Stem Cell Renewal and Cell-Cell Communication: Methods and Protocols [1 ed.] 149392589X, 978-1-4939-2589-6, 978-1-4939-2590-2

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Methods in Molecular Biology 1212

Kursad Turksen Editor

Stem Cell Renewal and Cell-Cell Communication Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

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

.

Stem Cell Renewal and Cell-Cell Communication Methods and Protocols

Edited by

Kursad Turksen Ottawa Hospital Research Institute Ottawa, ON, Canada

Editor Kursad Turksen Ottawa Hospital Research Institute Ottawa, ON, Canada

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-2589-6 ISBN 978-1-4939-2590-2 (eBook) DOI 10.1007/978-1-4939-2590-2 Library of Congress Control Number: 2015939235 Springer New York Heidelberg Dordrecht London # Springer Science+Business Media New York 2015 This work is subject to copyright. All rights are reserved 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, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. Printed on acid-free paper Humana Press is a brand of Springer Springer Science+Business Media LLC New York is part of Springer Science+Business Media (www.springer.com)

Preface Two topics—cell–cell interactions and stem cell renewal—are now inexorably linked as we strive to understand the stem cell niche and its function. I have thus selected a number of representative protocols to provide readers with approaches for studying these complimentary aspects of stem cells. As always, I am grateful to the contributors to this volume, for their generosity in sharing their protocols and the numerous details that will help others use these methods to advance their own studies. This volume, as others that I have been involved with, would not have materialized without the strong support and encouragement of the Editor in Chief of the Methods in Molecular Biology series, Dr. John Walker. Similarly, I am grateful to the Executive Editor of Methods in Molecular Biology, Patrick Marton, for being a great cheerleader during the maturation of this volume. Finally, I would like to acknowledge the Editor of Methods in Molecular Biology, David Casey, for his tireless efforts to make the volume flawless. Thanks to all for their efforts in bringing this volume to fruition! Ottawa, ON, Canada

Kursad Turksen

v

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

v ix

Laboratory Control and Basic Culture Protocols for Stem Cell Self-Renewal . . . . . . Jeong Mook Lim, Yoon Mi Jo, and Ji Yeon Ahn Advanced Fixation for Transmission Electron Microscopy Unveils Special Extracellular Matrix Within the Renal Stem/Progenitor Cell Niche . . . . . . . Will W. Minuth and Lucia Denk Multiplex Immunoassays for Quantification of Cytokines, Growth Factors, and Other Proteins in Stem Cell Communication . . . . . . . . . . . . . . Ivona Valekova, Helena Kupcova Skalnikova, Karla Jarkovska, Jan Motlik, and Hana Kovarova Porous Membrane Culture Method for Expansion of Human Pluripotent Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jin-Su Kim, Seung-Taeh Hwang, and Soo-Hong Lee Tuning Differentiation Signals for Efficient Propagation and In Vitro Validation of Rat Embryonic Stem Cell Cultures. . . . . . . . . . . . . . . . . . . Stephen Meek, Linda Sutherland, and Tom Burdon Generation, Expansion, and Differentiation of Human Pluripotent Stem Cell (hPSC) Derived Neural Progenitor Cells (NPCs). . . . . . . . . . . . . . . . . . . . . David A. Brafman Isolation, Long-Term Expansion, and Differentiation of Murine Neural Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alexandra Bizy and Sacri R. Ferro´n Generation, Expansion, and Differentiation of Cardiovascular Progenitor Cells from Human Pluripotent Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . Nan Cao, He Liang, and Huang-Tian Yang A Practical Guide for the Isolation and Maintenance of Stem Cells from Tendon . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pauline Po Yee Lui A Human Colonic Crypt Culture System to Study Regulation of Stem Cell-Driven Tissue Renewal and Physiological Function . . . . . . . . . . . . . . . . Alyson Parris and Mark R. Williams Ca2+ Handling in Mouse Embryonic Stem Cell-Derived Cardiomyocytes. . . . . . . . . Wenjie Wei and Jianbo Yue Potential Application of Extracellular Vesicles of Human Adipose Tissue-Derived Mesenchymal Stem Cells in Alzheimer’s Disease Therapeutics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Takeshi Katsuda, Katsuyuki Oki, and Takahiro Ochiya Application of Fluid Mechanical Force to Embryonic Sources of Hemogenic Endothelium and Hematopoietic Stem Cells . . . . . . . . . . . . . . . . . . . . Nan Li, Miguel F. Diaz, and Pamela L. Wenzel

1

vii

21

39

65

73

87

103

113

127

141 163

171

183

viii

Contents

Electrophysiological Recordings from Neuroepithelial Stem Cells . . . . . . . . . . . . . . . Masayuki Yamashita In Vivo Stem Cell Transplantation Using Reduced Cell Numbers . . . . . . . . . . . . . . . Takeo W. Tsutsui

195

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

209

201

Contributors JI YEON AHN  Department of Agricultural Biotechnology, Seoul National University, Seoul, South Korea ALEXANDRA BIZY  Departamento de Biologı´a Celular, Universidad de Valencia, Valencia, Spain DAVID A. BRAFMAN  School of Biological and Health Systems Engineering, Arizona State University, Tempe, AZ, USA TOM BURDON  The Roslin Institute and R(D)VS, University of Edinburgh, Midlothian, UK NAN CAO  Key Laboratory of Stem Cell Biology, Institute of Health Sciences, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences (CAS), Shanghai, China; Shanghai Jiao Tong University School of Medicine, Shanghai, China LUCIA DENK  Molecular and Cellular Anatomy, University of Regensburg, Regensburg, Germany MIGUEL F. DIAZ  Department of Pediatric Surgery, Center for Stem Cell and Regenerative Medicine, Institute of Molecular Medicine, University of Texas Medical School, Houston, TX, USA SACRI R. FERRO´N  Departamento de Biologı´a Celular, Universidad de Valencia, Valencia, Spain SEUNG-TAEH HWANG  Department of Biomedical Science, CHA University, Gyunggi-do, South Korea KARLA JARKOVSKA  Institute of Animal Physiology and Genetics, Academy of Sciences of the Czech Republic, Libechov, Czech Republic YOON MI JO  Department of Agricultural Biotechnology, Seoul National University, Seoul, South Korea TAKESHI KATSUDA  Division of Molecular and Cellular Medicine, National Cancer Center Research Institute, Tokyo, Japan JIN-SU KIM  Department of Biomedical Science, CHA University, Gyunggi-do, South Korea HANA KOVAROVA  Institute of Animal Physiology and Genetics, Academy of Sciences of the Czech Republic, Libechov, Czech Republic SOO-HONG LEE  Department of Biomedical Science, CHA University, Gyunggi-do, South Korea NAN LI  Department of Pediatric Surgery, Center for Stem Cell and Regenerative Medicine, Institute of Molecular Medicine, University of Texas Medical School, Houston, TX, USA HE LIANG  Key Laboratory of Stem Cell Biology, Institute of Health Sciences, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences (CAS), Shanghai, China; Shanghai Jiao Tong University School of Medicine, Shanghai, China JEONG MOOK LIM  Department of Agricultural Biotechnology and Research Institute of Agriculture and Life Sciences, Seoul National University, Seoul, South Korea; GreenBio Research Institute, Seoul National University, Kangwon, South Korea PAULINE PO YEE LUI  Headquarter, Hospital Authority, Hong Kong, SAR, China

ix

x

Contributors

STEPHEN MEEK  The Roslin Institute and R(D)VS, University of Edinburgh, Midlothian, UK WILL W. MINUTH  Molecular and Cellular Anatomy, University of Regensburg, Regensburg, Germany JAN MOTLIK  Institute of Animal Physiology and Genetics, Academy of Sciences of the Czech Republic, Libechov, Czech Republic TAKAHIRO OCHIYA  Division of Molecular and Cellular Medicine, National Cancer Center Research Institute, Tokyo, Japan KATSUYUKI OKI  Research and Development Department, SEEMS Inc., Tokyo, Japan; Department of Pathology, Tokyo Medical University, Tokyo, Japan ALYSON PARRIS  School of Biological Sciences, University of East Anglia, Norwich, Norfolk, UK HELENA KUPCOVA SKALNIKOVA  Institute of Animal Physiology and Genetics, Academy of Sciences of the Czech Republic, Libechov, Czech Republic LINDA SUTHERLAND  The Roslin Institute and R(D)VS, University of Edinburgh, Midlothian, UK TAKEO W. TSUTSUI  Department of Pharmacology, School of Life Dentistry at Tokyo, The Nippon Dental University, Tokyo, Japan IVONA VALEKOVA  Institute of Animal Physiology and Genetics, Academy of Sciences of the Czech Republic, Libechov, Czech Republic WENJIE WEI  Department of Physiology, University of Hong Kong, Hong Kong, China PAMELA L. WENZEL  Children’s Regenerative Medicine Program, Department of Pediatric Surgery, University of Texas Medical School, Houston, TX, USA; Center for Stem Cell and Regenerative Medicine, The Brown Foundation Institute of Molecular Medicine, University of Texas Health Science Center at Houston, Houston, TX, USA MARK R. WILLIAMS  School of Biological Sciences, University of East Anglia, Norwich, Norfolk, UK MASAYUKI YAMASHITA  Centre for Medical Science, International University of Health and Welfare, Ohtawara, Japan HUANG-TIAN YANG  Key Laboratory of Stem Cell Biology, Institute of Health Sciences, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences (CAS), Shanghai, China; Shanghai Jiao Tong University School of Medicine, Shanghai, China JIANBO YUE  Department of Physiology, University of Hong Kong, Hong Kong, China

Methods in Molecular Biology (2015) 1212: 1–20 DOI 10.1007/7651_2014_176 © Springer Science+Business Media New York 2014 Published online: 15 February 2015

Laboratory Control and Basic Culture Protocols for Stem Cell Self-Renewal Jeong Mook Lim, Yoon Mi Jo, and Ji Yeon Ahn Abstract In this report, we introduce a standard protocol for stem cell self-renewal in vitro. Both fundamental and major procedures of stem cell manipulation, which are required for somatic cell coculture and self-renewal, are briefly described since they are important for stabilization and data normalization. In this chapter, information on the basic preparation of stem cell culture such as labware washing, equipment sanitization, microbe control, and mycoplasmosis prevention is provided. In addition, protocols for cell retrieval and preservation, proliferation assays, and basic manipulation techniques for the coculture of stem cells with somatic cells are described. Keywords: Embryonic stem cell (ESC), Embryonic/fetal fibroblasts, Coculture, Standard operation protocol (SOP), Cryopreservation, Proliferation, Mycoplasmosis, Labware and equipment, Subpassage

1

Introduction Stem cells can transform into various cells and create functional tissues. To initiate stem cell differentiation in vivo, various signals and environmental factors are coordinated. The mobilization of multipotent and pluripotent stem cells from embryonic and somatic tissues and their subsequent maintenance in vitro are important for utilizing stem cells for clinical purposes. In the early stages of stem cell research, stem cells were obtained from intact tissues such as inner cell masses of embryos and terminally differentiated somatic tissues. Recent developments in the field of basic research and biotechnology have allowed the artificial establishment of pluripotent stem cells. Genetic engineering of cells can be employed to obtain induced pluripotent stem cells, and many target genes have been manipulated alone and in combination (1–3). Direct cell reprogramming has been used to create functional cells and to avoid various disadvantages of genetic manipulation and isolation of pluripotent cells such as uncontrolled differentiation and limitations of genetic manipulation, thus allowing the isolation of clinical-grade stem cells. Several cells with cellular plasticity have been used to isolate functional cells, and

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Jeong Mook Lim et al.

cellular niches have been used to overcome limitations of stem cell engineering. Characterizing cell-to-cell interactions and manipulating cellular interactions are important for determining cell fate as well as for both cell reprogramming and self-renewal. The basic protocols for stem cell self-renewal are introduced in this guidebook, for which cell-to-cell interaction studies are mainly applied. Since the coculture of embryonic stem cells (ESCs) onto the monolayer of mouse fetal fibroblasts is a general method for stem cell self-renewal, the basic protocol and the preparation of fibroblast coculture are discussed. General methods for cell culture such as monitoring cell proliferation and preparing culture materials are introduced, and we explain how labware cleaning and sanitization, as well as the prevention of microbial contamination, is accomplished. Prevention and curation of mycoplasma infection is described in detail based on its importance in laboratory quality control. In addition, culturing different cell types may increase the cellular plasticity of somatic cells or mobilization of stem cells mixed with terminally differentiated somatic cells. Examples of coculture methods described herein, which is based on cell-to-cell interactions, will be helpful for the establishment of somatic stem cells.

2

Materials

2.1 Preparation for Aseptic Cell Culture

1. 7 detergent (Mpbio, Cat. No. 76-670-94, Rue Geiler de Kayserberg, Illkirch, France) (see Note 2)

2.1.1 Reagent (See Note 1)

2. 0.25 % Trypsin–EDTA (Gibco, Cat. No. 25200072, Grand Island, NY) (see Note 3). 3. 70 % Alcohol (see Note 4) 4. Antibiotic-antimycotic solution (AA) (Gibco, Cat. No. 15240062, Grand Island, NY) (see Note 5) 5. Collagenase type I (Sigma-Aldrich, Cat. No. C0130, St. Louis, MO) 6. Dimethyl Sulfoxide (DMSO) (Bionichepharma, Cat. No. bip-1, Lake forest, CA). 7. Distilled water 8. Dulbecco’s Modified Eagle’s Medium (DMEM) (Gibco, Cat. No. 10566-016, Grand Island, NY) 9. Dulbecco’s Phosphate-Buffered Saline (DPBS) (Gibco, Cat. No. 14190144, Grand Island, NY) 10. Fetal bovine serum (FBS) (see Notes 6 and 3) (a) Non-ES-grade FBS (WelGENE, Cat. No. #S001-01, Daegu, Republic of Korea) (b) ES-grade FBS (Hyclon, Cat. No. SH30070.03, South Logan, UT)

Laboratory Control and Basic Culture Protocols for Stem Cell Self-Renewal

3

11. Gelatin (Sigma-Aldrich, Cat. No. G9391, St. Louis, MO) 12. Incuwater Germany)

(Applichem,

Cat.

No.

A5219,

Darmstadt,

13. Leukemia inhibitory factor (LIF) (Merck millipore, Cat. No. ESG1107, Temecula, CA) (see Note 7) 14. L-Glutamine (Gibco, Cat. No. 25030081, Grand Island, NY) 15. Mitomycin C (MMC) (Sigma-Aldrich, Cat. No. m4287, St. Louis, MO) 16. Mycoplasma detection kit 17. Nonessential amino acids (NEAA) (Gibco, Cat. No. 11140050, Grand Island, NY) 18. Pharmacidal sprays (Biological Industries, Cat. No. IC110110-L, Kibbutz Beit Haemek, Israel) 19. Plasmocin (Invivogen, Cat. No. ant-mpt, San Diego, CA) 20. Trypan blue solution (Sigma-Aldrich, Cat. No. T8154, St. Louis, MO) 21. β-mercaptoethanol (Gibco, Cat. No. 21985-023, Grand Island, NY) 2.1.2 Equipment

1. Aspiration pump (vacuum pump) Specification Model

Oil-less diaphragm type

Max. pressure

6 kgf/cm2

Vacuum

650  30 mmHg

2. Autoclave Specification Dimension

Chamber (400  670), body (600  650)

Temperature

121  1  C

Pressure

0–3 kg/cm2

Accessories

Digital timer, vent tank, safety valve, temperature sensor and controller, heater

3. Biosafety cabinet class II (clean bench or laminar flow hood) Specification Average velocity

Inflow: 0.53 m/s Down flow: 0.35 m/s

Filter efficiency

HEPA filter/Class 100

Sterilization

Germicidal UV lamp

4

Jeong Mook Lim et al.

4. Centrifuge Specification Max. speed

4,000 rpm or 2,469  g

Drive motor

High torque DC motor

5. ChemiDoc (UV transilluminator) (Bio-RAD, California, Hercules) Specification Application

Chemiluminescence, fluorescence, Gel documentation, colorimetry/densitometry

Illumination modes

Trans UV, white, epi white

Temperature

10–28  C (21  C recommended)

Humidity

95 % humidity, and 5 % CO2. 3. Water bath set at 37  C. 4. Benchtop cell culture centrifuge. 5. Benchtop orbital shaker. 6. Pipet Controller. 7. Serological pipettes (1, 5, 10, and 25 ml). 8. 10-, 20-, 200-, and 1,000-μl micropipettes. 9. 10-, 20-, 200-, and 1,000-μl micropipette tips. 10. Tissue culture treated polystyrene dishes: 6-well, 12-well, and 24-well and 100 mm. 11. Ultra-low attachment 6-well multi-well plates (Corning Inc., cat.no. 3471). 12. 1.5 ml microcentrifuge tubes. 13. Polystyrene conical tubes: 15- and 50-ml. 14. Hemacytometer. 15. Inverted light microscope with 4 and 10 phase objectives. 16. Polyethylene cell lifter (Corning Inc., cat.no. 3008).

2.2 Stock Solutions and Reagents

1. mTeSR™2 animal protein free, defined, feeder-independent medium for maintenance of undifferentiated human ESCs and iPSCs (Stem Cell Technologies, cat.no. 05860). Make aliquots of 50 ml and store at 4  C. 2. Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F-12; Life Technologies, cat.no. 11320-033). 3. 100 N-2 supplement (Life Technologies; cat.no. 17502-048). Make aliquots of 0.5 ml in 1.5 ml microcentrifuge tubes and store at 20  C. 4. 50 B-27 serum-free supplement (Life Technologies, cat.no. 17504-44). Make aliquots of 1.0 ml in 1.5 ml microcentrifuge tubes and store at 20  C.

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5. 100 MEM non-essential amino acids solution (NEAA; Life Technologies; cat.no. 11140-050). Make aliquots of 5 ml and store at 4  C. 6. Penicillin-Streptomycin (P/S) 5,000 U/ml (Life Technologies; cat.no. 15070-063). Make aliquots of 5 ml and store at 20  C. 7. GlutaMAX™ supplement (Life Technologies; cat.no. 35050-061). Make aliquots of 5 ml and store at 20  C. 8. Phosphate-Buffered Saline (PBS), ph 7.4 (Life Technologies, cat.no. 10010023). 9. StemPro® Accutase® cell dissociation reagent (Life Technologies, cat.no. A1110501). Make aliquots of 10 ml and store at 20  C. 10. Matrigel™, Growth Factor Reduced (BD Biosciences, cat.no. 354230). Make aliquots of 250 μl and store at 20  C. 11. Rho-associated protein kinase inhibitor (ROCKi, Y-27632; EMD Millipore, cat.no. SCM075). Dissolve in DMSO at a concentration of 5 mM. Make aliquots of 50 μl in 1.5 ml microcentrifuge tubes and store at 20  C. Protect from light. A final concentration of 5 μM will be used for the first day after passaging hPSCs and the first day of embryoid body (EB) formation. 12. Human recombinant Noggin (R&D Systems, cat.no. 6057NG). Reconstitute in sterile, distilled water at a concentration of 100 μg/ml. Make aliquots of 25 μl in 1.5 ml microcentrifuge tubes and store at 20  C. A final concentration of 50 ng/ml will be used during NPC generation. 13. Dorsomorphin dihydrochloride (DM; Tocris Biosciences, cat. no. 3093). Dissolve in DMSO at a concentration of 25 mM. Make aliquots of 10 μl in 1.5 ml microcentrifuge tubes and store at 20  C. A final concentration of 0.5 μM will be used during NPC generation. 14. Human recombinant bFGF (Life Technologies, cat.no. PHG6014). Reconstitute in sterile, distilled water at a concentration of 30 μg/ml. Make aliquots of 50 μl in 1.5 ml microcentrifuge tubes and store at 20  C. A final concentration of 30 ng/ml will be used to culture and expand NPCs. 15. Human recombinant EGF (R&D Systems, cat.no. 236-EG). Reconstitute in sterile, distilled water at a concentration of 30 μg/ml. Make aliquots of 50 μl in 1.5 ml microcentrifuge tubes and store at 20  C. A final concentration of 30 ng/ml will be used to culture and expand NPCs. 16. Human recombinant BDNF (R&D Systems, cat.no. 248-BD005). Reconstitute in sterile, distilled water a concentration of 20 μg/ml. Make aliquots of 25 μl in 1.5 ml microcentrifuge tubes and store at 20  C. A final concentration of 20 ng/ml will be used during the 4 weeks of neuronal differentiation.

Derivation of hPSC-Derived NPCs

91

17. Human recombinant GDNF (R&D Systems, cat.no. 212-GD). Reconstitute in sterile, distilled water at a concentration of 20 μg/ml. Make aliquots of 25 μl in 1.5 ml microcentrifuge tubes and store at 20  C. A final concentration of 20 ng/ml will be used during the 4 weeks of neuronal differentiation. 18. N-[(3,5-Diflurophenyl)acetyl-]-L-alanyl-2-phenyl]gylcine1,1-dimethlethyl ester (DAPT) γ-secretase inhibitor (Tocris Biosciences; cat.no. 2634). Dissolve in DMSO at a concentration of 5 mM. Make aliquots of 50 μl and store at 20  C. A final concentration of 1.0 μM will be used during the 4 weeks of neuronal differentiation. 19. N6,2’-O-Dibutyryladenosine 30 , 50 -cyclic monophosphate sodium salt (db-cAMP; Sigma, cat.no. D0260). Dissolve in sterile, distilled water at a concentration of 25 mM. Make aliquots of 1.0 ml and store at 20  C. A final concentration of 0.5 mM will be used during the 4 weeks of neuronal differentiation. 20. Poly-L-ornithine 10 mg/ml solution (PLO; Sigma, cat.no. P4957). Make aliquots of 1.0 ml in 1.5 ml microcentrigue tubes and store at 4  C. 21. Laminin (LN) from Engelbreth-Holm-Swarm murine sarcoma basement membrane, 1 mg/ml solution (Sigma, cat.no. L2020). Make aliquots of 1.0 ml in 1.5 ml microcentrifuge tubes and store at 20  C. 22. BD Cytofix™ Fixation Buffer (BD Biosciences, cat.no. 554655). 23. BD Phosflow™ Perm Buffer III (BD Biosciences, cat.no. 558050). 24. Mouse anti-SOX1 antibody (BD Biosciences, cat.no. 560749). 25. Goat anti-SOX2 antibody (Santa Cruz Biotechnology, cat.no. SC-17320). 26. Mouse anti-Nestin 560341).

antibody

(BD

Biosciences,

cat.no.

