, Brown GCJ et al (2018) Automated 3D bioassembly of microtissues for biofabrication of hybrid tissue engineered constructs. Biofabrication 10(2): 024103. https://doi.org/10.1088/17585090/aa9ef1 11. Horder H, Guaza Lasheras M, Grummel N et al (2021) Bioprinting and differentiation of adipose-derived stromal cell spheroids for a 3D breast cancer-adipose tissue model. Cells 10(4):803. https://doi.org/10.3390/ cells10040803 12. Guerre-Millo M (2002) Adipose tissue hormones. J Endocrinol Investig 25(10): 8 5 5 – 8 6 1 . h t t p s : // d o i . o r g / 1 0 . 1 0 0 7 / BF03344048 13. Trayhurn P, Beattie JH (2001) Physiological role of adipose tissue: white adipose tissue as an endocrine and secretory organ. Proc Nutr Soc 60(3):329–339. https://doi.org/10. 1079/pns200194 14. Sethi JK, Vidal-Puig AJ (2007) Thematic review series: adipocyte biology. Adipose tissue function and plasticity orchestrate nutritional adaptation. J Lipid Res 48(6):1253–1262. https://doi.org/10.1194/jlr.R700005JLR200 15. Ritter A, Kreis NN, Hoock SC et al (2022) Adipose tissue-derived mesenchymal stromal/ stem cells, obesity and the tumor microenvironment of breast cancer. Cancers (Basel)

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Chapter 16 Generating and Characterizing Adipose Spheroids from Adipose-Derived Stromal/Stem Cells Charles Amurgis, W. Vincent Nerone, and Lauren Kokai Abstract Advances in technology and automation over the past several decades have made it feasible to perform highthroughput compound screening with cell spheroids, a valuable approach for drug discovery. It is entirely feasible to generate multiple 384-well plates containing adipose spheroids from cryopreserved, singledonor, adipose stem cells, thus incorporating genetic diversity into the discovery stages of research. In this protocol, we describe our method for isolating primary human adipose stem cells and synthesizing cell spheroids comprised of mature adipocytes and stromal cells. Also included are representative outcome measurements useful for characterizing adipocyte metabolism and health. Wherever possible, we describe technologies that can be used to automate characterization and increase throughput. Key words Adipose tissue, Cell spheroids, Three-dimensional cell culture, Drug testing, Adiposederived stromal/stem cells, Cell spheroid characterization

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Introduction While in vitro differentiation of immortalized cell lines or primary human cells in a two-dimensional (2D) environment is the most commonly used approach for studying adipocytes due to a wealth of comparative literature, cost, and ease of use, inherent limitations exist in the simplicity of the method. For example, 2D models lack complete unilocular adipocyte differentiation, matrix–cell interactions, and cell-to-cell spatial organization. Furthermore, though 2D adipogenesis replicates acute phenotypes directed by transcription factors, the post-translational modifications that define mature adipocyte gene regulation in vivo are not present in this environment. For many studies, this lack of physiological relevance is acceptable; however, for applications such as disease modeling and drug screening, 2D in vitro platforms are insufficient. To overcome these challenges, while maintaining logistical flexibility of using cryopreserved cells, we generated adipose tissue de novo

Jeffrey M. Gimble et al. (eds.), Adipose-Derived Stem Cells: Methods and Protocols, Methods in Molecular Biology, vol. 2783, https://doi.org/10.1007/978-1-0716-3762-3_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2024