27. Rabbit anti-MAP2 (Millipore, cat.no. AB5622). 28. Mouse anti-B3T (Fitzgerald, cat.no. 10R-T136A). 29. Alexa Fluor 647 conjugated donkey anti-mouse IgG (Life Technologies, cat.no. A31571). 30. Alexa Fluor 647 conjugated donkey anti-goat IgG (Life Technologies, cat.no. A21447). 31. Alexa Fluor 647 conjugated donkey anti-rabbit IgG (Life Technologies, cat. no. A31573). 32. Alexa Fluor 488 conjugated donkey anti-mouse IgG (Life Technologies, cat. no. A21202). 33. Hoechst 33342 10 mg/ml (Life Technologies, cat.no. H3570).

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David A. Brafman

Medium

1. Neural Induction Medium (NIM). In a 50 ml conical tube, combine 47 ml DMEM/F-12, 1.0 ml 50 B27 Supplement, 0.5 ml 100 N2 Supplement, 0.5 ml NEAA, 0.5 ml GlutaMAX, 0.5 ml P/S, 100 μl of 100 μg/ml Noggin, and 1 μl of 25 mM DM. The medium may be stored at 4  C for up to 1 week. 2. NPC Expansion Medium (NEM). In a 50 ml conical tube, combine 47 ml Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F-12; Life Technologies, cat.no. 11320-033), 1.0 ml 50 B27 Supplement, 0.5 ml 100 N2 Supplement, 0.5 ml NEAA, 0.5 ml GlutaMAX, 0.5 ml P/S, 50 μl of 30 μg/ml EGF, and 50 μl of 30 μg/ml bFGF. The medium may be stored at 4  C for up to 1 week. 3. Neuronal Differentiation Medium (NDM). In a 50 ml conical tube, combine 47.75 ml Dulbecco’s Modified Eagle Medium/ Nutrient Mixture F-12 (DMEM/F-12; Life Technologies, cat.no. 11320-033), 0.5 ml 50 B27 Supplement, 0.25 ml 100 N2 Supplement, 0.5 ml NEAA, 0.5 ml GlutaMAX, 0.5 ml P/S, 50 μl of 20 ng/ml BDNF, 50 μl of 20 ng/ml GDNF, 10 μl of 5 mM DAPT, and 1.0 ml of 25 mM db-cAMP. The medium may be stored at 4  C for up to 1 week.

3

Methods This protocol outlines a serum free method based on the inhibition of TGF-β/Smad signaling that allows for the generation of NPCs and neurons from hPSCs (Fig. 1). First, hPSCs grown in feederindependent conditions on Matrigel-coated plates are forced into three-dimensional aggregates of cells termed embryoid bodies (EBs). Once formed, EBs are plated onto Matrigel-coated plates where they adhere and form neuroepithelial-like rosettes. NPCs are dissociated from EB-derived rosettes, dissociated, replated, and maintained as proliferative, multipotent cells in the presence of bFGF and EGF on laminin (LN) coated plates. Compared to hPSCs, NPCs do not express the pluripotency markers OCT4 and NANOG but express high levels of the pan-neural markers SOX1, SOX2, and NESTIN (Fig. 3a–c). Subsequent differentiation of NPCs to neurons is achieved through the withdrawal of FGF2 and EGF and addition of BDNF, GDNF, db-cAMP, and the Notch inhibitor DAPT. After 4 weeks of treatment in the presence of neuronal induction factors, cells acquire a neuronal morphology (Fig. 2e) and express high levels of the pan-neuronal markers microtubule-associated protein 2 (MAP2; Fig. 4a) and β-TubulinIII (β3T; Fig. 4b).

Derivation of hPSC-Derived NPCs

93

Fig. 2 Morphologies of cell types at each stage of the protocol. Phase contrast images of (a) hPSCs (scale bar ¼ 200 μm), (b) day 5 EBs (scale bar ¼ 200 μm), (c) day 7 neural rosettes (scale bar ¼ 500 μm), (d) passage 3 NPCs (scale bar ¼ 100 μm), and (e) week 4 neurons (scale bar 200 μm) 3.1 Maintenance and Expansion of Human Pluripotent Stem Cells (hPSCs) in Feeder-Independent Conditions

1. HPSCs are maintained in feeder-free conditions on Matrigel™-coated 100 mm plates with mTeSR™2 culture medium. 2. Prior to passaging hPSCs, prepare a working solution of Matrigel by thawing one aliquot of Matrigel™ on ice (see Note 1). 3. Make a 1:50 dilution of Matrigel™ in cold DMEM/F12 in a 50 ml conical. The working solution of Matrigel™ should be maintained on ice and can be stored at 4  C for up to 1 week. 4. Coat a 100 mm dished with 10 ml of the Matrigel™ working solution per dish at 37  C for 20 min (see Note 2). 5. Aspirate Matrigel™ solution and wash once with 10 ml of sterile PBS (see Note 3). 6. Warm mTeSR™2 medium and Accutase® solution in a 37  C water bath. 7. Aspirate medium from 100 mm plate of growing hPSCs and gently wash with 5 ml of PBS. 8. Add 5 ml of Accutase® solution and incubate in the CO2 incubator for 10 min. After 10 min, gently tap the sides of the plate against a solid surface to ensure complete cell dissociation. 9. Observe under the microscope to determine if additional incubation and tapping is required. 10. Add 5 ml of mTeSR™2 medium to the plate. 11. Using a 10 ml serological pipette, gently wash off the remaining attached cells until the plate is clear. 12. Gently triturate the cell suspension until all noticeable cell clumps are broken up. 13. Transfer the cell suspension to a 15 ml conical tube. 14. Take 10 μl of the cell suspension to perform a cell count using the hemacytometer. 15. Centrifuge the tube at 200  g for 5 min.

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16. Resuspend the cells in the appropriate amount of mTeSR™2 medium so that the final cell concentration is 5.0  105 cells/ml. Carefully pipette the cell suspension up and down 3–4 times with a 10 ml serological pipette. 17. In a separate 15 ml conical tube add the following: 1 ml of cell suspension, 9 ml of fresh mTeSR™2 medium, and 10 μl of 5 mM ROCKi (see Note 4). 18. Add the 10 ml of the cell suspension onto a Matrigel™ coated plate. 19. Place the plate in the CO2 incubator. Gently move the plate in several quick horizontal and vertical motions to disperse the cells evenly across the cell culture surface. 20. Change the medium daily by aspirating the old medium and adding 10 ml of fresh mTeSR™2 medium (see Note 5). After 2–3 days, dense, bright colonies should appear (Fig. 2a). HPSCs need to be passaged when the colonies begin to merge and the culture reaches approximately 75 % confluency, which typically occurs 4–6 days after passaging (see Note 6). 3.2 Embryoid Body (EB) Formation and Growth

1. Warm Neural Induction Medium (NIM) and Accutase® solution in a 37  C water bath. 2. Aspirate mTeSR™2 medium from healthy, 75 % confluent hPSC 100 mm plate grown. 3. Carefully wash the plate with 10 ml of PBS. 4. Add 5 ml of Accutase® solution and incubate in the CO2 incubator for 10 min. After 10 min, gently tap the sides of the plate against a solid surface to ensure complete cell dissociation. Observe under the microscope to determine if additional incubation and tapping is required. 5. Add 5 ml of NIM to the plate. 6. Using a 10 ml serological pipette, gently wash off the remaining attached cells until the plate is clear. 7. Gently triturate the cell suspension until all noticeable cell clumps are broken up. 8. Transfer the cell suspension to a 15 ml conical tube. 9. Take 10 μl of the cell suspension to perform a cell count using the hemacytometer. 10. Centrifuge the tube at 200  g for 5 min. 11. Resuspend the cells in NIM such that the final concentration of cells is 5.0  105 cells/ml (see Note 7). 12. Add 4 ml of cell suspension per well of an ultra-low attachment 6-well plate. A total of 2  106 cells will be present in each well. 13. Add 4 μl of 5 mM ROCKi per well.

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14. Place the plate on an orbital shaker that has been placed inside a CO2 incubator. Set the shaker speed at 95 RPM. 15. After 24 h, examine the cells under the microscope. Small clusters of cells should be visible. Do not change medium. 16. After 48 h, carefully aspirate approximately ½ (2 ml) of the medium from each well using a 5 ml serological pipette (see Note 8). 17. Add 2 ml of NIM to each well (see Note 9). 18. Place plate back on orbital shaker that has been placed inside a CO2 incubator. Changes of half of the medium should be made daily. After 2 days, large EB aggregates should be visible (Fig. 2b). 3.3 EB Plating and Formation of Neural Rosettes

1. After 5 days of EB growth, the EBs will be plated on Matrigel™ coated substrates to induce formation of neural rosettes. 2. Prepare a working solution of Matrigel by thawing one aliquot of Matrigel™ on ice. 3. Making a 1:50 dilution of Matrigel™ in cold DMEM: F12 in a 50 ml conical (see Note 1). The working solution of Matrigel™ should be maintained on ice and can be stored at 4  C for up to 1 week. 4. Coat 3 wells of a 6-well plate with 3 ml of the Matrigel™ working solution per well. Incubate at 37  C for 20 min (see Note 2). 5. Aspirate Matrigel™ solution and wash each well once with 3 ml of sterile PBS (see Note 3). 6. Add 12 ml of NIM to a 50 ml conical tube. 7. Carefully transfer all 4 ml of EB suspension to the 50 ml conical tube with NIM. 8. Gently invert the conical tube 2–3 times to mix the EBs evenly throughout the solution. 9. Quickly transfer 4 ml of the EB suspension to each Matrigel™ coated well (see Note 10). Place the plate in the CO2 incubator. Gently move the plate in several quick horizontal and vertical motions to disperse the EBs evenly across the cell culture surface (see Note 11). 10. After 24 h, carefully examine cells under the microscope. EBs should have settled and adhered to the plate (see Note 12). Do not change the medium. 11. After 48 h, carefully aspirate approximately ½ (2 ml) of the medium from each well using a 5 ml serological pipette. 12. Add 2 ml of NIM to each well.

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13. Place plate back inside a CO2 incubator. Changes of half of the medium should be made daily. After 3 days, the EBs should spread out on the Matrigel™ substrate and neural rosette structures should be visible (Fig. 2c). 3.4 Generation, Expansion, and Characterization of Neural Progenitor Cells (NPCs)

1. After 7 days of culture, rosettes should be dissociated and replated onto poly-L-ornithine/Laminin (PLO/LN) coated plates. 2. To generate PLO/LN coated plates, make a 50 μg/ml working solution of poly-L-ornithine (PLO) by adding 200 μl of 10 mg/ml PLO stock solution to 50 ml of sterile PBS in a 50 ml conical tube. 3. Coat a 100 mm dish with 10 ml of the PLO working solution at 37  C for 4 h. 4. Aspirate the PLO working solution and wash the PLO-coated 100 mm plates 3 times with 10 ml of PBS. 5. Thaw an aliquot of laminin (LN) on ice. Prepare a 20 μg/ml working solution of LN by adding 200 μl of 1 mg/ml LN to 50 ml of sterile PBS in a 50 ml conical tube. 6. Coat the PLO-coated 100 mm plate with 10 ml of LN working solution at 37  C for 4 h (see Note 13). 7. Aspirate the LN working solution and rinse the plates 1 time with 10 ml of PBS. 8. Warm Neural Expansion Medium (NEM) and Accutase® solution in a 37  C water bath. 9. Aspirate NIM from day 7 rosette cultures. 10. Add 3 ml of Accutase® solution to each well and incubate in the CO2 incubator for 10 min. After 10 min, gently tap the sides of the plate against a solid surface to ensure complete cell dissociation (see Note 14). 11. Using a 10 ml serological pipette, gently wash off the remaining attached cells until the plate is clear. 12. Gently triturate the cell suspension until all noticeable cell clumps are broken up. 13. Transfer the cell suspension to a 15 ml conical tube. 14. Take 10 μl of the cell suspension to perform a cell count using the hemacytometer. 15. Centrifuge the tube at 200  g for 5 min. 16. Resuspend the cells in the appropriate amount of NEM medium so that the final cell concentration is 1.1  106 cell/ml. Carefully pipette the cell suspension up and down 2–3 times with a 10 ml serological pipette until all cell aggregates are broken up.

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17. In a separate 15 ml conical tube add the following: 1 ml of cell suspension and 9 ml of fresh NEM (see Note 15). 18. Add the 10 ml of the cell suspension onto a PLO/LN coated plate. 19. Place the plate in the CO2 incubator. Gently move the plate in several quick horizontal and vertical motions to disperse the cells evenly across the cell culture surface. 20. Change the medium every other day by aspirating the old medium and adding 10 ml of fresh NEM medium. 21. Once the cells reach confluency, they should be Accutase® passaged at a density of 2.0  104/cm2 onto new PLO/LN coated plates which typically occurs every 5–7 days. After 2–3 passages, NPCs should appear with a morphology as displayed in Fig. 2d. 3.5 Characterization of NPCs by Immunofluorescent Staining

1. After 3 passages, NPCs can be characterized by immunofluorescent (IF) staining for expression of pan-neural markers SOX1, SOX2, and Nestin. 2. Warm NEM and Accutase® solution in a 37  C water bath. 3. Aspirate NEM from passage 3 NPC cultures. 4. Add 5 ml of Accutase® solution to each plate and incubate in the CO2 incubator for 5 min. After 5 min, gently tap the sides of the plate against a solid surface to ensure complete cell dissociation. 5. Observe under the microscope to determine if additional incubation and tapping is required. 6. Add 5 ml of NEM medium to the plate. 7. Using a 10 ml serological pipette, gently wash off the remaining attached cells until the plate is clear. 8. Gently triturate the cell suspension until all noticeable cell clumps are broken up. 9. Transfer the cell suspension to a 15 ml conical tube. 10. Take 10 μl of the cell suspension to perform a cell count using the hemacytometer. 11. Centrifuge the tube at 200  g for 5 min. 12. Resuspend the cells in the appropriate amount of NEM medium so that the final cell concentration is 1.0  105 cells/ml. Carefully pipette the cell suspension up and down 3–4 times with a 10 ml serological pipette. 13. Into 3 wells of a PLO/LN coated 24-well plate add 1.0 ml of cell suspension. 14. Place plate back inside a CO2 incubator. 15. After 24 h, aspirate the NEM medium from each well.

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16. Wash each well twice with 1 ml of sterile PBS. 17. Fix the NPCs by adding 1 ml of Fixation Buffer to each well. Incubate the cells in Fixation Buffer for 10 min at room temperature. 18. Aspirate the Fixation Buffer and wash each well twice with 1 ml of sterile PBS. 19. Permeablize cells by adding 1 ml of Perm Buffer to each well. Incubate the cells in Perm Buffer for 30 min at room temperature. 20. Aspirate the Perm Buffer and wash each well twice with 1 ml of sterile PBS. 21. Add the following primary antibodies to the following wells: (1) Well 1: 0.5 ml of mouse anti-SOX1 (1:10 dilution in PBS), (2) Well 2: 0.5 ml of goat anti-SOX2 (1:50 dilution in PBS), and (3) Well 3: 0.5 ml of mouse anti Nestin (1:10). Incubate the primary antibodies overnight at 4  C. 22. Aspirate the primary antibodies and wash each well twice with 1 ml of sterile PBS. 23. Add the following secondary antibodies to each well: (1) Well 1: 0.5 ml of Alexa Fluor 647 conjugated donkey anti-mouse IgG (1:200 dilution in PBS), (2) Well 2: 0.5 ml of Alexa Fluor 647 conjugate donkey anti-goat IgG (1:200 dilution in PBS), (3) Well 3: 0.5 ml of Alexa Fluor 647 conjugated donkey anti-mouse IgG (1:200 dilution in PBS). Incubate the secondary antibodies for 1 h at room temperature in the dark. 24. Aspirate the secondary antibodies and wash each well twice with 1 ml of sterile PBS. 25. Add 0.5 ml of Hoechst 3342 (1:5,000 dilution in PBS) to each well to counterstain nuclei. Incubate for 10 min at room temperature in the dark. 26. Aspirate the Hoechst 3342 and wash each well twice with 1 ml of sterile PBS. 27. Image NPCs using a fluorescent microscope (Fig. 3a–c). 3.6 Differentiation of NPCs to Neurons

1. Warm Neuronal Differentiation Medium (NDM) and Accutase® solution in a 37  C water bath. 2. Aspirate NEM from passage 3 NPC cultures. 3. Add 5 ml of Accutase® solution to each plate and incubate in the CO2 incubator for 5 min. After 5 min, gently tap the sides of the plate against a solid surface to ensure complete cell dissociation. 4. Observe under the microscope to determine if additional incubation and tapping is required. 5. Add 5 ml of NDM medium to the plate.

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Fig. 3 Immunofluorescent characterization of passage 3 NPCs. Immunofluorescence of (a) SOX1, (b) SOX2, and (c) Nestin (scale bar ¼ 200 μm)

6. Using a 10 ml serological pipette, gently wash off the remaining attached cells until the plate is clear. 7. Gently triturate the cell suspension until all noticeable cell clumps are broken up. 8. Transfer the cell suspension to a 15 ml conical tube. 9. Take 10 μl of the cell suspension to perform a cell count using the hemacytometer. 10. Centrifuge the tube at 200  g for 5 min. 11. Resuspend the cells in the appropriate amount of NDM medium so that the final cell concentration is 1.5  105 cell/ml. Carefully pipette the cell suspension up and down 2–3 times with a 10 ml serological pipette until all cell aggregates are broken up. 12. Add 1.5 ml of cell suspension to each well of a 12-well PLO/LN coated plate. 13. Place the plate in the CO2 incubator. Gently move the plate in several quick horizontal and vertical motions to disperse the cells evenly across the cell culture surface. 14. Change the medium every other day by aspirating the old medium and adding 10 ml of fresh NDM medium. 15. After 4 weeks in NDM medium cells should acquire a neuronal morphology (Fig. 2e). 3.7 Characterization of Neurons by Immunofluorescent Staining

1. Fix the neurons by adding 1.5 ml of Fixation Buffer to each well. Incubate the cells in Fixation Buffer for 10 min at room temperature. 2. Aspirate the Fixation Buffer and wash each well twice with 2 ml of sterile PBS.

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Fig. 4 Immunofluorescent characterization of week 4 neurons differentiated from passage 3 NPCs. Immunofluorescence of (a) β3T and (b) MAP2 (scale bar ¼ 200 μm)

3. Permeablize cells by adding 1.5 ml of Perm Buffer to each well. Incubate the cells in Perm Buffer for 30 min at room temperature. 4. Aspirate the Perm Buffer and wash each well twice with 2 ml of sterile PBS. 5. Add 1.5 ml of mouse anti-β3T (1:1,000 dilution in PBS) and rabbit anti-MAP2 (1:500 dilution in PBS) primary antibodies to each well. Incubate the primary antibodies overnight at 4  C. 6. Aspirate the primary antibodies and wash each well twice with 2 ml of sterile PBS. 7. Add 1.5 ml of Alexa Fluor 488 conjugated donkey anti-mouse IgG (1:200 dilution in PBS) and Alexa Fluor 647 conjugated donkey anti-rabbit IgG (1:200 dilution in PBS). 8. Incubate the secondary antibodies for 1 h at room temperature in the dark. 9. Aspirate the secondary antibodies and wash each well twice with 1 ml of sterile PBS. 10. Add 0.5 ml of Hoechst 3342 (1:5,000 dilution in PBS) to each well to counterstain nuclei. Incubate for 10 min at room temperature in the dark. 11. Aspirate the Hoechst 3342 and wash each well twice with 1 ml of sterile PBS. 12. Image neurons using a fluorescent microscope (Fig. 4b).

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Notes 1. Matrigel™ should always be kept on ice during thawing in order to prevent gelling. 2. If not used immediately, the culture plate should be sealed to avoid evaporation of Matrigel™ working solution. Culture plates can be sealed with Parafilm® and can be stored at 4  C for up to 1 week after coating. Prior to using stored Matrigel™ coated plates, allow the plates to equilibrate with room temperature for 15 min. 3. Aspirate Matrigel™ solution immediately prior to seeding of cells. Do not allow the Matrigel™ coated surfaces to dry out. 4. HPSCs can be grown and maintained in other tissue culture plate sizes (e.g., multi-well dishes). Cells should be plated in these other formats at a density 9.0  103/cm2. 5. ROCKi is only added during passaging to aid in hPSC survival. 6. If the colonies are passaged too late, then the cultures will begin to display signs of differentiation, such as cell types with noncolony morphology. These cells can be removed using manual dissection techniques prior to passaging. 7. Optimal cell density for EB formation may be hPSC-line dependent. When working with a new cell line, experiment with a range of cell densities. 8. Use caution when aspirating medium as not to disturb the EBs in suspension. 9. Do not add ROCKi. ROCKi is added on the first day of EB formation to aid with cell survival. 10. If the transfer of the EBs to the Matrigel™ coated plates is not performed in a timely manner, the EBs may begin to settle to the bottom of the 50 ml conical. 11. Failure to evenly disperse the EBs throughout the cell culture surface may result in aggregation of EBs into the center of the well and failure of neural rosettes to form. 12. Do not move the plate prior to 24 h to allow for sufficient time for the EBs to settle and adhere. On average, approximately 5–10 % of the EBs will be of cystic or non-uniform shape and will not attach to the Matrigel™ coated plates. 13. If not used immediately, the PLO/LN culture plate should be sealed to avoid evaporation of laminin working solution. Culture plates can be sealed with Parafilm® and can be stored at 20  C for up to 2 months after coating. Prior to using stored PLO/LN coated plates, allow the plates to thaw at 37  C for 15 min.

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14. Complete dissociation of the rosettes from the cell culture surface may require the use of a cell lifter. 15. Dissociated rosettes can be plated in other culture plate sizes (e.g., multi-well dishes). Cells should be plated in these other formats at a density 2.0  104/cm2.

Acknowledgements The research was supported by funding from the University of California-San Diego Stem Cell Program, a gift from Michael and Nancy Kaehr, and the California Institute for Regenerative Medicine (RT2-01889 and RB3-05086). References 1. Lunn JS, Sakowski SA, Hur J, Feldman EL (2011) Stem cell technology for neurodegenerative diseases. Ann Neurol 70:353–361 2. Yoo J, Kim HS, Hwang DY (2012) Stem cells as promising therapeutic options for neurological disorders. J Cell Biochem 114:743–753 3. Lindvall O, Kokaia Z (2006) Stem cells for the treatment of neurological disorders. Nature 441:1094–1096 4. Lindvall O, Kokaia Z (2010) Stem cells in human neurodegenerative disorders–time for clinical translation? J Clin Invest 120:29–40 5. Chambers SM, Fasano CA, Papapetrou EP, Tomishima M, Sadelain M, Studer L (2009) Highly efficient neural conversion of human ES and iPS cells by dual inhibition of SMAD signaling. Nat Biotechnol 27:275–280 6. Elkabetz Y, Panagiotakos G, Al Shamy G, Socci ND, Tabar V, Studer L (2008) Human ES cell-derived neural rosettes reveal a functionally distinct early neural stem cell stage. Genes Dev 22:152–165

7. Shin S, Mitalipova M, Noggle S, Tibbitts D, Venable A, Rao R, Stice SL (2006) Long-term proliferation of human embryonic stem cellderived neuroepithelial cells using defined adherent culture conditions. Stem Cells 24:125–138 8. Bretzner F, Gilbert F, Baylis F, Brownstone RM (2011) Target populations for first-inhuman embryonic stem cell research in spinal cord injury. Cell Stem Cell 8:468–475 9. Schwartz SD, Hubschman JP, Heilwell G, Franco-Cardenas V, Pan CK, Ostrick RM, Mickunas E, Gay R, Klimanskaya I, Lanza R (2012) Embryonic stem cell trials for macular degeneration: a preliminary report. Lancet 379:713–720 10. Marchetto MC, Winner B, Gage FH (2010) Pluripotent stem cells in neurodegenerative and neurodevelopmental diseases. Hum Mol Genet 19:R71–R76

Methods in Molecular Biology (2015) 1212: 103–112 DOI 10.1007/7651_2014_91 © Springer Science+Business Media New York 2014 Published online: 26 July 2014

Isolation, Long-Term Expansion, and Differentiation of Murine Neural Stem Cells Alexandra Bizy and Sacri R. Ferro´n Abstract Stem cells are capable of extensive self-renewal while preserving the ability to generate cell progeny that can differentiate into different cell types. Here, we describe some methods for the isolation of neural stem cells (NSCs) from the adult murine subependymal zone (SEZ), their extensive culturing and the assessment of their full developmental potential, particularly with respect to their differentiation capacity. The procedure includes chemically defined conditions such as absence of serum and addition of specific growth factors, in which differentiated cells die and are rapidly eliminated from the culture. In contrast, undifferentiated precursors become hypertrophic and proliferate, forming clonal spherical clusters called “neurospheres.” Experimental manipulation of NSCs identifies populations of cells with differential restriction in their selfrenewal potential and introduces a great interest in defining the conditions that guide their differentiation into a variety of neuronal and glial subtypes, aspects that have important implications for their use in future clinical purposes. Keywords: Neural stem cells, SEZ, Neurogenic niche, Self-renewal, Multipotency, Mitogens, Serum-free medium

1

Introduction Stem cells are defined as undifferentiated cells capable of extensive proliferation and self-renewal and that can give rise to functional differentiated progeny to regenerate the tissues/organs in which they reside. In contrast, the term progenitor cell is used to define undifferentiated cells possessing limited proliferation capacity and a more restricted developmental potential. In the adult brain, the SEZ is a very active germinal center in which continuous neurogenesis appears to be supported by a relatively quiescent NSC population. SEZ stem cells (named type B cells) proliferate at a low rate, selfrenew and contribute to transit-amplifying progenitors (C cells). C cells are proliferating cells that rapidly divide and give rise to neuroblasts (A cells) that migrate out of the SEZ and terminally differentiate into local-circuitry neurons in the olfactory bulb (1–4). NSCs have provided a unique model system for understanding the

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molecular mechanisms that control cell-fate specification in the mammalian nervous system. Prospective isolation of pure NSCs from the SEZ niche has been widely used in the field for the study of their fundamental properties and potential (5–10). The methodology described in this chapter refers to the isolation, culturing, longterm expansion and differentiation of NSCs under chemically defined conditions. The specific conditions required for the neural stem cells to become the main cell type in the culture are: (1) absence of serum and of a cell adhesion substrate, (2) addition of the appropriate growth factors (i.e., EGF, epidermal growth factor and FGF2, fibroblast growth factor 2), and (3) low cell density (5  104 cells/cm2). Under these selective conditions differentiated cells die and are rapidly eliminated from the culture. Conversely, a very small fraction of undifferentiated precursors enter into an active proliferative state, become hypertrophic and round up, forming clonal spherical clusters called “neurospheres” (Fig. 1). These neurospheres contain cells capable of generating new clusters (self-renewal) and can be subcultured several times. When harvested under the same culture conditions, they can be propagated for months and their growth rate increases exponentially with time in culture. Furthermore, neurospheres display a steady capacity to generate neurons, astrocytes, and oligodendrocytes upon passages, also demonstrating their multipotency capacity in vitro. The frequency of long-lived NSCs in dissociated SEZ is often estimated as the number of cells capable of generating such primary spheres. Therefore, although there is still controversy about the real functional relationship between neurosphere-forming cells and the stem cell population in vivo (8), this system represents an optimal model to investigate the progressive restriction and the potential use of neural stem cells for restorative neurogenesis in disease or trauma.

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Materials Prepare all solutions in sterile conditions using sterile-filtered components and sterile material. For cell culture, use pre-warmed medium and store at 4 ºC unless stated otherwise. The following materials and media are the same for both fetal and adult NSCs isolation and expansion.

2.1 Primary Culture Preparation Components

1. Dissection tools: large and small scissors, small curved forceps, small spatula, small and regular scalpels (see Note 1). 2. Plates: 35-mm, 100-mm dishes and p24 well plates. Add cold sterile PBS to sterile 35-mm and 100-mm dishes. Place mouse head and brain in the 100-mm dishes for dissection. Transfer the dissected pieces of tissue in 35-mm plates with fresh PBS and leave on ice until tissue dissociation (see Note 2). Prepare p24 well plates to seed the NSCs.

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Fig. 1 Neural stem cells culture established from the adult SEZ. (a) Phase contrast images showing the process of neurosphere formation after 1, 2, 4, and 7 days in vitro. (b) Electron microscopy image of a neurosphere grown after 4 days. (c) Schematic drawing depicting the different steps for isolation of NSCs from adult NSCs and primary neurosphere formation. The long-term expansion of neurosphere cultures is also showed together with the differentiation of NSCs after mitogens removal and addition of Matrigel to give rise to astrocytes (GFAP, red ), neurons (βIII-tubulin, green), and oligodendrocytes (O4, blue). Scale bars: in a: 25 μm; in b: 10 μm; in c, left panel, 30 μm; right panel, 60 μm

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3. Enzymatic mixture for tissue dissociation: 0.9 mg/ml papain (Worthington DBA), 0.2 mg/ml L-cysteine (Sigma), and 0.2 mg/ml EDTA (Sigma) in EBSS (Earle’s Balanced Salts Solution, Gibco BRL) (see Note 3). 4. 70 % EtOH (see Note 4). 2.2 Expansion Medium Components

1. 10 hormone mix: DMEM/F12 (Dulbecco’s Modified Eagles Medium/F12) supplemented with 0.5 % glucose, 0.09 % NaHCO3, 4 mM HEPES, 0.8 mg/ml apo-t-transferrin (Sigma), 0.2 mg/ml insulin (bovine, Sigma), 90 μg/ml putrescine (Sigma), 160 nM progesterone (Sigma), and 240 nM Na2SeO3 (Sigma). Prepare 0.2 mg/ml insulin solution: dilute 100 mg insulin in 4 ml 0.1 N HCl sterile and 36 ml ultrapure H2O. Prepare 90 μg/ml putrescine solution: weigh 38.6 mg putrescine and dilute in 40 ml of ultrapure H2O (see Note 5). 2. Control medium: DMEM/F12 supplemented with 0.6 % glucose (Sigma), 0.1 % NaHCO3 (Sigma), 5 mM HEPES (Sigma), 2 mM L-glutamine (Gibco BRL), 50 U ea./ml penicillin/ streptomycin (Gibco BRL), and 10 % hormone mix (10). Sterile-filter the medium through a 0.22 μm-pore filter and keep at 4 ºC until use (see Note 6). 3. Complete medium: control medium supplemented with 4 mg/ml BSA (bovine serum albumin, Sigma), 0.7 U/ml heparin (Sigma), 20 ng/ml EGF (epidermal growth factor, human recombinant, Gibco BRL), and 10 ng/ml FGF2 (basic fibroblast growth factor 2, human recombinant, Sigma). Sterile-filter the medium through a 0.22 μm-pore filter and keep at 4 ºC until use (see Note 7). 4. Accutase solution (Sigma): aliquot and store at 20  C until use. 5. 0.1 % Trypan blue (Sigma) in PBS.

2.3 Differentiation Medium Components

1. Matrigel®: growth factor reduced (Becton Dickinson). Stock solution at 15 mg/ml, diluted 1:100 in control medium for coating (see Note 8). 2. Differentiation medium I: Control medium + BSA (Sigma) + 0.7 U/ml Heparin (Sigma) + 10 ng/ml FGF2 (Sigma). 3. Differentiation medium II: Control medium + BSA + 0.7 U/ ml Heparin + 10 % FBS (Fetal Bovine Serum, Gibco BRL).

2.4 Immunodetection Components

1. Round glass coverslips (O. Kindler GmbH & Co.) sterilized by autoclaving at 120 ºC for 30 min. 2. Fixative: PFA 4 %, 20 min at room temperature. 3. Blocking solution: 0.1 M PBS, 0.2 % Triton X-100, 1 % glycine, and 10 % FBS (Gibco BRL) or CS (Calf Serum, Gibco BRL).

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Sterile-filter the solution through a 0.22 μm-pore filter and store at 4 ºC until use. 4. Primary antibodies: –

Monoclonal rabbit anti-βIII-tubulin (1:300 dilution, Sigma).



Polyclonal chicken anti-GFAP (1:500 dilution, Millipore).



Monoclonal mouse anti-O4 (1:2 dilution, Hybridoma bank).

Prepare fresh primary antibodies in the blocking solution and store at 4 ºC until use. 5. Secondary antibodies: –

Cy3-AffiniPure F(ab’)2 Frag Donkey Anti-mouse IgG secondary antibody (1:500 dilution, Jackson Immunochemicals).



Alexa Fluor® 488-AffiniPure F(ab’)2 Fragment Donkey Anti-Rabbit IgG secondary (1:500 dilution, Molecular Probes).



Alexa Fluor® 647-AffiniPure F(ab’)2 Fragment Donkey Anti-Chicken IgG secondary antibody (1:800 dilution, Molecular Probes).

Prepare fresh secondary antibodies in the blocking solution. Keep in the dark at 4 ºC until use. 6. DAPI (40 -6-Diamindino-2-phenylindole dihydrochloride hydrate) (Sigma). Working concentration: 2 μg/ml in ultrapure H2O. Keep in the dark at 4 ºC. 7. Mounting medium: FluorSave (Calbiochem) (see Note 9).

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Methods

3.1 Isolation and Primary Culture of Neural Stem Cells. Neurosphere Formation

Clean and disinfect the working surface with 70 % ethanol to avoid contamination. Have all the materials and surgical instruments ready before starting the dissection. Adult mice 2–4 months old are used. Sacrifice animal, remove and dissect the brain outside the laminar flow hood. SEZ tissue from one brain is sufficient to start a culture. 1. Add cold sterile PBS to sterile 100-mm dishes where mouse head and brain will be placed for dissection and to several 35mm dishes to subsequently wash dissected tissues. 2. Kill animal by cervical dislocation and cut the head off using large scissors. Rinse the head with 70 % ethanol. Using small scissors remove the skin of the head and make a longitudinal cut through the skull along the sagittal suture. Using curved forceps peel off the skull, scoop out the brain with a spatula

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and place it in a petri dish containing sterile PBS. Place the brain under a dissecting microscope (use 10 magnification). Use a scalpel to slice off the olfactory bulbs. Make a second incision caudally to get a 2–3 mm-thick slice containing the lateral ventricles. Discard the rest of the brain caudal to the slice. Using fine micro-scalpels trim a thin layer of tissue adjacent to the ventricles (use 25 magnification), excluding the striatal parenchyma and the corpus callosum. Place each dissected SEZ into a 35-mm petri dish containing sterile PBS. 3. Using fine scalpels chop each SEZ into small pieces and transfer them to a 15-ml tube containing the enzymatic solution (5 ml per animal). Incubate the tubes at 37  C for 30 min to allow digestion (see Note 10). 4. Add 4 ml of control medium to inactivate papain and pellet the tissue by centrifuging at 300  g for 5 min. Remove supernatant and add 3 ml of control medium. Dissociate by gently triturating 10–20 times using a fire-polished glass Pasteur pipette until all the pieces are dissociated and cell suspension is homogeneous (see Note 11). Add 3 ml of control medium to wash cells and pellet them by centrifuging at 300  g for 10 min. Remove the supernatant, add 1 ml of pre-warmed complete medium and resuspend the pellet by pipetting up and down until suspension is homogeneous. 5. Seed cells obtained from 1 SEZ in two wells of a 24-well culture plate (surface area: 1.9 cm2; 1 ml final volume per well: 0.5 ml complete medium plus 0.5 ml cell suspension). Incubate at 37  C in a 5 % CO2 humidified incubator for 10 days (see Note 12). 3.2 Expansion of Neural Stem Cells

Cells proliferate to form neurospheres that can be easily observed under phase contrast and electronic microscopy (Fig. 1). 1. After 10 days, collect medium with primary neurospheres from each well and transfer it to a 15-ml tube containing 5 ml of pre-warmed PBS. Pellet cell suspension by centrifugation at 30  g for 10 min (see Note 13). 2. Remove the supernatant, add 1 ml of Accutase solution and incubate for 10 min at room temperature (see Note 14). Add 3 ml of control medium to wash cells and pellet them by centrifugation at 300  g for 10 min. Remove the supernatant, add 1 ml of pre-warmed complete medium and resuspend the pellet by pipetting up and down until suspension is homogeneous. 3. Count viable cells by trypan blue exclusion by mixing 10 μl of cell suspension with 10 μl of dye solution and score the number of viable cells using a Neubauer chamber.

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4. Seed cells in pre-warmed complete medium at the appropriate density in untreated tissue flasks. For bulk expansion, plate cells at 10,000 per cm2 (50 cells/μl). For self-renewal experiments, seed cells at the very low density of 500 cells per p96-well (2.5 cells/μl or in 0.3 cm2) in 0.2 ml of complete medium, to ensure clonogenicity. Incubate at 37 ºC/5 % CO2 in a humidified incubator. The total cell number should increase two- to five folds at each passage (see Note 15). 3.3 Differentiation of Neural Stem Cells and Assessment of Multipotency

Plating neural stem cells onto an adhesion substrate like Matrigel and removing the growth factors from the culture medium is sufficient to promote their spontaneous differentiation into neuronal and glial cells (Fig. 1). 1. Add 0.25 ml of 1 Matrigel (diluted in control medium) to a p24 wells plate containing the round glass coverslips. Incubate at 37  C for at least 2 h (see Note 16). 2. Collect neural stem cells from the plates or flasks and transfer to a 15 ml sterile plastic conical tube using a sterile plastic pipette. Pellet cell suspension by centrifugation at 30  g for 10 min (see Note 17). 3. Remove the supernatant, add 1 ml of accutase solution and incubate for 10 min at room temperature. To wash cells from growth factors, remove supernatant and resuspend cells gently in 10 ml of control medium (see Note 18). 4. Centrifuge for 10 min at 300  g and remove supernatant. Resuspend the cells in 1 ml of pre-warmed differentiation medium 1. Count viable cells by trypan blue exclusion using a Neubauer chamber. 5. Aspirate Matrigel and wash twice the p24 well plate coverslips with ultrapure water. Seed 80,000 cells per cm2 and incubate at 37 ºC during 48 h (see Note 19). 6. Remove the differentiation medium 1 and add the differentiation medium 2. Incubate at 37 ºC for 5 days. Under these conditions simultaneous detection of the three cell phenotypes is usually successful 7 days after plating (Fig. 1). 7. Remove the differentiation medium 2 and wash once with PBS. For fixation, add PFA 4 % during 20 min at room temperature. Rinse twice with PBS. Store in PBS with 0.05 % sodium azide at 4 ºC or proceed directly to immunocytochemistry as follows. 8. Dilute primary antibodies including chicken anti-GFAP (1:500, Millipore) and rabbit anti-βIII-tubulin (1:300 dilution, Sigma) in blocking solution. Mouse anti-O4 (1:2, Hybridoma Bank) antibody must be diluted in blocking

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solution without Triton. Incubate primary antibodies overnight at 4 ºC or 1.30 h at 37 ºC. 9. Rinse 2–3 times, 5 min each time with PBS 0.1 M. Dilute secondary antibodies including Cy3 anti-mouse (1:500, Jackson), Alexa 488 anti-rabbit (1:500, Jackson) and Alexa 647 anti-chicken (1:500, Jackson) in blocking solution for 1 h at room temperature in the dark. 10. Rinse 2–3 times, 10 min each time with PBS 0.1 M and stain with 2 μg/ml of DAPI for 5 min in the dark. 11. Rinse in PBS 0.1 M and mount with FluorSave. 12. Determine the presence of the three CNS cell types by analyzing the coverslips under fluorescence microscopy.

4

Notes 1. It is necessary to have all the materials and instruments ready before starting the dissection. Sterilize dissection tools by autoclaving at 120 ºC for 30 min. 2. To prevent cell death, it is important to keep tissue in ice cold PBS during the whole dissection process (Fig. 2). 3. For optimal efficiency, it is recommended to prepare the enzymatic solution the day of the dissociation. Prepare the mix in a sterile plastic tube and keep 30 min at 37 ºC or until the solution is clear. Do not forget to sterile-filter through a 0.22 μm-pore filter and keep at 4 ºC until use. 4. Ethanol must be used to maintain the tools sterile during the dissection, as well as disinfecting the working surfaces and rinse the decapitated head or the abdomen in case of fetal dissociation. Before dissection, introduce tools in 70 % EtOH in a beaker with cotton on the bottom to avoid spoiling the tips of the forceps and scissors. 5. Prepare 500 ml of 10 stock hormone mix solution. Mix well and filter. To avoid repeated freezing and thawing, prepare 50 ml aliquots of stock solution and store at 20 ºC. 6. The medium should be discarded after 15 days kept at 4 ºC (Fig. 2). 7. Add fresh mitogens the day of use. Prepare control medium plus BSA and heparin. Sterile-filter the medium through a 0.22 μm-pore filter and keep at 4 ºC until use. The medium does not need to be filtered after the addition of mitogens to avoid the mitogens attach to the filter. 8. Thaw Matrigel® stock overnight on ice at 4  C. Aliquot Matrigel® using precooled pipettes, tips, and tubes and keep at

Isolation, Long-Term Expansion, and Differentiation of Murine Neural Stem Cells PROBLEM

POSSIBLE REASONS Excessive enzymatic digestion Insufficient enzymatic digestion

Low primary neurosphere yield

Excessive mechanical trituration Tissue degradation Aged medium

Low efficiency in secondary neurospheres

Wrong number of cells plated Excessive enzymatic digestion Excessive mechanical trituration Aged medium Wrong number of cells plated

Neurosphere aggregation

Shaking of the plate

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POSSIBLE SOLUTIONS Do not exceed 20 min in papain Make sure that the cultures are started from single-cell suspensions Triturate gently Reduce pipetting Keep tissue in ice cold PBS during microdissection Do not exceed microdissection time in 1h Prepare fresh medium just before cultures Make sure that cell density is 2.5 cells per µl Do not exceed 10 min in accutase Triturate gently Reduce pipetting Prepare fresh medium just before cultures Make sure that cell density is correct Handle plates carefully Avoid frequent monitoring

Fig. 2 Troubleshooting advice

80  C. One day before plating cells, treat glass coverslips of p24 plates with 1 Matrigel® diluted in cold control medium and place at 37 ºC overnight. Do not let dry. Rinse the plates with ultrapure H2O twice before seeding the cells. 9. Do not use an excessive quantity of FluorSave since it may favor apparition of bubbles. Wait for the mounting medium to dry before observing at the microscope. 10. The 5 ml volume of papain solution is sufficient for a good digestion of tissue from one animal. Pooling tissues from more than three animals may require a larger volume of enzymatic solution, in order to avoid poor enzymatic digestion. It is crucial not to exceed the time of digestion (Fig. 2). 11. To prevent tissue and/or cell sticking to the walls of the Pasteur pipette rinse the pipette several times with medium before every dissociation step, or when a new pipette is used. During trituration avoid foaming and bubbles. 12. Make sure that the cultures are started from suspensions of cells dissociated to the single-cell level. We usually obtain approximately 10,000 cells per SEZ and, therefore final seeding density is approximately 5 cells per μl or 2,600 cells per cm2. Cell density is a critical parameter as cells aggregate if density is too high influencing clonality. Additionally neurosphere forming efficiency is influenced by autocrine/paracrine signals, thus different densities affect neurospheres numbers. Is therefore important to maintain low cell density (2.5–5 cells per μl). Movement of dishes also causes aggregation, thus avoid moving the plates during neurosphere formation (Fig. 2).