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by creating adipospheres comprised of primary human stem cells differentiated toward adipocytes. Compared to 2D cultures, adipose organoids can bridge the gap between developmental in vitro and in vivo models by providing cell–cell interaction, improved unilocular lipid formation, and physiologically mature gene expression profiles that improve in vitro tissue recapitulation. There are several benefits to utilizing adipospheres created from human cells as opposed to using in vivo animal models. Animal models face limitations in their predictive validity of the human adipose microenvironment as they possess unique metabolic signatures and distinctive interspecies genetic expression compared to humans. As a result, human-derived cells provide a more accurate representation of human disease outcomes. For instance, despite being the most common species for modeling obesity, rodents diverge from humans in overall lipogenic capacity due to an upregulation of SREBP-1c, a key transcriptional factor of fatty acid storage [1]. Therefore, we have elected to use human cells to better mimic disease outcomes relevant to human health. During spheroid synthesis and completion of the adipogenic differentiation protocol, there are several methods to nondestructively track cell viability and differentiation efficiency. In our studies, we elected to track viability by grossly observing morphological changes with microscopy, measuring the model’s metabolic functionality via glucose consumption and determining the presence of free lactate dehydrogenase (LDH) in the conditioned media. Additional methods for tracking cell metabolic function include using phenol-red-containing media and monitoring for color change, measuring precise media acidification with a microneedle pH meter or measuring oxygen consumption rate, metabolic fuel preference, or other metabolic tests with more sophisticated technologies such as the Agilent Seahorse Bioanalyzer and associated test kits [2]. In many instances, the described methods for measuring viability and metabolism are easily adapted for compatibility with drug screening experiments. To ensure successful adipogenesis has occurred within adipospheres and to measure patient-specific baseline efficiencies, spheroids may be characterized with microscopy, including nondestructive and higher throughput methods utilizing 4′,6-diamidino-2-phenylindole (DAPI), BODIPY, and phalloidin [3]. However, due to the limited stain penetration of these threedimensional (3D) models, we also present an alternative method for sectioning and staining to examine inner structures with or without immunohistochemistry. To quantify genetic changes in cells, we also present robust methods for RNA extraction for downstream quantification with quantitative reverse transcriptase polymerase chain reaction. Finally, to assess adiposphere metabolism, such as the ability to store and release energy in the form of triglycerides, we recommend

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measuring spheroid triglyceride content normalized to cell number, quantifying adipocyte size, and measuring isoproterenolmediated glycerol release. Triglycerides serve as the primary storage form of fatty acids in adipocytes, representing a key hallmark of adipogenic differentiation and maturation [4]. By quantifying triglyceride levels, the extent of adipocyte metabolic function can be compared. Triglyceride storage is closely related to adipocyte size, which is also a measure of maturity. Osmium tetroxide (OsO4) fixation of the spheroids, followed by urea-based digestion, provides a robust method for quantifying adipocyte size distributions across experimental test groups using either microscopic image analysis or more throughput methods with a Coulter counter [5, 6]. Finally, stimulating beta-adrenergic receptors on adipocytes with isoproterenol leads to activation of lipase enzymes, which catalyzes the hydrolysis of triglycerides stored within adipocytes [7–9]. As a result, glycerol and fatty acids are released into the surrounding medium and can be measured. These approaches are more labor intensive and less amenable to initial drug screening protocols; however, they do yield vital information regarding adipocyte health and function.

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Materials

2.1 Adipose-derived Stromal/Stem Cell (ASC) Isolation and Culture Expansion

1. Collagenase solution: 1 mg/mL collagenase type 2 in Hanks’ Balanced Salt Solution (HBSS) with 3.5% bovine serum albumin. Can be stored as prepared at -80 °C for years without loss of activity (see Notes 1–3). 2. Erythrocyte lysis buffer: 1000 mL deionized water, 8.23 g of 154 mM ammonium chloride (NH4Cl), 1.0 g of 10 mM potassium bicarbonate (KHCO3), and 36 mg of Open. . . and then select the image from the browser. Once the image is open, ensure that the western blot lanes are perfectly horizontal. If they are not, go to Image > Transform > Rotate. . . and rotate the image until the lanes are horizontal. 3. Next, box select the lanes that you want to analyze by using the rectangle selection tool in the main ImageJ window (Figs. 3 and 4). 4. Going to Analyze > Gels > Select First Lane will mark these lanes with a “1.” Then, go to Analyze > Gels > Plot Lanes. ImageJ will then generate a series of peaks based on the chemiluminescence image as illustrated in Fig. 5.

Fig. 2 The ImageJ main window

Fig. 3 Selecting the rectangle selection tool in the ImageJ main window

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Fig. 4 Chemiluminescence image of the western blot open in ImageJ

Fig. 5 The peaks produced by ImageJ’s interpretation of the chemiluminescence image of the western blot

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Fig. 6 Selecting the line drawing tool in the ImageJ main window

Fig. 7 Arrows point to the troughs that incompletely separate each peak and must be sealed with the line drawing tool

5. Now, select the line tool and ensure that each peak is completely separated from its neighbors. This process is shown in Figs. 6 and 7. 6. Once the peaks have been separated using the line tool, use the wand (tracing) tool and select each peak. Clicking on each peak once will suffice. A table listing the areas of each peak will be produced. Additionally, once all the peaks have been selected, you may go to Analyze > Gels > Label Peaks to generate area percentages for each peak (see Figs. 8 and 9). These areas can then be used to calculate the relative expression of each protein sample on the western blot.