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13. In primary cultures from adult brain a lot of debris is normally present together with adherent cells. To reduce debris you may increase centrifugation time to 20–30 min. 14. Resuspend the pellet in the Accutase solution by gently shaking to allow the enzyme digest the cell aggregates. It is crucial not to exceed the time of digestion. 15. All subsequent passages are performed after 6–7 days in vitro but, generally, primary spheres seem to require a slightly longer time to grow to the optimal size for passage. Culture can be expanded and propagated indefinitely by repeated subculturing. 16. Overnight incubations can be used. Do not let dry the wells. 17. In order to have a good cell yield do not use neurospheres which have overgrown. 18. Upon EGF removal, neural stem cells differentiate into the three major CNS cell types. Neurons are generated first, followed by astroglia and by oligodendrocytes. 19. Neuronal cells can be detected as early as 2 days after plating. Still detection of astrocytes and oligodendrocytes can be facilitated by the addition of FBS at a final concentration of 2 %, 2 days after plating. Simultaneous detection of the three cell phenotypes is usually possible at 7 days after plating. References 1. Doetsch F (2003) A niche for adult neural stem cells. Curr Opin Genet Dev 13:543–550 2. Ihrie RA, Alvarez-Buylla A (2011) Lake-front property: a unique germinal niche by the lateral ventricles of the adult brain. Neuron 70:674–686 3. Ma DK, Bonaguidi MA, Ming GL, Song H (2009) Adult neural stem cells in the mammalian central nervous system. Cell Res 19:672–682 4. Riquelme PA, Drapeau E, Doetsch F (2008) Brain micro-ecologies: neural stem cell niches in the adult mammalian brain. Philos Trans R Soc Lond B Biol Sci 363:123–137 5. Reynolds BA, Weiss S (1992) Generation of neurons and astrocytes from isolated cells of the adult mammalian central nervous system. Science 255:1707–1710 6. Bickenbach JR, Chism E (1998) Selection and extended growth of murine epidermal stem cells in culture. Exp Cell Res 244:184–195

7. Rietze RL, Valcanis H, Brooker GF, Thomas T, Voss AK, Bartlett PF (2001) Purification of a pluripotent neural stem cell from the adult mouse brain. Nature 412:736–739 8. Singec I, Knoth R, Meyer RP, Maciaczyk J, Volk B, Nikkhah G, Frotscher M, Snyder EY (2006) Defining the actual sensitivity and specificity of the neurosphere assay in stem cell biology. Nat Methods 3:801–806 9. Ferron SR, Andreu-Agullo C, Mira H, Sanchez P, Marques-Torrejon MA, Farinas I (2007) A combined ex/in vivo assay to detect effects of exogenously added factors in neural stem cells. Nat Protoc 2:849–859 10. Pastrana E, Silva-Vargas V, Doetsch F (2011) Eyes wide open: a critical review of sphereformation as an assay for stem cells. Cell Stem Cell 8:486–498

Methods in Molecular Biology (2015) 1212: 113–125 DOI 10.1007/7651_2014_119 © Springer Science+Business Media New York 2014 Published online: 11 September 2014

Generation, Expansion, and Differentiation of Cardiovascular Progenitor Cells from Human Pluripotent Stem Cells Nan Cao, He Liang, and Huang-Tian Yang Abstract Cardiovascular progenitor cells (CVPCs) derived from human embryonic stem cells and human induced pluripotent stem cells represent an invaluable potential source for the study of early embryonic cardiovascular development and stem cell-based therapies for congenital and acquired heart diseases. To fully realize their values, it is essential to establish an efficient and stable differentiation system for the induction of these pluripotent stem cells (PSCs) into the CVPCs and robustly expand them in culture, and then further differentiate these CVPCs into multiple cardiovascular cell types. Here we describe the protocols for efficient derivation, expansion, and differentiation of CVPCs from hPSCs in a chemically defined medium under feeder- and serum-free culture conditions. Keywords: Human pluripotent stem cells, Human embryonic stem cells, Human induced pluripotent stem cells, Cardiovascular progenitor cells, Directed differentiation, Progenitor maintenance, Chemically defined medium, Cardiomyocytes, Smooth muscle cells, Endothelial cells

1

Introduction Pluripotent stem cells (PSCs) including human embryonic stem cells (hESCs) and human induced pluripotent stem cells (hiPSCs) possess unlimited proliferation capacity and can differentiate into derivatives of the three primary germ layers in vitro, including major cell types that form the heart (1–4). Thus they hold tremendous promise for the study of cardiac development and regenerative medicine (5, 6). Heart development is a well-organized process that involves the sequential induction of mesoderm, multipotent cardiovascular progenitor cells (CVPCs), and functional derivatives (7). The process of hPSC differentiation into cardiomyocytes (CMs) is thought to proceed through a similar hierarchy of CVPCs (4, 8). The CVPCs derived from hPSCs are definitively committed cells at earlier developmental stage than CMs and are capable of populating multiple cardiovascular lineages including CMs, smooth muscle cells (SMCs), and endothelial cells (ECs) (9–14). The in vitro

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differentiation of PSCs into CVPCs allows investigators (1) to study human early embryonic developmental processes during the differentiation of PSCs into specialized CVPCs; (2) to identify factors, reagents, and drugs that promote or suppress the formation and maintenance of CVPCs; (3) to understand the characteristics of cardiovascular progenitors and the mechanisms for their selfrenewal; and (4) to evaluate the therapeutic value of hCVPCs for myocardial regeneration. To realize these full values, methodologies for efficient generation, expansion, and differentiation of the CVPCs must be developed. In this chapter, we describe methods to efficiently induce and expand CVPCs from hPSCs. Multipotent CVPCs are generated from monolayer-cultured hESCs and hiPSCs in a chemically defined medium within 3 days by a combined treatment of bone morphogenetic protein 4 (BMP4), glycogen synthase kinase-3 (GSK3) inhibitor CHIR99021, and ascorbic acid. These CVPCs stably self-renew and expand >107 fold in feeder- and serum-free conditions when the differentiation-inducing signals from BMP, GSK3, and Activin/Nodal pathways are synchronizedly inhibited. Furthermore, the protocols are presented for the differentiation of these CVPCs into major cardiovascular lineages including CMs, SMCs, and ECs in vitro when guided by appropriate extrinsic influences.

2 2.1

Materials Cells

1. Human ESC lines H1 and H9 (WiCell). 2. hiPSC line hAFDC-iPS-36 generated from human amniotic fluid-derived cells via ectopic expression of human OCT4, SOX2, KLF4, and C-MYC (15). The use of human cell lines is subject to regulatory guidelines in related country.

2.2

Reagents

1. DMEM/F12 (Life Technologies, cat. no. 11330-057). 2. mTeSR1 (STEMCELL Technologies, cat. no. 05850). 3. RPMI1640 (Life Technologies, cat. no. 11875-119). 4. B27 supplement (Life Technologies, cat. no. 17504-044). 5. B27 supplement without vitamin A (Life Technologies, cat. no. 12587-010). 6. B27 supplement without insulin (Life Technologies, cat. no. 0050129SA). 7. Dulbecco’s PBS (D-PBS) Ca- and Mg-free (Life Technologies, cat. no. 14200067). 8. N2 supplement (Life Technologies, cat. no.17502048).

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9. A83-01 (Stemgent, cat. no. 04-0014). 10. Accutase (Stem Cell Technologies, cat. no. 07920). 11. Ascorbic acid (AA) (Sigma, cat. no. A4544-25G). 12. bFGF (Life Technologies, cat. no. 13256-029). 13. β-mercaptoethanol (Sigma, cat. no. M3148). 14. BMP4 (R&D Systems, cat. no. 314-BP-010). 15. CHIR99021 (Stemgent, cat. no. 04-0004). 16. Dispase (Life Technologies, cat. no. 17105041). 17. Dorsomorphin (Sigma, cat. no. p5499). 18. Growth factor-reduced matrigel (BD Biosciences, cat. no. 354277). 19. L-glutamine (Life Technologies, cat. no. 25030-081). 20. IWR1 (Calbiochem, cat. no. 681669-10MG). 21. Nonessential amino acids (Life Technologies, cat. no. 11140-050). 22. PDGF-BB (R&D Systems, cat. no. 220-BB-010). 23. Penicillin/streptomycin (Life Technologies, cat. no. 15140-122). 24. TGF-β1 (R&D Systems, cat. no. 240-B-010/CF). 25. 1-Thioglycerol (Sigma, cat. no. M6145). 26. VEGF (R&D Systems, cat. no. 493-MV-005/CF). 27. Y-27632 (Calbiochem, cat. no. 688000). 28. Anti-α-Actinin (sarcomeric) clone EA-53 (Sigma, cat. no. A7811). 29. Anti-α-SMA (Sigma, cat. no. A2547). 30. Anti-GATA4 (Santa Cruz Biotechnology, cat. no. sc-25310). 31. Anti-ISL1 (Developmental Studies Hybridoma Bank, clone 39.4D5). 32. Anti-MEF2C (Cell Signaling, cat. no. 5030). 33. Anti-MESP1/2 (Aviva ARP39374_P050).

Systems

Biology,

cat.

no.

34. Anti-PECAM1 clone 9G11 (R&D System, cat. no. BBA7) PE-conjugated Anti-SSEA1 (eBioscience, cat. no. 12-8813-42). 2.3 Cell Culture Media (See Note 1)

1. Basal CVPC induction medium (CIM) (500 mL): in a sterile environment, mix 480 mL of DMEM/F12, 10 mL of B27 supplement without vitamin A, 5 mL of 200 mM L-glutamine, 5 mL penicillin/streptomycin, 400 μM 1-thioglycerol. 2. CIM (500 mL): in a sterile environment, mix 480 mL of DMEM/F12, 10 mL of B27 supplement without vitamin A, 5 mL of 200 mM L-glutamine, 5 mL penicillin/streptomycin,

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400 μM 1-thioglycerol, 50 μg/mL AA, 25 ng/mL BMP4, and 3 μM GSK3 inhibitor CHIR99021. 3. Basal CVPC propagation medium (CPM) (500 mL): in a sterile environment, mix 470 mL of DMEM/F12, 10 mL of B27 supplement without vitamin A, 5 mL of N2 supplement, 5 mL of nonessential amino acids, 5 mL of 200 mM L-glutamine, 5 mL penicillin/streptomycin, 400 μM 1-thioglycerol, 0.1 mM β-mercaptoethanol. 4. CPM (500 mL): in a sterile environment, mix 470 mL of DMEM/F12, 10 mL of B27 supplement without vitamin A, 5 mL of N2 supplement, 5 mL of nonessential amino acids, 5 mL of 200 mM L-glutamine, 5 mL penicillin/streptomycin, 400 μM 1-thioglycerol, 0.1 mM β-mercaptoethanol, 3 μM CHIR99021, 2 μM BMP inhibitor dorsomorphin, and 0.5 μM Activin/Nodal inhibitor A83-01. 5. Cardiac differentiation medium 1 (CDM1) (500 mL): in a sterile environment, mix 480 mL of RPMI1640, 10 mL of B27 supplement without insulin, 5 mL of 200 mM L-glutamine, 5 mL penicillin/streptomycin, 10 ng/mL BMP4, and 5 μM Wnt antagonist IWR1. 6. Cardiac differentiation medium 2 (CDM2) (500 mL): in a sterile environment, mix 480 mL of RPMI1640, 10 mL of B27 supplement, 5 mL of 200 mM L-glutamine, and 5 mL penicillin/streptomycin. 7. SMC differentiation medium (SDM) (500 mL): in a sterile environment, mix 480 mL of DMEM/F12, 10 mL of B27 supplement without vitamin A, 5 mL of 200 mM L-glutamine, 5 mL penicillin/streptomycin, 400 μM 1-thioglycerol, 10 ng/ mL PDGF-BB, and 2 ng/mL TGF-β1. 8. EC differentiation medium (EDM) (500 mL): in a sterile environment, mix 480 mL of DMEM/F12, 10 mL of B27 supplement without vitamin A, 5 mL of 200 mM L-glutamine, 5 mL penicillin/streptomycin, 400 μM 1-thioglycerol, 50 ng/ mL VEGF, and 10 ng/mL bFGF. 2.4

Equipment

1. Flow cytometry FACS Aria (Becton Dickinson). 2. Humidified tissue culture incubator (Thermo, 37  C, 5 % CO2). 3. Inverted TS100).

phase-contrast

microscope

(Nikon,

ECLIPSE

4. Conical tubes (15 and 50 mL; Corning Biosciences, cat. nos. 430791 and 430829). 5. Flow round-bottom tube (5 mL; BD Biosciences, cat. no. 352052).

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6. Microcentrifuge tube (1.5 mL; Axygen, cat. no. MCT-150-C). 7. Plates (6-, 12-, 24-, and 96-well; Nunc, cat. nos. 140675, 150628, 142475, and 165306). 8. Serological pipettes (1, 5, 10, and 25 mL; Corning, cat. nos. 4011, 4050, 4100, and 4250). 9. Stericup filtration system (Millipore, cat. no. SCGPU02RE, 250 mL). 10. Stericup filtration system (Millipore, cat. no. SCGPU05RE, 500 mL).

3

Methods

3.1 Preparation of Matrigel-Coated Plates

1. In a sterile hood, add 23 mL of cold (4  C) DMEM/F12 to a 50-mL conical tube and keep it cold by placing it on ice. 2. Remove one matrigel aliquot (2 mg) from the 80  C freezer, and add 1 mL of cold DMEM/F12 to it. Gently pipette the Matrigel solution with a P1000 tip to thaw and dissolve the Matrigel. 3. After that, immediately transfer the Matrigel solution to the 50-mL conical tube that contains 23 mL of cold DMEM/F12, mix them well with pipette, and then add Matrigel in DMEM/ F12 at a volume of 1 mL/well for 6-well plates, 0.5 mL/well for 12-well plates, 250 μL/well for 24-well plates, or 100 μL/ well for 96-well plates. 4. The Matrigel-coated plates are stored in a freezer at 4  C overnight before use and they can be stored up to 1 week (see Note 2).

3.2 Thawing and Recovering hPSCs in Feeder-Free Culture

1. Pre-warm the required volume of mTeSR1 medium at room temperature until it is no longer cool to the touch. 2. Take a Matrigel-coated 6-well plate from 4  C and put it at 37  C for at least 1 h. 3. Remove the vial of hPSCs from liquid nitrogen storage using metal forceps and immerse the vial in a 37  C water bath without submerging the cap. Swirl the vial gently. When only an ice crystal remains, remove the vial from the water bath. 4. Spray the vial with 70 % ethanol and place it in tissue culture hood. Pipet cells gently into a sterile 15-mL conical tube, add 4 mL of pre-warmed mTeSR1 medium drop-wise using a 5-mL sterile pipette. While adding the medium, gently shake the tube to mix the PSCs. This reduces osmotic shock to the cells. 5. Centrifuge the cells at 200  g for 5 min. Aspirate and discard the supernatant with a sterilized Pasteur pipette. Aspirate the liquid from the wells of the Matrigel-coated plate.

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6. Resuspend the cell pellet in 2 mL mTeSR1 medium. Slowly add 2 mL of cell suspensions into each well of the Matrigel-coated 6-well plate. Place the plate gently into the incubator. Move the plate in several quick, short, back-and-forth, and side-to-side motions to further disperse cells across the surface of the wells. 7. The next day, replace the spent medium with fresh mTeSR1 medium. 8. Replace the medium daily thereafter until the cells are approximately 80–90 % confluent (see Note 3). 3.3 Passaging hPSCs (See Note 4)

1. Pre-warm the required volume of mTeSR1 medium at room temperature until it is no longer cool to the touch. 2. Take a Matrigel-coated 6-well plate from 4  C and put it at 37  C for at least 1 h. 3. Aspirate the old medium from the vessel containing PSCs with a Pasteur pipette, and rinse the vessel with D-PBS. 4. Add 1 mL of 1 mg/mL Dispase solution (room temperature) to each well and incubate the plate at 37  C for 7 min. 5. Aspirate the Dispase solution with a Pasteur pipet. Remove the Dispase carefully without disturbing the attached cell layer. 6. Gently wash the cells with 2 mL of DMEM/F12, aspirate the medium, and repeat twice. 7. Add 2 mL pre-warmed mTeSR1 medium to the well and remove the cells from the well(s) by gently scraping them into small clusters. 8. After the cells are removed from the surface of the well, pool the contents of the well into a sterile conical tube containing 12 mL mTeSR1 medium. Gently mix 2–4 times using a 5 mL pipette and seed 2 mL of the cell suspension into each well of a Matrigel-coated 6-well plate (split ratio of 1:6 is performed here). 9. Put the plate back into the incubator. Move the plate in several quick, short, back-and-forth, and side-to-side motions to further disperse cells across the surface of the wells. 10. The next day, replace the spent medium with fresh mTeSR1 medium; change medium daily until cells are 80–90 % confluent.

3.4 Induction of CVPCs from hPSCs

1. Pre-warm the required volume of CIM at room temperature until it is no longer cool to the touch. 2. Take a Matrigel-coated plate from 4  C and put it at 37  C for at least 1 h. 3. Take hPSCs cultured on Matrigel-coated 6-well plates in mTeSR1 medium at 80–90 % confluence (see Note 5).

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Aspirate the medium and rinse the vessel twice with D-PBS. Add 1 mL of room-temperature Accutase to each well. Put the plate in an incubator and wait for exactly 5 min (see Note 6). 4. Add 2 mL of mTeSR1 medium into each well and pool all of the cells in a 15-mL conical tube. Count the total cell number with a hemocytometer. Centrifuge the cells at 200  g for 5 min at room temperature. 5. Aspirate the supernatant, resuspend the cells in CIM + 5 μM Y27632 at a cell density of 0.25 million cells per mL, and plate them onto Matrigel-coated culture plates at a density of 5  104 cells/cm2 (see Note 7). 6. Put the plate back into the incubator. Move the plate in several quick figure eight motions to disperse cells across the surface of the wells. This time point corresponds to day 0 of differentiation. 7. Twenty-four hours later, aspirate the old medium and then replace it with room-temperature CIM. Place the plate in a 37  C, 5 % CO2 incubator for 2 days without changing the medium. 8. The monolayer-cultivated cells were harvested at differentiation day 3 for further examination or passaging. 9. The CVPCs exhibit homogenous morphology and express several CVPC markers including SSEA1, MESP1/2, ISL1, GATA4, and MEF2C but not pluripotent markers, which can be monitored by fluorescence-activated cell sorter (FACS), immunostaining analyses, and quantitative reverse transcription PCR (Fig. 1) as previously described (12). 3.5 Propagation of CVPCs from hPSCs

1. Pre-warm the required volume of CPM at room temperature until it is no longer cool to the touch. 2. Take a Matrigel-coated 6-well plate from 4  C and put it at 37  C for at least 1 h (see Note 8). 3. When CVPCs are induced on day 3 (typically 80–90 % confluence) (see Note 9), aspirate the medium and rinse the vessel twice with D-PBS. Add 1 mL of room-temperature Accutase to each well of a 6-well plate. Place the plate in an incubator and wait for exactly 5 min. 4. Add 2 mL of CPM into each well and pool all of the cells in a 15-mL conical tube. Centrifuge the cells at 200  g for 5 min at room temperature. 5. Remove the supernatant and resuspend in 6 mL CPM + 5 μM Y27632 (see Note 10). Gently mix 5–10 times using 5 mL pipette and seed 2 mL of the cell suspension into each well of a Matrigel-coated 6-well plate (split ratio of 1:3 is performed here) (see Note 11).

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a

b CIM

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89.7 ± 1.9

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15

86.8 ± 2.2

H9

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94.4 ± 2.4

hiPSC

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86.5 ± 2.2

hiPSC

3

92.4 ± 4.2

0 1 3 Differentiation time (days)

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Fig. 1 Induction of cardiovascular progenitor cells (CVPCs) from human PSCs. (a) An outline of the differentiation protocol. CIM, CVPC induction medium; Y, Y27632. (b) Intracellular flow cytometry analysis of induction efficiency of CVPCs in various hPSC lines. (c) Immunofluorescence analysis showing the expression of CVPC markers in cells at differentiation day 3 following CIM treatment. Scale Bars ¼ 25 μm. (d) qPCR analysis showing the downregulation of pluripotency markers and upregulation of CVPC markers during cardiovascular induction (n ¼ 3) (reproduced from (12) with permission from Cell Res)

6. Place the plate back into the CO2 incubator. Move the plate in several quick, short, back-and-forth, and side-to-side motions to further disperse cells across the surface of the wells. 7. The medium is renewed daily until cells reach 80–90 % confluence (typically 3–4 days) that is ready for passage. 8. The expanded CVPCs can actively proliferate and form roundshaped colonies with clear edge during expansion. They uniformly express primitive CVPC markers including SSEA1, MESP1/2, and ISL1 (Fig. 2) as previously described (12).

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b

Basal CPM

# Cells

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93.7

6.8

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MESP1/2

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MESP1/2

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Fig. 2 Maintenance of hPSC-derived CVPCs. (a) Phase contrast images (left panels) and percentage of MESP1/ 2+ cells analyzed by intracellular flow cytometry (right panels. CPM, CVPC propagation medium; Basal CPM, CPM without CHIR99021, dorsomorphin and A83-01). (b) Representative images showing the typical morphology (left panels), and the SSEA1 and MESP1 expression analyzed by intracellular flow cytometry (middle and right panels). The control sample used in this assay was a 1:1:1 mixture from all three types of cells examined. (c) Representative images of the ISL1 and MESP1 expression analyzed by immunostaining of CVPC colonies at passage 15. Scale bar ¼ 100 μm (reproduced from (12) with permission from Cell Res) 3.6 Differentiation of CVPCs into CMs

1. Pre-warm the required volume of CDM1 at room temperature until it is no longer cool to the touch. 2. Take a Matrigel-coated 6-well plate from 4  C and put it at 37  C for at least 1 h. 3. Aspirate the medium containing CVPCs with a Pasteur pipette and rinse the vessel twice with D-PBS. Add 1 mL of roomtemperature Accutase to each well of a 6-well plate. Place the plate in an incubator and wait for exactly 5 min. 4. Add 2 mL of CDM1 into each well and pool all of the cells in a 15-mL conical tube. Count the total cell number with a

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hemocytometer. Centrifuge the cells at 200  g for 5 min at room temperature. 5. Aspirate the supernatant, resuspend the cells in CDM1 at a cell density of four million cells/mL and plate them onto Matrigelcoated culture plates at a density of 4  105 cells/cm2. 6. Put the plate back into the incubator. Move the plate in several quick, short, back-and-forth, and side-to-side motions to further disperse cells across the surface of the wells. 7. Cultivate the cells for 3 days without changing the medium. 8. Three days later, aspirate the CDM1 medium and add 3 mL of room-temperature CDM2 medium to each well of a 6-well plate. 9. Cultivate the cells in CDM2 for another 9 days. Medium is renewed every 2–3 days. 10. After 12 days of differentiation, majority of the cells should exhibit hallmarks of CMs including spontaneous beating and cardiac-specific marker expression (Fig. 3) as previously described (12). 3.7 Differentiation of CVPCs into SMCs or ECs

1. Pre-warm the required volume of SDM or EDM at room temperature until they are no longer cool to the touch. 2. Take Matrigel-coated 6-well plates from 4 C and put them at 37  C for at least 1 h. 3. Aspirate the medium containing CVPCs with a Pasteur pipette and rinse the vessel twice with D-PBS. Add 1 mL of roomtemperature Accutase to each well of a 6-well plate. Put the plate in the incubator and wait for exactly 5 min. 4. Add 2 mL of SDM or EDM into each well and pool all of the cells in a 15-mL conical tube. Count the total cell number with a hemocytometer. Centrifuge the cells at 200  g for 5 min at room temperature. 5. Aspirate the supernatant, resuspend the cells in SDM or EDM at a cell density of 0.1 million cells/mL, and plate them onto Matrigel-coated culture plates at a density of 104 cells/cm2 in SDM or EDM respectively. 6. Put the plate back into the incubator. Move the dish in several quick figure eight motions to disperse cells across the surface of the wells. 7. Cultivate the cells for 12 days. Medium is renewed every 2–3 days. 8. After 12 days of differentiation, majority of the cells should express SMC marker α-SMA or EC marker PECAM1 respectively (Fig. 3) as previously described (12).

hPSC-Derived Cardiovascular Precursor Cells

PDGF-BB+TGF β 1 in Basal CIM

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Fig. 3 In vitro differentiation potential of CVPCs. (a) An outline of the conditions for inducing CVPC differentiation. (b) Differentiation potential of P15 CVPCs into cardiomyocytes, smooth muscle cells, and endothelial cells determined by immunostaining analyses for Nkx2-5 (green), cTnT (red or green), α-actinin (red) and merged Nkx-2.5 with cTnT in cardiomyocytes, α-SMA (red) and SM-MHC (green) in smooth muscle cells, and PECAM1 (red), CDH5 (red), and CD34 (green) in endothelial cells derived from CVPCs at differentiation day 12. Scale Bars ¼ 50 μm (reproduced from (12) with permission from Cell Res)

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Notes 1. All the medium used in this protocol can be stored at 70  C for more than 3 months, but they are not stable in 4  C and should be used immediately. 2. Different batch of Matrigel may affect the induction and maintenance of CVPCs and should be tested before use. 3. Never let the hPSC colonies reach confluence and they may start to differentiate. 4. Pluripotent stem cell culture can be maintained by repeating passage before moving to the next step for CVPC induction. The induction efficiency of CVPCs is not significantly changed by using hPSCs with passages range from 20 to 60. 5. HPSCs are cultured on Madrigal in mTeSR1 for at least two passages. The homogeneous maintenance of hESCs in undifferentiated state is critical for the efficient induction of CVPCs.

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6. Incubation should be carried out for exactly 5 min under the hood. This is important to digest the hPSCs into single cells that is critical for the efficient CVPC without decreasing cell variability. 7. ROCK inhibitor Y27632 at 5 μM is helpful for increasing cell viability after thawing but it is better not over 24 h. 8. Different batch of Madrigal may affect the propagation efficiency of CVPCs and should be tested before use. 9. The high efficiency in CVPC induction is critical for their selfrenewal and propagation. Make sure the expression of CVPC markers in majority of the cells (>90 %) before long-term expansion (12). If not, the CVPCs can be sorted out using anti-SSEA-1 antibody-conjugated magnetic beads as previously described (11, 16). 10. During the initial five passages, overnight treatment of 5 μM Y27632 is used to improve cell survival, but it is not required in the following passages. 11. Split ratio for the CVPCs can be variable (1:2–1:5) between lines and passage times. A general rule is to observe the last split ratio and adjust the ratio according to the appearance of the CVPC colonies. If the cells are actively proliferating, increase the split ratio.

Acknowledgments The work was supported by grants of the Strategic Priority Research Program of CAS (XDA01020204), National Natural Science Foundation of China (31030050), National Basic Research Program of China (2011CB965300), National Science and Technology Project of China (2012ZX09501-001-001), and CAS (GJHZ1225). We are grateful to WiCell Research Institute for providing us H1 and H9 hESC lines and to Dr. Ying Jin (Institute of Health Sciences, China) for kindly providing the hiPSC line hAFDC-iPS-36. We also thank the members from our laboratory for valuable discussions. References 1. Thomson JA, Itskovitz-Eldor J, Shapiro SS et al (1998) Embryonic stem cell lines derived from human blastocysts. Science 282:1145–1147 2. Takahashi K, Tanabe K, Ohnuki M et al (2007) Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131:861–872

3. Park IH, Lerou PH, Zhao R et al (2008) Generation of human-induced pluripotent stem cells. Nat Protoc 3:1180–1186 4. Burridge PW, Keller G, Gold JD et al (2012) Production of de novo cardiomyocytes: human pluripotent stem cell differentiation and direct reprogramming. Cell Stem Cell 10:16–28

hPSC-Derived Cardiovascular Precursor Cells 5. Murry CE, Keller G (2008) Differentiation of embryonic stem cells to clinically relevant populations: lessons from embryonic development. Cell 132:661–680 6. Blin G, Neri T, Stefanovic S et al (2010) Human embryonic and induced pluripotent stem cells in basic and clinical research in cardiology. Curr Stem Cell Res Ther 5:215–226 7. Noseda M, Peterkin T, Simoes FC et al (2011) Cardiopoietic factors: extracellular signals for cardiac lineage commitment. Circ Res 108:129–152 8. Mummery CL, Zhang J, Ng ES et al (2012) Differentiation of human embryonic stem cells and induced pluripotent stem cells to cardiomyocytes: a methods overview. Circ Res 111:344–358 9. Bu L, Jiang X, Martin-Puig S et al (2009) Human ISL1 heart progenitors generate diverse multipotent cardiovascular cell lineages. Nature 460:113–117 10. Yang L, Soonpaa MH, Adler ED et al (2008) Human cardiovascular progenitor cells develop from a KDR+ embryonic-stem-cell-derived population. Nature 453:524–528 11. Blin G, Nury D, Stefanovic S et al (2010) A purified population of multipotent

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cardiovascular progenitors derived from primate pluripotent stem cells engrafts in postmyocardial infarcted nonhuman primates. J Clin Invest 120:1125–1139 12. Cao N, Liang H, Huang J et al (2013) Highly efficient induction and long-term maintenance of multipotent cardiovascular progenitors from human pluripotent stem cells under defined conditions. Cell Res 23:1119–1132 13. Elliott DA, Braam SR, Koutsis K et al (2011) NKX2-5(eGFP/w) hESCs for isolation of human cardiac progenitors and cardiomyocytes. Nat Methods 8:1037–1040 14. Kattman SJ, Witty AD, Gagliardi M et al (2011) Stage-specific optimization of activin/nodal and BMP signaling promotes cardiac differentiation of mouse and human pluripotent stem cell lines. Cell Stem Cell 8:228–240 15. Li C, Zhou J, Shi G et al (2009) Pluripotency can be rapidly and efficiently induced in human amniotic fluid-derived cells. Hum Mol Genet 18:4340–4349 16. Leschik J, Stefanovic S, Brinon B et al (2008) Cardiac commitment of primate embryonic stem cells. Nat Protoc 3:1381–1387

Methods in Molecular Biology (2015) 1212: 127–140 DOI 10.1007/7651_2014_92 © Springer Science+Business Media New York 2014 Published online: 20 July 2014

A Practical Guide for the Isolation and Maintenance of Stem Cells from Tendon Pauline Po Yee Lui Abstract Stem cells are unspecialized cells that can self-renew and have the ability to develop into cells of highly specialized functions. The study of stem cells holds enormous promise in the medical field ranging from their uses in cell therapies to their uses for greater understanding of tissue development and disease pathologies. Stem cells have been isolated from tendon tissue recently. These tendon-derived stem cells (TDSCs) are particularly relevant for tendon repair and the study of the potential roles of stem cells in tendon pathology as they are isolated from tendon tissues. This paper aims to describe the step-by-step protocol and the practical tips for the isolation and verification of stem cell characteristics of TDSCs. The cell seeding density and hence cell–cell contact has a significant impact on the isolation and expansion of TDSCs. Hence, I also describe our established protocol for the determination of the optimal seeding density for TDSC isolation and culture. Keywords: Tendon-derived stem/progenitor cells, Mesenchymal stem cell isolation, Seeding density, Stem cell culture, Stem cell characterization

1

Introduction Tendon transmits forces from muscle to bone and provides joint function. Tendon injuries are common (1). Both acute and chronic tendon injuries are difficult to manage (2). After acute tendon injury, tendon heals slowly and scar tissue, which has poor tissue quality and tensile mechanical properties, is formed (3). Chronic tendon injuries associated with tendon overuse is presented with defective healing with increase in cellularity and vascularity, increased deposition of proteoglycans, matrix degeneration, and tissue metaplasia (4, 5). The injured tendon is presented with swelling, local tenderness, and pain. As the pathogenesis of chronic tendon injuries is not clear, there is no effective treatment. Stem cells which have self-renewal and multi-lineage differentiation potential have received intense attention recently both as a cell source for tissue repair (6) and as the target cell in disease pathogenesis (7). Recently, stem cells with self-renewal and multi-lineage

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differentiation potential have been isolated from tendon tissues of different species including human (8), mouse (8), rat (9), rabbit (10), and horse (11) as well as from different origins of tendon tissue including hamstring tendon (12), patellar tendon (13), flexor tendon (9), Achilles tendon (10), long head of the biceps (14), and supraspinatus tendon (14). Our group named these stem cells isolated from tendon tissues as tendon-derived stem cells (TDSCs) to indicate the tissue from which these stem cells were isolated (9). The TDSCs are particularly relevant for tendon repair (15, 16) and the study of the potential roles of stem cells in tendon pathology (4, 16) as they are isolated from tendon tissues. This paper aimed to present the step-by-step protocol and the practical tips for the isolation and verification of stem cell characteristics of TDSCs. Since mesenchymal stem cells (MSCs) are clonogenic cells, the initial cell seeding density and hence cell–cell contact has a significant impact on their isolation. The seeding of TDSCs at lower cell density was found to result in a more purified culture and higher yield (9, 17, 18). The passaging density also affects TDSC’s proliferative potential and TDSCs seeded at a lower density was reported to expand more efficiently (9, 18). These findings are consistent with the previous reports that cell density and hence cell–cell interaction controlled the differentiation fate of cortical stem cells (19). Moreover, the presence of differentiated cells in the MSC culture was reported to induce spontaneous MSC differentiation (20–22). Hence, I also describe our established protocol for the determination of the optimal seeding density for TDSC isolation and culture.

2

Materials

2.1 Reagents for the Isolation and Culture of TDSCs

1. Phosphate buffered saline (PBS), 1 L, sterile: Dissolve 80 g of NaCl, 2 g of KCl, 14.4 g of Na2HPO4, 2.4 g of KH2PO4 into 1 L of Milli-Q H2O to make a 10 stock solution. Dilute 1:9 with Milli-Q H2O to prepare the working solution. Adjust the pH of the solution to 7.4 by hydrochloric acid and sterilized by autoclaving. Store at room temperature. 2. Complete culture medium, 1 L, sterile: Dissolve low-glucose Dulbecco modified Eagle medium (LG-DMEM, Gibco, Invitrogen corporation, Carlsbad, USA) powder in 1 L of Milli-Q H2O. Adjust the pH of the solution to 7.0–7.2 and filter with a 0.22 μm filter (Millipore, Billerica, MA, USA). Supplement the plain medium with final concentration of 10 % fetal calf serum (FBS), 50 μg/ml penicillin, 50 μg/ml streptomycin, and 100 μg/ml neomycin (all from Invitrogen Corporation, Carlsbad, USA). Store at 4  C.

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3. 3 mg/ml type I collagenase from Clostridium histolyticum, 100 ml, freshly prepared, sterile: Dissolve 0.03 g of type I collagenase from Clostridium histolyticum (Sigma-Aldrich, St. Louis, MO, USA) in 100 ml of sterile complete culture medium. Filter with a 0.22 μm filter (Millipore, Billerica, MA, USA). 2.2 Reagents for Determining the Optimal Plating Density for the Isolation of TDSCs

1. 0.5 % crystal violet, 100 ml: Dissolve 0.5 g of crystal violet (Sigma-Aldrich, St. Louis, MO, USA) in 100 ml of double distilled water (ddH2O). Store at room temperature.

2.3 Reagents for Osteogenic Differentiation Assay

1. 0.1 mM dexamethasone, 100 ml, sterile: Dissolve 0.00392 g of dexamethasone (D4902, Sigma-Aldrich, St. Louis, MO, USA) in 100 ml of ddH2O and filter with a 0.22 μm filter (Millipore, Billerica, MA, USA). Store at 20  C. 2. 50 mM ascorbic acid, 10 ml, sterile: Dissolve 0.08806 g of ascorbic acid (A0278, Sigma-Aldrich, St. Louis, MO, USA) in 10 ml of sterile PBS and filter with a 0.22 μm filter (Millipore, Billerica, MA, USA). Store at 20  C. 3. 1 M β-glycerophosphate, 10 ml, sterile: Dissolve 2.16 g of βglycerophosphate (G6736, Sigma-Aldrich, St. Louis, MO, USA) in 10 ml of sterile PBS and filter with a 0.22 μm filter (Millipore, Billerica, MA, USA). Store at 20  C. 4. Osteogenic medium, sterile: Complete culture medium supplemented with final concentration of 1 nM dexamethasone, 50 μM ascorbic acid, and 20 mM β-glycerophosphate. Store at 4  C. 5. 0.5 % Alizarin red S, 100 ml: Dissolve 0.5 g of Alizarin red S power (Sigma-Aldrich, St. Louis, MO, USA) in 100 ml of ddH2O and adjust the pH to 4.2. Store at room temperature. 6. Cetylpyridinium chloride (CPC), 100 ml: Dissolve 10 g of CPC in 100 ml of 10 mM sodium phosphate (10 % w/v) (SigmaAldrich, St. Louis, MO, USA) and adjust the pH to 7.0. Store at room temperature.

2.4 Reagents for Chondrogenic Differentiation Assay

1. 10 μg/ml transforming growth factor beta 3 (TGF-β3), 0.2 ml: Dissolve 2 μg of TGF-β3 (243-B3-002, R&D Systems, Inc., Minneapolis, MN, USA) in 0.2 ml of sterile 4 mM HCl containing at least 0.1 % BSA. Store at 20  C. 2. 10 μg/ml bone morphogenetic protein-2 (BMP-2), 1 ml: Dissolve 10 μg of BMP-2 (355-BM-010, R&D Systems, Inc., Minneapolis, MN, USA) in 1 ml of sterile 4 mM HCl containing at least 0.1 % BSA. Store at 20  C.

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3. 50 mg/ml ascorbate-2-phosphate, 10 ml, sterile: Dissolve 500 mg of ascorbate-2-phosphate (A8960, Sigma-Aldrich, St. Louis, MO, USA) in 10 ml of sterile PBS and filter with a 0.22 μm filter (Millipore, Billerica, MA, USA). Store at 20  C. 4. 40 mg/ml proline, 10 ml, sterile: Dissolve 400 mg of L-proline (P0380, Sigma-Aldrich, St. Louis, MO, USA) in 10 ml of sterile PBS and filter with a 0.22 μm filter (Millipore, Billerica, MA). Store at 20  C. 5. 100 mg/ml pyruvate, 10 ml, sterile: Dissolve 100 mg sodium pyruvate (Sigma-Aldrich, St. Louis, MO, USA) in 1 ml of sterile Milli-Q H2O. Store at a range from 20  C to 4  C. Dilute to 100 μg/ml when use. 6. Chondrogenic medium, 100 ml, sterile: LG-DMEM supplemented with final concentration of 10 ng/ml TGF-β3, 500 ng/ml BMP-2, 10 7 M dexamethasone, 50 μg/ml ascorbate-2-phosphate, 40 μg/ml proline, 100 μg/ml pyruvate and 1:100 diluted ITS + Premix (6.25 mg/ml insulin, 6.25 mg/ml transferrin, 6.25 mg/ml selenous acid, 1.25 mg/ml bovine serum albumin, 5.35 mg/ml linoleic acid) (Becton Dickinson, Franklin Lakes, USA). Store at 20  C. 7. Slide coating: Wash glass slides in acetone for 5 min. Immerse the glass slides in 2 % 3-aminopropyl-triethoxy-silane (SigmaAldrich, St. Louis, MO, USA) in acetone for 15 min. Wash the coated slides with distilled H2O and dry in an oven at 70  C until dry. Store at room temperature. 8. 0.1 % Safranin O, 500 ml: Dissolve 0.5 g of Safranin O (SigmaAldrich, St. Louis, MO, USA) in 500 ml of distilled H2O. Store at room temperature. 9. 0.1 % fast green, 500 ml: Dissolve 0.5 g of fast green (Merck Millipore, Darmstadt, Germany) in 500 ml of distilled H2O. Store at room temperature. 2.5 Reagents for Adipogenic Differentiation Assay

1. 50 mM isobutylmethylxanthine, 10 ml, sterile: Dissolve 0.11112 g of isobutylmethylxanthine (I5879, Sigma-Aldrich, St. Louis, MO, USA) in 10 ml of sterile DMSO and filter with a 0.22 μm filter (Millipore, Billerica, MA, USA). Store at 20  C. 2. 50 mM indomethacin, 10 ml, sterile: Dissolve 0.178895 g of indomethacin (I7378, Sigma-Aldrich, St. Louis, MO, USA) in 10 ml of sterile methanol and filter with a 0.22 μm filter (Millipore, Billerica, MA, USA). Store at 20  C. 3. 1 mg/ml insulin, 10 ml, sterile: Dissolve 10 mg of insulin (I5500, Sigma-Aldrich, St. Louis, MO, USA) in 10 ml of diluted hydrochloric acid (diluted to pH 2–3 with sterile PBS (pH 7.4)) and filter with a 0.22 μm filter (Millipore, Billerica, MA, USA). Store at 20  C.

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4. Adipogenic medium, 100 ml, sterile: Complete culture medium supplemented with final concentration of 500 nM dexamethasone, 0.5 mM isobutylmethylxanthine, 50 μM indomethacin, and 10 μg/ml insulin (all from Sigma-Aldrich, St. Louis, MO, USA). Store at 4  C. 5. 0.3 % Oil Red-O solution, 100 ml, freshly prepared: Dissolve 0.3 g of Oil Red-O powder in 100 ml of 100 % propylene glycol with heating to 95–100  C. Filter the solution with coarse filter paper (i.e., 25 μm filter paper) while the solution is still warm. Allow the solution to stand overnight at room temperature before use.

3

Methods

3.1 Isolation and Culture of TDSCs

1. Store the tendon tissue for TDSC isolation in sterile PBS with 10 % PSN (i.e., 500 μg/ml penicillin, 500 μg/ml streptomycin, and 1,000 μg/ml neomycin) for 5 min (see Notes 1, 2). 2. Wash the tendon tissue twice with PBS before TDSC isolation. 3. Add 1 ml 0.05 % trypsin–EDTA to cover and digest the tendon tissue for 5 min at 37  C, 5 % CO2. 4. Add complete culture medium to terminate the trypsin–EDTA action. 5. Vortex vigorously for 30 s. 6. Remove the complete culture medium and wash the tendon tissue twice with PBS. 7. Mincethetendontissueintosmallpieces(1mm  1mm  1mm) and digest it with 3 ml 3 mg/ml type I collagenase from Clostridium histolyticum for 2 h at 37  C, 5 % CO2. Vortex vigorously for 30 s at room temperature every 30 min of digestion (see Note 3). 8. Obtain the released cell using a 70 μm cell strainer (Becton Dickinson, Franklin Lakes, NJ, USA). 9. Wash the released cells with PBS twice and resuspend them in complete culture medium. 10. Count the number of nucleated cells. 11. Plate the isolated nucleated cells at an optimal low cell density and culture them at 37  C, 5 % CO2 to form colonies for the isolation of TDSCs (see Note 4). 12. Wash cells twice with PBS to remove the non-adherent cells at day 2 after initial plating and replace with fresh complete culture medium (see Note 5). 13. Trypsinize the cells with 0.05 % trypsin–EDTA (about 0.5 ml for a 100 mm dish) at day 7–10 when the cells reach confluence. The cells at this stage are considered to be at passage 0 (see Note 6).

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14. Replace with fresh complete culture medium every 3 days (see Note 7). 15. Subculture the TDSCs at 500 cells/cm2 when they reach 80–90 % confluence (see Notes 8, 9). 16. Determine the surface marker expression and multi-lineage differentiation potential of the isolated TDSCs to confirm their stem cell characteristics (see Note 10). 3.2 Determination of the Optimal Plating Density for the Isolation of TDSCs

1. Plate the nucleated cells isolated from tendon at 5, 50, 500, and 5,000 cells/cm2 in complete culture medium in 20-cm2 dishes in triplicate. 2. Culture the cells for 7–10 days at 37  C, 5 % CO2. 3. Replace with fresh complete culture medium every 3–4 days. 4. Remove the culture medium and wash the cells twice with PBS. 5. Stain the cells with 2 ml 0.5 % crystal violet for 15 min at room temperature. 6. Count the number of cell colonies per dish at different seeding densities (see Note 11). 7. The optimal plating density for TDSC isolation is the plating density at which the greatest number of colonies per nucleated cell is obtained.

3.3 Determination of Stem Cell-Related Surface Marker Expression

1. Wash TDSCs twice with PBS and trypsinize the cells with 0.05 % trypsin–EDTA. 2. Add complete culture medium to inhibit the activity of trypsin–EDTA and centrifuge the cell suspension at 350  g for 5 min at room temperature. 3. Wash cells twice with PBS and resuspend the cells in PBS (1 % BSA) at 2  105/ml for 15 min at 4  C. 4. Add primary antibodies of target cell surface marker at the dilution recommended by the company to 100 μl of cell suspension for 15 min to 1 h at 4  C (see Note 12). 5. Wash the cell suspension with ice-cold PBS (with 1 % BSA) once to remove the unbound antibodies. 6. Add secondary antibodies conjugated to a fluorescent probe at the dilution recommended by the company to 100 μl of cell suspension for binding detection for 15 min to 1 h at 4  C. 7. Wash the cell suspension with PBS at 400  g for 5 min to remove the unbound antibodies. 8. Resuspend the cells in 500 μl of ice-cold PBS (with 1 % paraformaldehyde). 9. Analyze the sample by flow cytometer (see Note 13).

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1. Seed TDSCs at 4  103 cells/cm2 in complete culture medium in a 6-well culture plate at 37  C, 5 % CO2 until the cells reached confluence. 2. Replace with either 2 ml of complete culture medium or osteogenic medium and incubate at 37  C, 5 % CO2 for 14–21 days for calcium nodule formation. 3. Replace with fresh complete culture medium or osteogenic medium every 3.5 days. 4. Assess the formation of calcium nodules by Alizarin red S staining. 5. Wash the cells twice with PBS. 6. Fix the cells with 2 ml 70 % ethanol for 10 min. 7. Stain the cells with 2 ml of 0.5 % of Alizarin red S for 30 min. 8. Remove Alizarin red S and wash the cells with ddH2O. 9. View the calcium nodules under light microscope (see Note 14). 10. Add 2 ml of 10 % (w/v) CPC to the cells for 15 min at room temperature with gentle shaking to elute the dye. 11. Pipette the solution several times to ensure that the Alizarin red S dye is in the solution. 12. Transfer the solution to a 1.5-ml eppendorf tube. 13. Measure the dye color intensity at OD 562 nm with 10 % (w/v) CPC as blank. 14. Normalize the OD value with the protein concentration as determined in an identical culture.