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Fig. 8 The peaks after being selected with the wand tool

Fig. 9 The results window of ImageJ showing each peak’s area and percent area ig

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Notes 1. An effective scraping strategy is to hold the plate at about a 15–30° angle. Dip the scraper into the pooled reagent and scrape cells from the top of the plate into the pooled reagent at the bottom. Once scraping stops producing any new mucoid cell debris, the cells have been sufficiently detached and lysed. The entire scraping process should take no more than 90 s per dish. 2. Once all samples have been collected, the reagent–cell debris mixtures can be stored at -80 °C for later extraction; however,

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Table 8 Example RT-PCR reactions Sample

RNA concentration

1 μg RNA

cDNA superscript

Nuclease-free water

1

0.15 ng/μL

6.7 μL

4 μL

9.3 μL

2

0.07 ng/μL

14.3 μL

4 μL

1.7 μL

continuing the RNA extraction to completion without an intervening storage step is highly recommended. If using frozen lysed cell samples, allow the samples to completely thaw at room temperature. Otherwise, continue to the next step. 3. It is recommended to prepare new 1.5 mL tubes to collect supernatant during this centrifugation step. 4. Beware not to collect any of the mucoid waste or pink phenol. If the sample is disrupted, return all samples to the 1.5 mL tubes and repeat centrifugation to reseparate the organic and aqueous phases. 5. We note that adding glycogen has resulted in observable increases in RNA yield, and so we strongly recommend usage. 6. This will situate the RNA pellets on the hinge-side of each tube and allow for easier identification of the RNA pellets as well as decrease the likelihood of accidentally aspirating an RNA pellet. 7. For example, after evaluating the concentrations of hypothetical RNA samples 1 and 2 below, it is determined that we can use 1 μg of each sample. 6.7 μL of sample 1 RNA will be added to sample 1’s RT-PCR, and 14.3 μL of sample 2 RNA will be added to sample 2’s RT-PCR. Refer to Table 8. Note that if the concentration of RNA in a sample is lower 0.0625 μg/μL, 16 μL of RNA sample will not be enough RNA solution to add 1 μg of RNA to the RT-PCR mixture. In this case, calculate the amount of RNA in 16 uL of the lowest concentration sample and use that same quantity of RNA for each subsequent sample. Then, calculate the mass of RNA that was added to that reaction and normalize all other volumes of RNA sample added to all other concurrent reactions based on that mass. This ensures that equal amounts of RNA are added to each concurrent RT-PCR. 8. We recommend primer pairs that generate product sizes approximately 100 bp in length. We have observed that higher-quality qPCR data were obtained from primer pairs that produce amplicons in that range. As such, these primers should also be used in endpoint PCR as it is often necessary to optimize primers before use in qPCR.

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9. These steps are sequentially repeated between 20 and 40 cycles. Total PCR product amplification is 2n, where n is the number of cycles of PCR. More PCR cycles increases amplicon detection sensitivity. Less PCR cycles may allow observation of differential gene expression at endpoint. 10. Primer annealing temperature (Ta) is ideally 5 °C below the primer melting temperature (Tm), which varies per primer based largely on the primer’s length and the number of hydrogen bonds needed to anneal. Annealing temperatures too low may encourage off-target annealing. Temperatures too high may discourage primer annealing altogether. Finding the best annealing temperature for a primer pair is an optimization process involving trial and error by running PCR at varying annealing temperatures. The best annealing temperature is that which produces clear, bright product bands of uniform amplicon size. 11. An agarose concentration of 2.0% is ideal for PCR product sizes between 50 bp and 2 kbp. 12. DNA, due to its negative phosphate groups, will always run to the anode (red electrode) under an electric current. 13. If running reactions in technical triplicate, three wells will be required per target gene per treatment group. Also consider that one target gene should be a housekeeping gene such as gapdh. With one housekeeping gene, one other target gene, and one control cDNA sample and one treatment cDNA sample, expect to use a minimum of 12 wells. Using a biological triplicate of cDNA samples in a single qPCR will expand well usage threefold up to 36 (one-third of the total on a single 96-well plate). This map should designate the control or treatment cDNA sample used and the target gene for that well. An example 96-well plate map is provided in Fig. 10. 14. Note that SYBR® Green may not be compatible with all qPCR equipment. Ensure equipment compatibility with SYBR® Green before use. If SYBR® Green is not compatible with available equipment, choose a different dye. 15. Note that programs for qPCR are faster than endpoint PCR programs. Choose primer pairs that produce products that are around 100 bp in length to account for shortened extension time. 16. Note that qPCR is highly sensitive to pipetting error. The following preparation steps have been designed around an effort to minimize pipetting error as much as possible. For this reason, please pay attention to the creation of two master mixes in tandem, called the “sample” master mix and “primer” master mix, respectively.