3.5 Chondrogenic Differentiation

1. Pellet 8  105 of TDSCs by centrifuging the cells at 450  g for 10 min in a 15-ml conical polypropylene tube. 2. Add 2 ml of complete culture medium or chondrogenic medium to the cell pellet and incubate the cell pellet at 37  C, 5 % CO2 for 14–21 days. 3. Change half of the culture medium (i.e., 1 ml) every 3–4 days. 4. Remove the culture medium and fix the cell pellet with buffered formalin. 5. Dehydrate the cell pellet with increasing concentrations of ethanol using the tissue processing machine (Shandon Pathcentre, Thermo Fisher Scientific, MA, USA). 6. Embed the cell pellet in paraffin. 7. Section the pellet at the thickness of 5 μm. 8. Mount the section onto coated slide. 9. Deparaffinize the section twice with xylene for 5 min each. 10. Immerse the section in 100 % ethanol for 3 min.

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11. Rehydrate the section by immersion in 90 % and 70 % ethanol for 2 min each. 12. Stain the section with 0.1 % fast green for 5 min and then with 1 % acetic acid for 1 min. 13. Stain the section with 0.1 % Safranin O for 1 h for the examination of proteoglycan deposition. 14. Immerse the section in tap water for 1 min. 15. Dehydrate the section by immersion in 70 %, 80 % and 90 % ethanol for 10 s each and then in 100 % ethanol twice for 5 min each. 16. Immerse the section in xylene twice for 5 min each. 17. Mount the section with p-xylene-bis-pyridinium bromide (DPX) (Sigma-Aldrich, St. Louis, MO, USA). 18. Cover the section with glass cover slide. 19. View the slide under a light microscope (see Note 15). 3.6 Adipogenic Differentiation

1. Seed TDSCs at 4  103 cells/cm2 in complete culture medium in a 6-well culture plate at 37  C, 5 % CO2 until the cells reached confluence. 2. Replace with either 2 ml of complete culture medium or adipogenic medium and incubate at 37  C, 5 % CO2 for 21 days for the formation of oil droplets. 3. Assess the formation of oil droplets by Oil Red-O staining. 4. Wash the cells with PBS and fix the cells with 2 ml 70 % ethanol for 10 min. 5. Stain the cells with 2 ml freshly prepared 0.3 % Oil Red-O solution (Sigma-Aldrich, St. Louis, MO, USA) for 2 h at room temperature. 6. Remove the Oil Red-O solution and wash the cells twice with ddH2O. 7. View the cells under light microscope (see Note 16). 8. Remove the water. 9. Add 1 ml 100 % isopropanol and incubate for 10 min with gentle shaking to elute the Oil Red-O dye. 10. Pipette the solution several times to ensure that all the Oil Red-O dye is in the solution. 11. Transfer the solution to a 1.5-ml eppendorf tube. 12. Measure OD at 500 nm with 100 % isopropanol as blank. 13. Normalize the OD value with the protein concentration as determined in an identical culture.

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Notes 1. The species and the origin of tendon tissue affect the isolation efficiency. The age of the animals or individual also affect the success of getting TDSCs as there is depletion of TDSC pool in aged tendon tissues (23). We found that rat TDSCs were more easily isolated compared to human TDSCs, probably because of age and species differences. As a result, the isolation of TDSCs from young individuals is preferred. 2. It is important to have the target tissue for stem cell isolation free from contamination of the surrounding tissue. It is known that stem cells are present at the tendon mid-substance (24), peritendinous tissue (24), the tendon–bone junction (24), and the fat pad surrounding the tendon (25). Hence, for the isolation of stem cells from the tendon mid-substance, the peritendinous connective tissue, fat pad and the tendon–bone junction have to be completely removed. We scrape carefully the surface of the tendon with the top of the scalpel to strip away the surrounding tissues. 3. Some research groups added dispase in addition to collagenase type I for digestion (8, 10). 4. The number of colonies per nucleated cell varies with the initial plating density. More TDSCs and hence a purer culture is obtained at a relatively low initial plating density (9, 17, 18) (Fig. 1). The optimal initially seeding density varies with tendon tissue harvested from different species, origins and age of individual or animal. In my laboratory, the optimal seeding density for the isolation of stem cells from 8-week-old Sprague–Dawley

Fig. 1 Formation of cell colonies of tendon-derived nucleated cells isolated from the flexor tendon of 8-weekold Sprague–Dawley rats after culturing for 7 days at different initial seeding densities. The cell colonies were stained with crystal violet. Distinctive colonies meeting the criteria of colony counting were observed at the initial seeding density of 50 and 500 cells/cm2. An initial seeding density of 50 cells/cm2 achieved the highest number of single cell colonies per tendon-derived nucleated cell and was used as the optimal initial seeding for subsequent experiments. Magnification: 1 (Reprinted from Rui, Y.F., Lui, P.P.Y., Li, G., Fu, S.C., Lee, Y.W., and Chan, K.M. (2010) Isolation and characterization of multi-potent rat tendon-derived stem cells. Tissue Eng Part A 16(5), 1549–1558.)

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(SD) rat flexor tendon, 4–6-week-old SD rat patellar tendon, human patellar tendon, and human hamstring tendon were 50 cells/cm2 (9), 500 cells/cm2 (17), 500 cells/cm2 (18), and 500 cells/cm2 (unpublished observation, manuscript under revision), respectively. The procedure for the determination of the optimal seeding density is shown in the section titled “Determination of the optimal plating density for the isolation of TDSCs.” The optimal seeding density determined can be used as a guide for the isolation of TDSCs from same tendon tissue in the future. 5. The TDSCs attach to the plate and remain quiescent for several days before they divide rapidly to form colonies. 6. The morphology of TDSC is heterogeneous at P0 and the culture becomes more homogeneous with sub-culture. One research group removed tenocytes by picking TDSC colonies by local application of trypsin under microscopic visualization (10). The TDSC colonies were then pooled and subcultured for experiments. This method has the theoretical advantage of reducing the contamination by tenocytes but the efficiency of getting a “pure” TDSC culture is not clear and needs further experiments. 7. The renewal and multi-lineage differentiation potential of TDSCs decreased with passaging (17). Hence, the author recommended the use of only the early passage of TDSCs (P1-P5) for basic science, pathology and tissue engineering research. 8. TDSCs proliferated faster and have a shorter population doubling time at a lower seeding density (9, 18) (Figs. 2 and 3a, b). The exact optimal cell density for TDSC propagation can be determined by culturing TDSCs at different cell densities, cell counting, and calculation of the cumulative population doublings or population doubling time. The population doublings 2 Þ log ðN 1 Þ of TDSCs is calculated using the formula PD ¼ log ðNlogð2Þ where N1 is the initial cell number during cell seeding and N2 is the cell number at harvest. The cumulative population doublings (CPD) is calculated by adding the population doublings for each passage to the population doublings of the previous passages. The population doubling time is calculated using the formula PDT ¼ t  loglogð2Þ ðN1 =N2 Þ . where t is the time lapsed in hours (18). 9. We routinely culture TDSCs at 20 % O2 tension. However, our recent data showed that TDSCs expanded more efficiently and showed higher colony-forming ability at 2 % O2 tension compared to 20 % O2 tension (18) (Fig. 3). 10. As there are no specific markers for the isolation of TDSCs, TDSCs are isolated by an enrichment process that favors the

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Fig. 2 Graph showing the proliferative potential of rat flexor tendon TDSCs at passage 1 at different seeding densities. Cells seeded at the lowest density (100 cells/cm2) showed the highest fold increase of cell number than those seeded at higher densities. The results shown were mean  standard deviation of three Petri dishes for each time point. (Reprinted from Rui, Y.F., Lui, P.P.Y., Li, G., Fu, S.C., Lee, Y.W., and Chan, K.M. (2010) Isolation and characterization of multi-potent rat tendon-derived stem cells. Tissue Eng Part A 16(5), 1549–1558.)

proliferation of TDSCs compared to the other cell types. The TDSCs isolated are not pure stem cell culture. However, the selection of an optimal low density for TDSC isolation (Isolation and culture of TDSCs—step 11) would favor the isolation of purer culture. 11. Only colonies of size at least 2 mm in diameter with no colonyto-colony contact inhibition and stained clearly with crystal violet are counted. 12. According to the minimal criteria for defining human MSCs as suggested by the International Society for Cellular Therapy (26), 95 % of the MSC population must express CD105, CD73, and CD90. These cells must also lack expression (2% positive) of CD45, CD34, CD14 or CD11b, CD79α or CD19 and HLA class II. 13. TDSCs treated with fluorescent-labeled isotype-matched control should be included as negative controls to ensure the specificity of binding. 14. The ossified deposits appear red.

Fig. 3 Boxplots showing the fold change of the number of human patellar TDSCs incubated at 20 % or 2 % oxygen (O2) tension at (a) low plating density (500 cells/cm2) and at (b) high plating density (5,000 cells/cm2). (c) Boxplot comparing the doubling time of human TDSCs at the exponential phase at the low plating density at 20 % and 2 % O2 tension. The data shown was the results of six independent experiments (N ¼ 6/group). Double hash symbols indicated p  0.01 for comparing 2 % versus 20 % O2 tension groups at each time point. Open circle and asterisk in boxplots represented outliner and extreme value, respectively, of the dataset and they were also included in the data analysis. (Reprinted from Lee, W.Y., Lui, P.P., and Rui, Y.F. (2012) Hypoxia-mediated efficient expansion of human tendon-derived stem cells in vitro. Tissue Eng Part A 18(5–6), 484–498.)

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15. The proteoglycans appear orange-red. Chondrocytes appear as hypertrophic cells embedded in the lacunae. 16. The lipid droplets appear red.

Acknowledgement The author would like to thank Ms. Yin Mei Wong and Ms. Yuk Wa Lee for their help in proofreading the manuscript. References 1. Maffulli N, Wong J, Almekinders LC (2003) Types and epidemiology of tendiniopathy. Clin Sports Med 22(4):675–692 2. Hoffmann A, Gross G (2006) Tendon and ligament engineering: from cell biology to in vivo application. Regen Med 1(4):563–574 3. Miyashita H, Ochi M, Ikuta Y (1997) Histological and biomechanical observations of the rabbit patellar tendon after removal of its central one-third. Arch Orthop Trauma Surg 116(8):454–462 4. Lui PP (2013) Histopathological changes in tendinopathy – potential roles of BMPs? Rheumatology (Oxford). 52(12):2116–2126 5. Lui PP, Maffulli N, Rolf C, Smith RK (2011) What are the validated animal models for tendinopathy? Scand J Med Sci Sports 21(1):3–17 6. Lui PP, Ng SW (2013) Cell therapy for the treatment of tendinopathy – a systematic review on the pre-clinical and clinical evidence. Semin Arthritis Rheum 42(6):651–666 7. Billings PC, Fiori JL, Bentwood JL, O’Connell MP, Jiao X, Nussbaum B, Caron RJ, Shore EM, Kaplan FS (2008) Dysregulated BMP signaling and enhanced osteogenic differentiation of connective tissue progenitor cells from patients with fibrodysplasia ossificans progressive (FOP). J Bone Miner Res 23(3):305–313 8. Bi Y, Ehirchiou D, Kilts TM, Kilts TM, Inkson CA, Embree MC, Sonoyama W, Li L, Leet AI, Seo BM, Zhang L, Shi S, Young MF (2007) Identification of tendon stem/progenitor cells and the role of the extracellular matrix in their niche. Nat Med 13(10):1219–1227 9. Rui YF, Lui PPY, Li G, Fu SC, Lee YW, Chan KM (2010) Isolation and characterization of multi-potent rat tendon-derived stem cells. Tissue Eng Part A 16(5):1549–1558 10. Zhang J, Wang JHC (2010) Characterization of differential properties of rabbit tendon stem cells and tenocytes. BMC Musculoskelet Disord 11:10

11. Lovati AB, Corradetti B, Lange Consiglio A, Recordati C, Bonacina E, Bizzaro D, Cremonesi F (2011) Characterization and differentiation of equine tendon-derived progenitor cells. J Biol Regul Homeost Agents 25(2 Suppl):S75–84 12. Ruzzini L, Abbruzzese F, Rainer A, Longo UG, Trombetta M, Maffulli N, Denaro V (2013) Characterization of age-related changes of tendon stem cells from adult human tendons. Knee Surg Sports Traumatol Arthrosc. doi:10.1007/s00167-013-2457-4 13. Rui YF, Lui PP, Ni M, Chan LS, Lee YW, Chan KM (2011) Mechanical loading increased BMP-2 expression which promoted osteogenic differentiation of tendon-derived stem cells. J Orthop Res 29(3):390–396 14. Randelli P, Conforti E, Piccoli M, Ragone V, Creo P, Cirillo F, Masuzzo P, Tringali C, Cabitza P, Tettamanti G, Gagliano N, Anastasia L (2013) Isolation and characterization of 2 new human rotator cuff and long head of biceps tendon cells possessing stem cell-like self-renewal and multipotential differentiation capacity. Am J Sports Med 41(7):1653–1664 15. Lui PP, Wong OT (2012) Tendon stem cells: experimental and clinical perspectives in tendon and tendon-bone junction repair. Muscles Ligaments Tendons J 2(3):163–168 16. Lui PP, Chan KM (2011) Tendon-derived stem cells (TDSCs): from basic science to potential roles in tendon pathology and tissue engineering applications. Stem Cell Rev 7(4):883–897 17. Tan Q, Lui PP, Rui YF (2012) Effect of in vitro passaging on the stem cell-related properties of tendon-derived stem cells – Implications for tissue engineering. Stem Cells Dev 21(5):790–800 18. Lee WY, Lui PP, Rui YF (2012) Hypoxiamediated efficient expansion of human tendon-derived stem cells in vitro. Tissue Eng Part A 18(5–6):484–498

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19. Tsai RY, McKay RD (2000) Cell contact regulates fate choice by cortical stem cells. J Neurosci 20(10):3725–3735 20. Tsai MT, Lin DJ, Huang S, Lin HT, Chang WH (2012) Osteogenic differentiation is synergistically influenced by osteoinductive treatment and direct cell-cell contact between murine osteoblasts and mesenchymal stem cells. Int Orthop 36(1):199–255 21. Wang T, Xu Z, Jiang W, Ma A (2006) Cell-tocell contact induces mesenchymal stem cell to differentiate into cardiomyocyte and smooth muscle cell. Int J Cardiol 109(1):74–81 22. Canseco JA, Kojima K, Penvose AR, Ross JD, Obokata H, Gomoll AH, Vacanti CA (2012) Effect on ligament marker expression by directcontact co-culture of mesenchymal stem cells and anterior cruciate ligament cells. Tissue Eng Part A 18(23–24):2549–2558 23. Kohler J, Popov C, Klotz B, Alberton P, Prall WC, Haasters F, Muller-Deubert S, Ebert R, Klein-Hitpass L, Jakob F, Schieker M, Docheva

D (2013) Uncovering the cellular and molecular changes in tendon stem/progenitor cells attributed to tendon aging and degeneration. Aging Cell 12(6):988–999 24. Tan Q, Lui PP, Lee YW (2013) In vivo identity of tendon stem cells and the roles of stem cells in tendon healing. Stem Cells Dev 22(23):3128–3140 25. Buckley CT, Vinardell T, Thorpe SD, Haugh MG, Jones E, McGonagle D, Kelly DJ (2010) Functional properties of cartilaginous tissues engineered from infrapatellar fat pad-derived mesenchymal stem cells. J Biomech 43(5):920–926 26. Dominici M, Le Blanc K, Mueller I, SlaperCortenbach I, Marini FC, Krause DS, Deans RJ, Keating A, Prockop DJ, Horwitz EM (2006) Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy 8(4):315–317

Methods in Molecular Biology (2015) 1212: 141–161 DOI 10.1007/7651_2015_197 © Springer Science+Business Media New York 2015 Published online: 12 March 2015

A Human Colonic Crypt Culture System to Study Regulation of Stem Cell-Driven Tissue Renewal and Physiological Function Alyson Parris and Mark R. Williams Abstract The intestinal epithelium is one of the most rapidly renewing tissues in the human body and fulfils vital physiological roles such as barrier function and transport of nutrients and fluid. Investigation of gut epithelial physiology in health and disease has been hampered by the lack of ex vivo models of the native human intestinal epithelium. Recently, remarkable progress has been made in defining intestinal stem cells and in generating intestinal organoid cultures. In parallel, we have developed a 3D culture system of the native human colonic epithelium that recapitulates the topological hierarchy of stem cell-driven tissue renewal and permits the physiological study of native polarized epithelial cells. Here we describe methods to establish 3D cultures of intact human colonic crypts and conduct real-time imaging of intestinal tissue renewal, cellular signalling, and physiological function, in conjunction with manipulation of gene expression by lentiviral or adenoviral transduction. Visualization of mRNA- and protein-expression patterns in cultured human colonic crypts, and cross-validation with crypts derived from fixed mucosal biopsies, is also described. Alongside studies using intestinal organoids, the near-native human colonic crypt culture model will help to bridge the gap that exists between investigation of colon cancer cell lines and/or animal (tissue) studies, and progression to clinical trials. To this end, the near native human colonic crypt model provides a platform to aid the development of novel strategies for the prevention of inflammatory bowel disease and cancer. Keywords: Human, Intestine, Colon, Crypt, Stem cells, Culture, Signalling, Immunocytochemistry, In situ hybridization, Imaging, Proliferation, Migration, Tissue renewal

1

Introduction The human large intestinal epithelium is a monolayer of polarized epithelial cells that is exquisitely organized into millions of blindending flask-like invaginations called colonic crypts (Fig. 1). In the absence of villi (i.e., finger-like protrusions found in the small intestine), each colonic crypt is linked to its neighbor by adjoining surface epithelial cells that surround the crypt opening. Approximately ten billion cells are shed from the gut surface epithelium each day. These are continuously replaced by intestinal stem cell progeny that emerge from a population of proliferating LGR5/OLFM4positive stem cells located at the base of each colonic crypt (1).

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Shedding cell Mucus secreƟon

MigraƟon Enterocyte Enteroendocrine cell TuŌ cell

Goblet cell MitoƟc cell ProliferaƟon Progenitors & Stem Cell Niche Progenitor/stem cell

Fig. 1 Hierarchy of stem cell-driven renewal of the human colonic epithelium. The human colonic crypt base contains a mixture of proliferating stem cells, mitotic progenitor cells, and post-mitotic differentiated cells. Cells in the upper half of the crypt are mostly post-mitotic and migrate in an upward direction to the surface epithelium. On reaching this destination they are shed into the crypt lumen. These cells are replaced by stem cell progeny that emanate from the stem cell niche and differentiate en route to the surface epithelium

Stem cell progeny (i.e., transit amplifying cells) proliferate, migrate, and differentiate (into absorptive enterocytes, mucus-secreting goblet cells, hormone-secreting enteroendocrine cells and tuft cells that secrete prostaglandins and opioids) along the crypt-axis before they are shed from the surface epithelium. The prevailing dogma is that a hierarchy of rapid tissue renewal (approx. 5–7 days) minimizes the lifelong accumulation of molecular damage and disease risk (2). Long-lived stem cells are positioned in a relatively safe harbor (e.g., sterile) at the crypt-base, from where they fuel the

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constant replenishment of short-lived progeny that migrate up the crypt-axis and differentiate before being shed into the gut lumen within a matter of days. Each crypt houses all of the secretory and absorptive cell types that contribute to regular flushing of the crypt lumen (with fluid and mucus secretions) and maintenance of barrier function. As such, individual crypts not only represent the selfrenewing unit of the tissue but also fulfil the physiological functions of the intestinal epithelium. Isolation of intact colonic crypts has long-held the promise that functional self-renewing units of the human intestinal epithelium (i.e., crypts) could be studied ex vivo in real-time when placed in 3D culture conditions. Intact colonic crypts can be readily isolated following disaggregation of the lamina propria extracellular matrix by chelation of divalent cations or enzymatic treatment (3, 4). However, in this configuration, devoid of physical cell–matrix interactions, isolated crypts are prone to anoikis (5). Rescue of isolated crypt viability by embedding live crypts in a collagen matrix to restore cell–matrix interactions was an important first step towards reconstituting a viable self-renewing crypt culture system (3, 6). Although we and others demonstrated that embedding colonic crypts in a collagen gel maintained crypt morphology, cell polarity, and physiological function for a few days (3, 7), sustained cultured crypt viability required activation of mitogenic signals in combination with suppression of anti-proliferative signalling pathways. The Clevers’ Laboratory paved the way by capitalizing on their seminal work on Wnt signalling, intestinal stem cells and cancer (8–10), and developed the mouse small intestinal organoid culture system (11, 12). This “minigut” culture system formed the basis for the development of human organoid cultures from the intestinal epithelium (1, 13, 14) and from (inducible) pluripotent stem cells (15), both of which hold the potential for therapeutic transplantation (16, 17). For the study of native intact human colonic crypts, we adopted a similar approach whereby crypts are embedded in a basement membrane-derived matrix, Matrigel (18), instead of collagen I (7), and serum-free culture medium is supplemented with the intestinotrophic factors IGF-1 (activator of RAS signalling pathway) and R-spondin (activator of Wnt signals) in combination with inhibitors of BMP (Noggin or Gremlin proteins) and TGFβ (SB431542 or A83-01 small molecules) signalling pathways (18). The established human colonic crypt culture system provides a near-native model of a self-renewing tissue unit whereby epithelial cell polarity, tissue topology, and the hierarchy of stem cell-driven tissue renewal are maintained. Two major hall marks of intestinal crypts that are desirable to recapitulate in culture are: (1) the hierarchy of crypt cell renewal along the topological axis of the tissue and (2) functional epithelial cell polarity. Spatial correlation of protein/gene expression patterns with these characteristics can give valuable clues about gene function. Comparison of expression patterns in human

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cultured crypts should be made with crypts derived from immediately fixed colorectal tissue samples for cross-validation of the near-native culture system. Development of a validated nearnative colonic crypt culture system ex vivo provides the opportunity to gain insights into the regulation of (patho)physiological function. For example, assessment of signalling pathway status in immediately fixed clinical human colorectal tissue samples, together with assessment of the effects of pathway activation/ inhibition on the processes of crypt (stem) cell proliferation, migration, and shedding in cultured crypts, demonstrated that activation of canonical Wnt signals in combination with suppressed TGFβ/BMP pathways promotes renewal of the native human colonic epithelium (18, 19). Similarly, comparison of polarized expression patterns for NKCC1 in immediately fixed clinical human colorectal tissue samples with near-native cultured human colonic crypts proved invaluable when studying the functional regulation of fluid secretion by calcium-dependent internalization of NKCC1 in native polarized colonic crypt cells (7). These studies required adaptation of standard methods for manipulation of gene expression and assays for signalling pathway activation, cell proliferation, migration, and shedding so that they may be applied to the 3D human colonic crypt culture system. The aim of this chapter is to provide protocols not only for human colonic crypt isolation and culture but also for the application of a range of techniques that we have found invaluable to date. For example, a significant amount of time has been invested in optimizing the conditions for immunocytochemistry, mRNA in situ hybridization, and bright-field time-lapse and fluorescence real-time imaging. These techniques translate into methodologies to assess labelling of markers for intestinal stem cells (e.g., LGR5/ OLFM4), goblet cells (e.g., muc2), absorptive enterocytes (e.g., FABP1), tuft cells (e.g., COX1, COX2), enteroendocrine cells (e.g., chromogranin A, serotonin), cell signalling pathway status (e.g., beta catenin; phospho-smad 1,5,8; calcium waves), protein trafficking (e.g., NKCC1 internalization), crypt cell proliferation (e.g., BrDU, Ki67), crypt cell migration (time-lapse imaging), and crypt cell shedding (time-lapse imaging) (7, 18).

2

Materials

2.1 Human Colonic Crypt Isolation

1. Dissecting scissors, tissue forceps, and knives (25 mm blades, Roboz RS-6270). 2. 25 ml (Universal) screw top plastic bottles (RL Slaughter, UK). 3. Crypt isolation buffer: NaCl 140 mM, KCl 5 mM, Hepes (N-2-hydroxyethylpiperazine-N2-ethanesulfonic acid) 10 mM, D-glucose 5.5 mM, Na2HPO4 1, EDTA

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(diaminoethanetetraacetic acid disodium salt) 10 mM, dithiothreitol (1 mM), bovine serum albumin (0.1 % w/v) (All from Sigma, UK). 4. 25 ml Universal plastic bottle filled with 20 ml AdF12/DMEM culture medium (Invitrogen). 2.2 Adenoviral and Lentiviral Transduction [Optional]

1. Hexadimethrine bromide (#H9268, Sigma, UK; 8 mg/ml stock).

2.3 Human Colonic Crypt Culture

1. Matrigel 400 μl aliquot (Becton Dickinson, UK; #BD354230).

2. Adenovirus or lentivirus particles containing, for example, a gene reporter (e.g., Wnt reporter, HIV-based, VSV-G; SABiosciences) or dominant negative mutant (e.g., adenoviral CMV-DN-TCF4; type 5, dE1E3; Vector Biolabs).

2. AdF12/DMEM culture medium (Invitrogen, UK; #12634-028). 3. Penicillin/Streptomycin (10,000/10,000 U/ml; Invitrogen #15140-122). 4. Glutamine (100; Invitrogen; #25030-081). 5. B27 growth supplement (#12587-010, Invitrogen, UK). 6. N2 growth supplement (#17502-048, Invitrogen, UK). 7. N-acetylcysteine (0.5 M stock in PBS, Sigma; #A9165-5G). 8. Hepes (Invitrogen, #15630-056). 9. A83-01 (25 mM stock in DMSO; Tocris, UK; #2939). 10. Human recombinant R-spondin (250 μg/ml stock in 0.1 % BSA/PBS; Sino Biological, China or R&D Systems, UK). 11. Human recombinant Wnt 3A (100 μg/ml stock in 0.1 % BSA/ PBS; R&D Systems). 12. Human recombinant Noggin (200 μg/ml stock in 0.1 % BSA/ PBS; Peprotech, UK). 13. Human recombinant IGF-1 (100 μg/ml stock in 0.1 % BSA/ PBS; Sigma, UK). 14. Prostaglandin E2 (1 mM stock in DMSO; Caymen Chemicals). 15. Formulate 15 ml of human colonic crypt growth medium; 15 ml AdF12/DMEM, 10 μM hepes, 0.5 μM A83-01, 100 nM PGE2, 1 mM n-acetylcysteine, 200 ng/ml human recombinant Noggin, 100 ng/ml human recombinant Wnt 3A, 500 ng/ml human recombinant R-spondin-1, 50 ng/ml human recombinant IGF-1. 16. 4  12 well cell culture plates (Nunc multidish, or Corning) or glass bottomed multiwall dish (MatTek) for time-lapse or fluorescence imaging (Section 2.1, step 4). 17. Borosilicate glass, zero thickness, 16 mm diameter coverslip (VWR 631-0151), autoclaved.

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2.4 Bright-Field and Fluorescence TimeLapse Imaging

1. Live cell imaging system including climate control for temperature and CO2, automated stage, inverted microscope, range of objective lenses, bright-field DIC optics and fluorescence light source with range of excitation and emission filters. We use both Zeiss-Axiovision and Nikon-NIS elements based systems for time-lapse imaging and a Photon Technology Image Ratio Pro system based on a Nikon inverted microscope for calcium imaging 2. Glass bottom multiwell plate (e.g., MatTek) or POC chamber (PeCon, Germany) 3. Hepes-buffered saline (HBS): NaCl 140 mM, KCl 5 mM, Hepes (N-2-hydroxyethylpiperazine-N2-ethanesulfonic acid) 10 mM, D-glucose 5.5 mM, Na2HPO4 1 mM, CaCl2 1 mM, MgCl2 0.5 mM. 4. Fluorescence probes: e.g., Fura2 (calcium-sensitive dye, 1 mM stock DMSO; Invitrogen), calcein-AM (viable cell dye, 5 mM stock, Invitrogen), and propidium iodide (dead cell label, 1 mg/ml stock, Invitrogen).

2.5 Immunocytochemistry and Confocal Microscopy

For 20 coverslips, each with crypts cultured in a polymerized droplet of Matrigel, the following will be needed: 1. Phosphate buffered saline solution (PBS) (100 ml) prepared from dissolving 1 tablet (Oxoid BR0014G) per 100 ml deionized water in a reusable glass bottle. Autoclave before use. 2. Paraformaldehyde, 4 % (PFA) (10 ml) in PBS, made by adding 4 g of PFA powder (Sigma P6148) per 100 ml of previously autoclaved PBS or DEPC-treated PBS, gently stirred whilst warming on a hotplate (covered, in a fume hood) so that the temperature never exceeds 65  C until the solution is clear (1–4 h). It is cooled to 4  C, pH adjusted to 7.4 with 4 M NaOH, and aliquoted into 7 ml plastic screw top single use bottles (“Bijoux” E1412-0711 Starlab) or similar 25 ml bottles (“Universal,” 25 ml, R&D Slaughter code U128) and stored for up to 1 month in a 80  C freezer, thawed only immediately before use. 3. Triton X solution, 0.5 % (Roche) (10 ml) in PBS prepared immediately before use (Caution: light-sensitive). 4. Ammonium chloride, 100 mM NH4Cl2 (Sigma A9434) (10 ml) solution in PBS. 5. Optional antigen retrieval solution of sodium dodecyl sulfate (SDS, Melford B2008) (5 ml) which is only sparingly soluble in aqueous solutions. It is best prepared at least 24 h in advance by adding 1 % powder to PBS in a glass bottle and left on a rocker overnight until all has dissolved. It will store for 1 month at RT. Alternative antigen retrieval solutions such as 1 M HCl in PBS

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(5 ml) or citrate buffer (10 mM tri-sodium citrate (Sigma W302600) in water adjusted to pH 6 by addition of HCl) (100 ml) may be required as specified by antibody suppliers. 6. Blocking solution (5 ml) prepared up to 1 h before use by addition of 1 % Bovine serum albumin (BSA) (Sigma A2153) to PBS. Optional 10 % normal donkey serum (Sigma 9663) and/or 10 % normal goat serum (Abcam Ab7481) can be added, depending on the species of secondary antibody to be used. 7. Coverslip mounting media (200 μl) to prevent photobleaching (containing nuclear counterstain if required, e.g., propidium iodide or DAPI) such as VECTASHIELD™ (Vector labs H1000, H1200 or H1300), coated microscope slides (10) (Polysine™ VWR), and plain varnish (0.5 ml) (e.g., clear nail varnish) to seal around edges of coverslips. 8. Primary antibodies (1–10 μl) suitable for immunofluorescence, fluorophore conjugated secondary antibodies (0.5–5 μl). 2.6 mRNA In Situ Hybridization

1. For 20 coverslips the following are required 2. DEPC water or DEPC PBS are used throughout and are made by adding 0.1 % (1 μl per ml) diethylpyrocarbonate (DEPC, sigma) to water (or PBS solution) and left uncovered overnight at room temperature (in a fume hood). Next day, cover and autoclave solutions to remove the DEPC. 3. Hybridization buffer is prepared with 25 ml Formamide, 3.25 ml 20 SSC pH 5, 2.5 ml Chaps (10 % or 1 g/10 ml), 1 ml Tween 20 (10 %), 0.5 ml 0.5 M EDTA pH 8, 100 μl Heparin* (50 mg/ml), 125 μl tRNA& (from yeast, 20 mg/ ml), make up to 50 ml with DEPC water and store at 20  C, only remove from freezer immediately before required. Use within 1 month. *Heparin sodium salt from porcine intestinal mucosa SIGMA H3149; &Yeast type III RNA S. cervisae SIGMA R6750. 4. Salmon sperm DNA (SS DNA, Sigma D7656) is denatured by immersing a 1.7 ml flip top centrifugation tube with the lid covered with Parafilm containing a small volume (75 μl) SS DNA in a vacuum flask containing water heated to 95  C for 5 min. The tube is removed and allowed to cool (2 min) before hybridization buffer and an appropriate concentration of probe is added (see Note 11), and pre-warmed for at least 1 h at the incubation temperature to be used for hybridization reaction (Hybridization mix). 5. PBST (30 ml), 0.1 % Tween 20 in DEPC-treated PBS (0.5 ml 10 % Tween 20 per 500 ml of PBS) 6. PBST–MeOH (5 ml), a 50:50 mix of PBST and methanol

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7. HCl–PBST (5 ml), 1 M HCl in PBST 8. Make up a fresh stock of Proteinase K (BDH 390923P) in a flip top tube by adding 0.005 g PK to 0.5 ml DEPC-PBS. From this stock add 1 μl/ml to DEPC-PBS to make 5 ml Proteinase K working solution. 9. Hybridization humidity buffer (20 ml), formamide and 10 % 20 SSC (pH 5). 10. 20 SSC, 175.3 g NaCl + 88.2 g Sodium Citrate in 800 ml DEPC water pH adjusted (pH 7 with 10 N NaOH, or pH 5 1 M citric acid). Make up volume to 1 l, dispense into aliquots and sterilize by autoclaving. This stock concentrate can be used to make intermediate dilutions such as 5 ml 5 SSC (pH 5) or 5 ml 0.2 SSC (pH 5) by dilution with DEPC water. 2.7 Double Immunocytochemistry and In Situ Hybridization

1. Post-hybridization blocking solution (5 ml), 2 % BBR blocking reagent (Roche 11096176001), 20 % normal donkey serum (Sigma D9663) in Tris buffered saline (TBS). 2. TBS (10 ml), 50 mM Tris–HCl, pH 7.5 150 mM NaCl, dissolve 6.05 g Tris and 8.76 g NaCl in 800 mL of DEPC H2O. Adjust pH to 7.5 with 1 M HCl and make volume up to 1 L with DEPC H2O. 3. Normal human serum (40 μl). 4. Anti-fluorescein primary antibody (5 μl) (Abcam ab 19224), and 1–10 μl other primary and secondary antibodies suitable for immunofluorescence labelling. 5. Ten coated microscope slides (Polysine™, VWR), 200 μl mounting media (VECTASHIELD) and sealing varnish (1 ml).

2.8 Cross-Validation with Immediately Fixed Microdissected (MD) Crypts

Equipment as follows: Dissecting microscope with light source (Zeiss Discovery v1.2 microscope, KL2500 light source, flexible light guide with twin spot illuminators). Dissecting stage made from 10 cm disposable petri dish with dark card inside and edges sealed with tape. 2 microdissecting knives with 25 cm blades (Section 2.1, item 1). Biopsies or surgically obtained mucosal strips as described in crypt isolation are fixed immediately on removal from the patient in PFA (4 % in PBS) at 4  C for 1 h. The fixed tissue is removed from the PFA and left in PBS to wash overnight at 4  C. If tissue is to be used for in situ hybridization it is fixed and processed in solutions treated with DEPC and used within 24 h. For protein localization tissue can be stored in fridge for up to 7 days (see Note 18).

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For 20 coverslips

1. Small piece of fixed, washed, mucosal tissue 2. Microdissection solution (25 ml) 1 mM CaCl2 in PBS. 3. Hydrophobic barrier pen (e.g., Vector H4000) 4. Triton X 0.5 % in PBS (5 ml) 5. PBS (10 ml) 6. Matrigel™ thawed on ice (2  100 μl) 7. Two 12-well culture plates each with ten coverslips as described in crypt isolation materials, pre warmed in 37  C incubator. 8. Two plain glass microscope slides (75 mm  25  1.5) and two 7 ml bijoux bottles.

3

Methods

3.1 Human Colonic Crypt Isolation

1. Collect a 9 cm2 (i.e., 3 cm  3 cm) piece of mucosa from colorectal surgery and cut into 0.5 cm square pieces, or collect 5 biopsy samples from endoscopy (see Note 1). 2. Remove residual muscularis mucosa with dissecting scissors and forceps and discard. 3. Wash mucosal tissue samples with crypt isolation solution (Section 2.1, item 3) and place in a 20 ml volume contained in a Universal plastic bottle (Section 2.1, item 2). Incubate at room temperature for 90 min (see Note 2). 4. Meantime, fill 5 Universal plastic bottles (Section 2.1, item 2) with 10 ml of crypt isolation solution (Section 2.1, item 3). 5. At the end of the 90 min incubation period (3.1.4), place 5 tissue biopsy samples obtained at endoscopy, or 5  0.5 cm square pieces of colorectal mucosa derived from surgery (step 2), into a Universal bottle prefilled with 10 ml of crypt isolation solution (3.1.5). Shake the solution vigorously to liberate whole, intact, isolated crypts (see Notes 3 and 4). Liberated crypts will sediment under gravity to the bottom of the conical of the Universal plastic bottle (see Notes 5 and 6). 6. Carefully remove the remainder of the mucosal tissue samples from the first Universal plastic bottle (3.1.6) and transfer them into the second of five Universals that have been prefilled with crypt isolation solution (3.1.5). 7. Repeat the process of vigorous shaking, crypt liberation, and sedimentation (3.16 and 3.17) for the remaining four Universal bottles that have been prefilled with crypt isolation solution. 8. At this stage you will have 5 Universal plastic bottles each with a pellet of isolated crypts at the bottom of the conical.

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Collect each pellet of crypts with a 200 μl pipette and tip and place into a single Universal plastic bottle filled with 20 ml of ice-cold AdF12/DMEM (Section 2.1, item 4). Crypts will sediment under gravity and form a pellet (see Note 6). 9. Repeat steps 6–10 until crypts cease to be liberated from mucosal tissue samples. 10. Remove AdF12/DMEM supernatant from accumulated pellet of isolated crypts (3.1.9) and adjust volume of residual crypt suspension to a density of 500 crypts/100 μl AdF12/DMEM. 3.2 Adenoviral and Lentiviral Transduction [Optional]

1. Incubate isolated crypt suspension (Section 3.1, step 11) on ice for 15 min with hexadimethrine (8 μg/ml) and the desired adenovirus or lentivirus vector (e.g., Section 2.2) at a multiplicity of infection of 250  103 TU/crypt.

3.3 Human Colonic Crypt Culture

1. Prepare 15 ml of human colonic crypt growth medium in advance of the isolation (Section 2.3, item 15). 2. Thaw a 400 μl frozen aliquot of Matrigel on ice for 30 min (Section 2.3, item 1). 3. Add 1 ml of PBS to the corner wells of each 12-well plate (Section 2.3, item 12) and add one glass coverslip (Section 2.2, item 13) to the remaining 8 wells of each 12-well plate. Prewarm 12-well plates at 37  C for 30 min (see Note 8). 4. Take 100 μl of isolated crypt suspension (Section 2.1, item 11) and add to thawed 400 μl aliquot of Matrigel on ice and mix carefully. 5. Dispense 20 μl of isolated crypt/Matrigel suspension (Section 3.3, step 3) on to each coverslip and return 4  12 well plates to 37  C incubator for 15 min until Matrigel has polymerized. 6. Remove from incubator and flood polymerized Matrigel/crypt droplet with 400 μl of human colonic crypt culture medium; return to incubator. 7. Change culture medium every other day. A typical field of view and appearance of crypt morphology following 7 days in culture is shown in Fig. 2. 8. Matrigel will destabilize progressively in colonic crypt culture. Cultured crypts will need re-plating into a fresh Matrigel droplet every week. Crypts can be liberated by incubation in crypt isolation buffer (Section 2.1, item 3) for 20 min on ice followed by mechanical agitation with a 1 ml pipette and tip. Liberated crypts should be collected by sedimentation in AdF12/DMEM and resuspended into fresh Matrigel as above (see Note 9).

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Fig. 2 Human colonic crypt culture. (a) and (b) A typical field of view for human colonic crypts following 7 days in culture. Open arrowheads indicate crypt bases and filled arrowheads indicate accumulation of shed cells at the crypt opening. (c) Higher magnification view of crypts over 4 days and 7 days of culture (adapted from ref. (18)) 3.4 Real-Time Bright-Field and Fluorescence Imaging

1. Mount single glass coverslips into POC chamber or use crypts that have been seeded onto a multiwell plate with glass bottom (Section 2.4, item 2). 2. To monitor crypt cell morphology, proliferation, and migration in real-time, place multiwell plate into climate controlled chamber (5 % CO2, 37  C) located on stage of inverted microscope and establish DIC illumination with a 20 objective (Note 10). 3. Crypt cell proliferation, migration, and shedding along the crypt-axis are optimally observed by taking an image every 5 min for the duration of the experiment (18) (see Fig. 3 for an example of crypt cell proliferation). 4. To monitor crypt cell volume (7) or viability (18), load crypts with calcein-AM (5 μM) for 1 h, wash twice with HBS (Section 2.4, item 3). Excite at 488 nm and visualize emission at 520 nm. 5. To monitor crypt cell viability, incubate crypts with propidium iodide (1 μg/ml) which will fluoresce when it binds to the nuclei of cells that have lost the plasma membrane integrity. Excite propidium iodide at 488 nm and visualize emission at

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Fig. 3 Real-time imaging of crypt cell division. DIC imaging of cell nucleus migration from the basal pole to the apical pole (20 min) followed by cytokinesis (35 min) and return of daughter cell nuclei to the basal pole (75 min)

620 nm. Figure 4 illustrates the live/dead (i.e., calcein/propidium iodide) labelling pattern of a human colonic crypt (18). 6. To monitor intracellular calcium signals, load crypts with Fura2AM (5 μM) for 1 h. Wash crypts with HBS and excite alternately with 340 and 380 nm. Intracellular calcium signals are visualized by monitoring the ratio of fluorescence emitted at 510 nm following excitation by 340 and 380 nm. Figure 5 illustrates a calcium signal invoked in the stem cell niche at the crypt base by stimulation of muscarinic acetylcholine receptors (7). 3.5 Immunocytochemistry

1. Select isolated, cultured crypts (in Matrigel affixed to glass coverslips) and if required stimulate with agonists or inhibitors and/or incubate with BrDU (10 μM). Alternatively prepare MD crypts (Section 3.8). Leave coverslips in the culture dish. 2. Aspirate off any culture medium and add sufficient 4 % PFA in PBS to cover each Matrigel (200 μl).

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Fig. 4 Hierarchy of calcein-labelled live cells and propidium iodide (PI)-positive dead cells. The number of PIpositive shed cells increases over time in culture (adapted from ref. (18))

Fig. 5 Live cell imaging of calcium signalling along the colonic crypt-axis. Fura2 loaded crypts were stimulated with acetylcholine (10 μM). The F340/F380 pseudocolor ratio images demonstrate that acetylcholine induces a calcium signal in the stem cell niche which propagates along the crypt-axis with respect to time (adapted from ref. (7))

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3. Aspirate off fixative, pipette on sufficient PBS to cover Matrigel as a wash, remove by aspirating and repeat. 4. Permeabilize crypts by covering in Triton X solution (omit for MD). Leave for 30 min. 5. Remove Triton X and wash twice in PBS (as step 3 above). 6. Reduce cross-linking of protein by adding ammonium chloride (NH4Cl2) solution for 13 min. 7. Remove ammonium chloride and wash twice in PBS (as step 3 above). 8. An optional step can be included here to improve epitope retrieval by addition of SDS solution, 1 M HCl solution (BrdU antigen), or by boiling the coverslips in citrate buffer for 5–10 min, as appropriate to the primary antibody to be used. 9. Wash twice in PBS if antigen retrieval step used. 10. Discourage nonspecific antigen binding by blocking with 1 % BSA solution (with optional 10 % normal goat and/or donkey serum) for 2 h. 11. Prepare primary antibodies by diluting according to suppliers instructions (range 1:25 to 1:500 usually) in PBS. 12. Remove block by aspirating but do not wash and add 20 μl of antibody per coverslip. Wrap culture dish in film to reduce evaporation loss and leave on for 12–24 h. 13. Prepare (fluorophore-conjugated) secondary antibody according to suppliers’ instructions (usually at half the concentration that the primary was used at) in PBS. 14. Remove primary antibody by adding 200 μl per coverslip of PBS and then aspirating off. Wash second time in PBS, remove and add 20 μl per coverslip of secondary antibody mix. 15. Leave secondary antibodies on for 2 h, preventing dish from exposure to ambient or artificial light by covering with foil or light-proof plastic box to avoid bleaching of fluorophore. 16. Remove secondary antibodies as in step 13 above, leave in PBS wash for 1 h. 17. Lift out coverslips from dish, clean back surface with ethanol on a tissue add a drop of mounting media and invert the coverslip onto a coated microscope slide. Seal around edges of coverslip with a transparent varnish and store in a light proof box prior to imaging by confocal microscopy (see Fig. 6). 3.6 mRNA In Situ Hybridization

1. For 20 coverslips cultured or MD crypts. 2. Fix crypts on coverslips for 30 min in PFA (200 μL per well) leaving them in the 12-well culture dish (see Note 12).