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Fig. 10 Example 96-well plate map for qPCR Table 9 Sample master mix example Sample master mix cDNA sample ®

6.4 μL

SYBR green qPCR supermix

32 μL

Nuclease-free water

19.2 μL

Total

57.6 μL

17. For example, if using two cDNA samples (a control group and an experimental group) and two gene targets (a housekeeping gene and another target gene), two sample master mixes (one for each cDNA sample) should be prepared according to Table 9. 18. Following the specific example above, four new primer master mixes would be made. These primer master mixes are prepared in 1.5 mL microcentrifuge tubes according to Table 10. Continuing the example, there would now be new primer master mixes for the control cDNA housekeeping gene, control cDNA target gene, experimental cDNA housekeeping

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Table 10 Primer master mix example Primer master mix Sample master mix

32 μL

100 μM forward primer

1.6 μL

100 μM reverse primer

1.6 μL

Total

35.2 μL

Table 11 qPCR master mix matrix Control cDNA sample Experimental cDNA sample Target gene

Primer master mix 1

Primer master mix 3

Housekeeping gene Primer master mix 2

Primer master mix 4

gene, and experimental cDNA target gene. This is illustrated in Table 11. 19. Three wells will contain 10 μL each of one primer master mix. Mix each 10 μL by pipetting up and down gently. Beware that overmixing may shear the cDNA. 20. Do not touch the adhesive film to prevent smudges. 21. There are many different points of failure for a qPCR. Since various comprehensive troubleshooting guides exist in other places, particularly on the websites of biotechnology manufacturers, a comprehensive troubleshooting guide for qPCR is not included here. 22. The standard data produced by qPCR equipment and software are in the form of Ct (cycle threshold) values. Ct values are abstract numerical representations of the cycle numbers in which qPCRs achieve a certain level of fluorescence that exceeds a predefined (usually machine or software default) threshold value. Because a qPCR involves precisely measuring the amount of cDNA included in a reaction, Ct is used to calculate the relative number of transcripts of a target gene present in the cDNA sample. This level of target gene expression is usually calculated via the ΔΔCt (double-delta Ct) method and presented as fold change. 23. We usually calculated the standard error of the mean of our 2-ΔΔCt values to represent our error bars. 24. We found that even for Mediator subunit MED31, a 13 kDa transcriptional co-activator, we were able to detect the presence

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of small and potentially underexpressed nuclear proteins without the extra steps involved to isolate nuclear protein fractions. 25. Note that maintaining the protein at a cold temperature during the extraction procedure is important for obtaining highquality samples for use in protein analysis protocols, such as the ones provided here. 26. Note that the standard curve will require 600 μL of Bradford buffer, and each protein sample will require another 100 μL of Bradford buffer, or 200 μL if assayed in technical duplicate. 27. For better results, prepare the protein samples in technical duplicates. 28. A sample layout is provided in Fig. 11. 29. An example standard curve is provided in Fig. 12. 30. Plugging in the slope (m) and y-intercept (b) from the graph yields the following equation: y þ 0:0012 =x 0:0709

ð7Þ

y-intercept equation with example values from Fig. 11 plugged in.

Fig. 11 Example 96-well plate map for the Bradford assay

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Fig. 12 Example standard curve scatterplot showing the y-intercept equation and coefficient of determination Table 12 Example protein sample preparations for SDS-PAGE Sample number

Concentration (mg/mL)

Protein sample (μL)

DI water (μL)

2× Laemmli buffer Total volume (μL) (μL)

1

1.78

12.58

7.42

20

40

2

2.44

9.18

10.82

20

40

3

1.12

20

0

20

40

The slope and y-intercept values are for demonstration purposes and will vary from assay to assay. In the case that the coefficient of determination (R2) deviates significantly from 1, the standard curve may need to be discarded and remade to ensure the accuracy of the Bradford assay. 31. An example is provided in Table 12. 32. This procedure does not contain instructions on the preparation of a polyacrylamide gel. We used precast polyacrylamide gels for our procedures, but handcast gels will also work. Detailed instructions for handcasting polyacrylamide gels are provided by biotechnology manufacturers. 33. Cooling the samples any longer than this may cause the solutions to become excessively viscous and interfere with pipetting.

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34. A plastic ice pack may also be added directly into the running buffer for additional cooling. Use of a Bio-Rad protein electrophoresis cell configured for only one or two gels allowed enough room for an ice pack to be placed into the running buffer without overflowing the cell. 35. For running larger proteins, allow an additional 30 min to 1 h for the proteins to properly resolve on the gel. 36. We have had success using semi-dry transfer with smaller proteins (