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Fig. 6 Labelling human intestinal stem/progenitor cells. Stem cell markers LGR5 and OLFM4 were labelled in native human colonic crypts microdissected from a fixed biopsy specimen. Microdissected crypts were subjected to either immunocytochemistry for LGR5 (OriGene mouse monoclonal antibody clone 2A2) and OLFM4 (Abcam rabbit polyclonal antibody) or double LGR5 mRNA in situ hybridization and OLFM4 immunocytochemistry. Similar observations were made for cultured crypts (18)

3. Whilst fixing prepare and pre-warm hybridization mix in hybridization oven. 4. Aspirate off fixative and re-cover the crypts with PBST to wash. Remove and repeat wash. 5. Add PBST–MeOH so that each coverslip is covered for no more than 30 s, aspirate off, immediately replace with HCl–PBST (without wash), and leave on for 5 min. 6. Remove HCl–PBST and wash twice.

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7. Add Proteinase-K solution for 10 min and then remove and replace with PFA for 10 min. 8. Remove fixative and wash twice. Remove as much of PBST as possible with aspirator and then leave coverslips uncovered to dehydrate on bench for 1 h. 9. After dehydration add 75 μL per coverslip of warm hybridization mix, cover plate and wrap in tissue soaked in hybridization humidity buffer (see Note 13). 10. Put plates into hybridization oven at chosen temperature for 12–16 h. 11. Remove one plate from the oven, add 200 μl 5 SSC re-cover and replace in oven for 5 min, repeat with other plate (see Note 14). 12. Remove plate(s) from oven and aspirate off SSC and replace with 200 μL 0.2 SSC and leave on bench for 1 h (see Note 15). 13. At this stage coverslips can be removed from dish and mounted on a slide for imaging as described in Section 3.5, step 17; however, FAM labelled probes are prone to rapid photobleaching under a confocal microscope and it is advantageous to perform double immunocytochemistry and in situ additional steps to label the hybridized mRNA with a more photo-stable fluorescent conjugated secondary antibody and simultaneous labelling of other proteins of interest. 3.7 Double Immunocytochemistry and In Situ Hybridization

1. To coverslips that have reached stage 12 of in situ protocol (Section 3.6, step 12) remove 0.2 SSC from crypts and place in post-hybridization blocking solution for 90 min. During the blocking time prepare the anti-FAM primary antibody by preabsorbing it in normal human serum (10 μl serum per 1 μl antibody) at room temperature for 1 h and 37  C (water bath) for 30 min (see Note 16). 2. Allow pre-absorbed primary antibody mix to cool, add other primary antibodies (if required) and make up volume with TBS and 2 % normal donkey serum to desired final concentration (see Note 17). 3. Aspirate off block and add 20 μl of antibody/serum/TBS cocktail to each coverslip. Wrap (Section 3.5, step 12) and leave for 20–24 h at room temperature. 4. Add 200 μl TBS to each coverslip, aspirate off. Repeat wash. Add secondary antibody made up in TBS. Protect from light and incubate for 2 h at room temperature. Repeat TBS wash and leave in TBS for 1 h. Mount up coverslips (Section 3.5, step 17) and image with a confocal microscope (see Fig. 6).

Human Colonic Crypt Culture

3.8 Cross-Validation with Immediately Fixed Microdissected (MD) Crypts

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1. A hydrophobic barrier pen is used to draw a rectangle about 1 cm by 2 cm on a plain glass slide and filled with dissection solution with so that a flattened dome forms (200–400 μl). The coverslip is positioned on a stage under a dissecting microscope (see Note 19). 2. A 1 mm square section of the fixed tissue is cut under the microscope and is then reduced to individual, intact, epithelial crypts by teasing them apart from the underlying fibroblasts using a pair of microdissecting knives. 3. When 5–10 individual crypts have been teased off it is advantageous to remove them from the coverslip for safe-keeping using 1 ml pipette tip to aspirate them. Transfer to a 7 ml bijoux on ice. The dome of dissection solution is replenished and more individual crypts are teased from the biopsy. 4. Repeat until at least 200 crypts have been obtained. 5. Dissected crypts are permeabilized for 1 h in a 0.5 % solution of Triton X (Roche) in PBS at 4  C and then washed by replacing the solution with PBS twice. 6. About 100 of the individual crypts should be picked up using a 200 μl pipette and placed into a 100 μl aliquot of Matrigel that has been thawed on ice (see Note 20). 7. A 10 μl drop of Matrigel is placed on each coverslip (in a 12-well culture dish) and put for 30 min in 37  C incubator to set the Matrigel (see Note 21).

4

Notes 1. Obtain ethical approval and informed consent for collection of mucosal tissue samples from colorectal surgery or from recto/ sigmoid endoscopy for research purposes. 2. The crypt isolation solution is devoid of calcium and magnesium which will serve to disaggregate the extracellular matrix proteins that secure the crypts in the lamina propria. 3. The vigor with which mucosal tissue samples are shaken at the end of the incubation period for isolation needs to be optimized in an empirical fashion so as to liberate single intact crypts or small groups of intact crypts that are not fragmented. 4. Crypt liberation will depend on the efficiency with which the calcium-/magnesium-free crypt isolation solution causes (1) disaggregation of the extracellular proteins of the lamina propria, within which the crypts are embedded, and (2) DTTmediated lysis of extracellular mucus, which can form a barrier between the tissue samples and the crypt isolation solution.

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The incubation time in the crypt isolation solution may need to be extended if the crypts are slow to liberate. 5. 0.1 % BSA contained within the crypt isolation solution (Section 2.1, item 2) increases the specific gravity such that liberated crypts sediment to the bottom of Universal plastic bottles under gravity, whereas single contaminating mesenchymal cells such as myofibroblasts and lymphocytes do not. 6. Centrifugation is not recommended as it increases the likelihood of disrupting the integrity of liberated intact whole colonic crypts. 7. As pointed out previously (12), crypt viability is compromised by prolonged exposure to EDTA. 8. Filling of peripheral wells with PBS and in-filling between wells with PBS will help prevent evaporation of culture medium. 9. Crypt viability can be maintained for up to 1 month, but there is an increasing propensity for organoid formation (13). 10. Plastic dishes/lids interfere with DIC optics. 11. Design and synthesize oligonucleotide probes with at least two 5/6-carboxyfluorescein succinimidyl ester (5/6FAM) molecules at 50 and 30 ends for the target mRNA (This protocol was developed using Exiqon Custom LN™ detection probes with 20-mer sequence flanked by 5/6FAM molecules at 50 and 30 ends). Scrambled probes should be used as a negative control. A positive control probe for a ubiquitously expressed gene such as beta actin should also be used (18). 12. Calculate and ensure enough reagents are available for protocol, some reagents are best made shortly before using so whilst crypts are fixing, prepare reagents for next steps in protocol, e.g., Proteinase K, denatured Salmon sperm DNA (SS DNA Sigma D7656) and add 2 μl probe (probe concentration in range 10–75 nM) to 1,433 μl hybridization buffer and 75 μl SS DNA in 1.7 ml flip top centrifuge tube and place mix in hybridization oven to warm (approx 90 min) to make hybridization mix. PBS–MeOH is also best used freshly prepared. 13. Once the hybridization mix has been added it is vital now that the coverslips never dry out again, so carefully wrap plate in an absorbent cloth soaked in formamide/4 SSC solution and then over-wrapped with plastic film such as catering cling-film. Place in hybridization oven (at a temperature depending on the probe but in the range of 40–70  C) for at least 12 h (up to 16 h determined by experiment). 14. Remove one plate at a time from the oven, unwrap but do not discard) and add 200 μl of 5 SSC to each coverslip (there will not be sufficient probe remaining to aspirate), wrap up to prevent desiccation and return to the oven (bottom shelf) for 5 min.

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15. Remove from oven, remove and discard wrapping, aspirate off 5 SSC and replace with sufficient 0.2 SSC to cover crypts, and leave for 1 h at room temperature. If double immunocytochemistry and in situ hybridization are attempted, then the following steps in the protocol prepare the hybridized crypts for the addition of primary antibodies. 16. It is essential that no phosphate containing buffers are used from here as it may interfere with the hybridized probe; only use TBS. 17. The addition of 2 % donkey serum to block is to reduce nonspecific binding when donkey fluorescence conjugated secondary antibodies are used (or substitute relevant secondary antibody species). 18. If MD crypts are to be used for in situ hybridization then all solutions must be made with DEPC-treated water, and particular attention paid to cleaning instruments and surfaces to avoid DNase contamination 19. Twin spot light sources of the dissecting microscope should be adjusted so that a near horizontal beam of light falls on the specimen from either side. Correct positioning of the light source aids depth perception of the specimen when dissecting. 20. The minimum amount of fluid should be taken up with the crypts to avoid excessive dilution of the Matrigel. Gravity sedimentation of crypts is preferable to pelleting by centrifugation as the latter can cause clumping of the dissected and permeabilized crypts when resuspending in Matrigel. 21. When the Matrigel has set crypts can be covered with 200 μl per well of PBS and stored at 37  C for 1 day, or fixed in PFA for immediate processing for immunocytochemistry or in situ hybridization.

Acknowledgements We are grateful to all the past and present members of the Williams laboratory for their input into the development and application of the human colonic crypt culture system. The work in the Williams laboratory has been funded by grants from the Biotechnology and Biological Sciences (BB/F015690/1, BB/D018196/1), the Humane Research Trust, Big C cancer charity, John and Pamela Salter Charitable Trust.

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References 1. Sato T, Clevers H (2013) Growing selforganizing mini-guts from a single intestinal stem cell: mechanism and applications. Science 340(6137):1190–1194. doi:10.1126/science. 1234852 2. Vermeulen L, Morrissey E, van der Heijden M, Nicholson AM, Sottoriva A, Buczacki S, Kemp R, Tavare S, Winton DJ (2013) Defining stem cell dynamics in models of intestinal tumor initiation. Science 342(6161):995–998. doi:10.1126/science.1243148 3. Whitehead RH, Brown A, Bhathal PS (1987) A method for the isolation and culture of human colonic crypts in collagen gels. In Vitro Cell Dev Biol 23(6):436–442 4. Gibson PR, van de Pol E, Maxwell LE, Gabriel A, Doe WF (1989) Isolation of colonic crypts that maintain structural and metabolic viability in vitro. Gastroenterology 96(2 Pt 1):283–291 5. Grossmann J, Maxson JM, Whitacre CM, Orosz DE, Berger NA, Fiocchi C, Levine AD (1998) New isolation technique to study apoptosis in human intestinal epithelial cells. Am J Pathol 153(1):53–62. doi:10.1016/s00029440(10)65545-9 6. Strater J, Wedding U, Barth TF, Koretz K, Elsing C, Moller P (1996) Rapid onset of apoptosis in vitro follows disruption of beta 1-integrin/matrix interactions in human colonic crypt cells. Gastroenterology 110(6):1776–1784 7. Reynolds A, Parris A, Evans LA, Lindqvist S, Sharp P, Lewis M, Tighe R, Williams MR (2007) Dynamic and differential regulation of NKCC1 by calcium and cAMP in the native human colonic epithelium. J Physiol 582(Pt 2):507–524. doi:10.1113/jphysiol.2007. 129718 8. Batlle E, Henderson JT, Beghtel H, van den Born MM, Sancho E, Huls G, Meeldijk J, Robertson J, van de Wetering M, Pawson T, Clevers H (2002) Beta-catenin and TCF mediate cell positioning in the intestinal epithelium by controlling the expression of EphB/ephrinB. Cell 111 (2):251–263, S0092867402010152 [pii] 9. van de Wetering M, Sancho E, Verweij C, de Lau W, Oving I, Hurlstone A, van der Horn K, Batlle E, Coudreuse D, Haramis AP, TjonPon-Fong M, Moerer P, van den Born M, Soete G, Pals S, Eilers M, Medema R, Clevers H (2002) The beta-catenin/TCF-4 complex imposes a crypt progenitor phenotype on colorectal cancer cells. Cell 111(2):241–250, S0092867402010140 [pii] 10. Barker N, van Es JH, Kuipers J, Kujala P, van den Born M, Cozijnsen M, Haegebarth A,

Korving J, Begthel H, Peters PJ, Clevers H (2007) Identification of stem cells in small intestine and colon by marker gene Lgr5. Nature 449(7165):1003–1007. doi:10.1038/ nature06196, nature06196 [pii] 11. Sato T, Vries RG, Snippert HJ, van de Wetering M, Barker N, Stange DE, van Es JH, Abo A, Kujala P, Peters PJ, Clevers H (2009) Single Lgr5 stem cells build crypt-villus structures in vitro without a mesenchymal niche. Nature 459(7244):262–265. doi:10.1038/ nature07935, nature07935 [pii] 12. Sato T, Clevers H (2013) Primary mouse small intestinal epithelial cell cultures. Methods Mol Biol (Clifton, NJ) 945:319–328. doi:10. 1007/978-1-62703-125-7_19 13. Sato T, Stange DE, Ferrante M, Vries RG, Van Es JH, Van den Brink S, Van Houdt WJ, Pronk A, Van Gorp J, Siersema PD, Clevers H (2011) Long-term expansion of epithelial organoids from human colon, adenoma, adenocarcinoma, and Barrett’s epithelium. Gastroenterology 141(5):1762–1772. doi:10. S0016-5085 1053/j.gastro.2011.07.050, (11)01108-5 [pii] 14. Jung P, Sato T, Merlos-Suarez A, Barriga FM, Iglesias M, Rossell D, Auer H, Gallardo M, Blasco MA, Sancho E, Clevers H, Batlle E (2011) Isolation and in vitro expansion of human colonic stem cells. Nat Med 17 (10):1225–1227. doi:10.1038/nm.2470 15. Spence JR, Mayhew CN, Rankin SA, Kuhar MF, Vallance JE, Tolle K, Hoskins EE, Kalinichenko VV, Wells SI, Zorn AM, Shroyer NF, Wells JM (2011) Directed differentiation of human pluripotent stem cells into intestinal tissue in vitro. Nature 470(7332):105–109. doi:10.1038/nature09691 16. Watson CL, Mahe MM, Munera J, Howell JC, Sundaram N, Poling HM, Schweitzer JI, Vallance JE, Mayhew CN, Sun Y, Grabowski G, Finkbeiner SR, Spence JR, Shroyer NF, Wells JM, Helmrath MA (2014) An in vivo model of human small intestine using pluripotent stem cells. Nat Med 20(11):1310–1314. doi:10. 1038/nm.3737 17. Yui S, Nakamura T, Sato T, Nemoto Y, Mizutani T, Zheng X, Ichinose S, Nagaishi T, Okamoto R, Tsuchiya K, Clevers H, Watanabe M (2012) Functional engraftment of colon epithelium expanded in vitro from a single adult Lgr5(+) stem cell. Nat Med 18(4):618–623. doi:10.1038/nm.2695, nm.2695 [pii] 18. Reynolds A, Wharton N, Parris A, Mitchell E, Sobolewski A, Kam C, Bigwood L, El Hadi A,

Human Colonic Crypt Culture Munsterberg A, Lewis M, Speakman C, Stebbings W, Wharton R, Sargen K, Tighe R, Jamieson C, Hernon J, Kapur S, Oue N, Yasui W, Williams MR (2014) Canonical Wnt signals combined with suppressed TGFbeta/BMP pathways promote renewal of the native human colonic epithelium. Gut 63(4):610–621. doi:10.1136/gutjnl-2012304067

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Methods in Molecular Biology (2015) 1212: 163–169 DOI 10.1007/7651_2014_97 © Springer Science+Business Media New York 2014 Published online: 26 July 2014

Ca2+ Handling in Mouse Embryonic Stem Cell-Derived Cardiomyocytes Wenjie Wei and Jianbo Yue Abstract Embryonic stem (ES) cells are alternative cell source for cell replacement therapy for cardiac diseases, thus it is important to verify if the cardiomyocytes derived from ES cells have comparable functional parameters similar to the mature cardiomyocytes. Ca2+ handling is one of the most important parameters of cardiomyocyte since it is involved in the regulation of several main functions of cardiomyocytes, e.g. the excitation–contraction coupling. By applying membrane-permeable fluorescent Ca2+ indicator and confocal microscopy detection system, change of intracellular Ca2+ concentration in ES cell-derived cardiomyocytes can be monitored in real-time manner. In this protocol, we describe a method of isolating mouse ES cell-derived cardiomyocytes and recording their global and local Ca2+ transients. Keywords: Mouse embryonic stem cells, Ca2+, Cardiomyocytes, Fluo-4 AM, Confocal microscopy

1

Introduction Ever since been discovered in 1980s, embryonic stem (ES) cells have been considered as one of the most promising cell sources for cell replacement therapy because of their pluripotency (1, 2). Cardiovascular diseases remain as one of the major threatens to human health worldwide today, and years of research on cardiogenesis from ES cells have indeed provided helpful insights to develop effective clinical cell therapy for human cardiovascular disease. Typical method to induce cardiomyocytes differentiation from ES cells is to form 3D cell aggregates called embryoid bodies (EB) (3). Since the first successful application of this approach to generate cardiomyocytes (4), this method has been continuously optimized (5). Meanwhile, detailed functional assays are performed to verify if these ES cell-derived cardiomyocytes are functional equivalent to nature adult cardiomyocytes. Ca2+, as an universal secondary messenger, regulates numerous cellular processes. For example, increased intercellular Ca2+ concentration serves as the trigger of various cellular activities, such as muscle contraction, neurotransmitter release, and fertilization (6). Intracellular Ca2+ release is also essential for excitation–contraction coupling of cardiomyoctyes (7).

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Thus, detection of intracellular Ca2+ change in the ES cell-derived cardiomyocytes would provide helpful information for the functional assessment. The most widely used intracellular Ca2+ indicators are chemical fluorescence probes, which exhibit an increase in fluorescence upon binding to Ca2+. These indicators can be divided into two groups: ratiometric such as Fura 2 (excitation ratio 340/380 nm) and Indo1 (excitation ratio 405/485 nm), and nonratiomentric such as Fluo 4 (excitation 488 nm) and rhod 2 (excitation 570 nm) (8). Ratiometric indicators can minimize the effects of photobleaching, leakage, and uneven loading, thereby delivering more robust and reproducible results, while nonratiometric indicators are highly sensitive for rapid Ca2+ changes (9, 10). Most fluorescence indicators are membrane impermeable; it is their acetoxymethyl ester (AM) form that is commonly applied. The AM group can help the indicator cross the cell membrane and are subsequently removed by cellular esterases to regenerate the indicator. In this protocol, using mouse ES cells as the model, we describe the procedure of deriving and isolating ES cell-derived cardiomyocytes, and how to measure global and local Ca2+ transitions in these cardiomyocytes by using Fluo-4 AM and laser scanning confocal microscopy (11).

2

Materials

2.1 ES Cells Cardiac Differentiation

1. ES cell culture medium: Millipore ESGRO Complete™ Clonal Grade Medium (Cat No. SF001-500). 2. 10x PBS stock: Dissolve 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 18 mM KH2PO4 in 800 mL of H2O, adjust pH to 7.1 with HCl, finally add dH2O to 1 L. Sterilize the solution by autoclaving for 20 min at 121  C. Store at room temperature. 3. 1x PBS solution: Add 900 mL H2O into 100 mL 10x PBS stock solution, filter with 0.22 μM filter, and autoclave for 20 min at 121  C to sterilize. Store at room temperature. 4. Accutase: Sigma (Cat No. A6964). 5. Cardiac differentiation medium: DMEM (Lifetech, Cat No.11965026), 20 % FBS (ES qualify, Lifetech, Cat No.10439024), 1 % L-glutamine (Lifetech, Cat No. 25030081), 1 % nonessential amino acid (Lifetech, Cat No. 11140050), 1 % penicillin and streptomycin (Lifetech, Cat No.15140122), 0.2 % 2-mercaptoethanol (Lifetech, Cat No. 21985023). 6. 0.1 % gelatin solution: 1 g gelatin (Sigma, Cat No.G9391) powder in 1 L dH2O, 121  C autoclave for 1 h to dissolve the powder, then filter with 0.45 μM filter. 7. 15 mL tubes, Petri dish. 8. Centrifuge.

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2.2 Cardiomyocytes Dissection and Culture

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1. Microscalpels (PARAGON, Cat No.900012). 2. 6-well plastic tissue culture plates, 18 mm cover glass. 3. Enzyme buffer: Dissolve 120 mmol/L NaCl, 5.4 mmol/L KCl, 5 mmol/L MgSO4, 5 mmol/L Na pyruvate, 20 mmol/L glucose, 20 mmol/L taurine, 10 mmol/L HEPES in MQ water, adjust pH to 6.9. Filter with 0.22 μM sterile filter before use. 4. Enzyme solution: Dissolve 1 mg/mL collagenase II (Lifetech Cat. No17101015), plus 30 μmol/L CaCl2 in enzyme buffer, filtered with 0.22 μM sterile filter, store at 4  C (Note 1). 5. KB buffer: Dissolve 85 mmol/L KCl, 30 mmol/L K2HPO4, 5 mmol/L MgSO4, 1 mmol/L EGTA, 2 mmol/L Na2ATP, 5 mmol/L Na pyruvate, 5 mmol/L creatine, 20 mmol/L taurine, 20 mmol/L glucose in MQ H2O, adjust pH to 7.2, filter with 0.22 μM sterile filter.

2.3 Ca2+ Measurement

1. 1 HBSS: (with Ca2+: Lifetech, Cat. No. 14025092). 2. Ca2+ dye: Fluo-4 AM (Lifetech, Cat. No.F14201), dissolve in DMSO to make a final concentration of 2 mM. 3. Chambers for cover glass (We made by our own, but they are also available from Corning and Nunc). 4. 10 % Pluronic F-127 solution: Sigma (Cat. No. P2443), add 1 g powder in 10 mL dH2O, put into 50  C water bath until the powder dissolve totally. Store as 0.5 mL aliquot in room temperature (Note 2). 5. Confocal imaging system (Olympus Fluoview System version 4.2 FV300 TIEMPO) connected with an inverted microscope (Olympus IX71).

3

Methods

3.1 ES Cells Cardiac Differentiation

1. Gelatin coating of tissue plates and cover glass: For 6 cm plate, add 3 mL of 0.1 % gelatin solution; for 6-well plate, add 1.5 mL of into each well; place the plates in 37  C incubator for 4 h, aspirate the gelatin solution before use. 18 mm coverglass can be placed in 6-well plate to go through the coating process. 2. Mouse ES cell culture: Plate cells at 3  105 cells/well in 6-well plate in 1.5 mL ESGRO complete medium, cells should be ready for passage in every 2 days. 3. Dissociate mouse ES cells: Grow mouse ES cells to 70–80 % confluence, wash with 1xPBS once, add 0.5 mL accutase/well, incubate at room temperature for 2–5 min to let cells detach. 4. Tap the bottom of the plate gently to remove all the colonies. Pipette up and down to get single cell suspension. Add the suspension into 5 mL basal medium (SF002-500) in a

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15 mL tube, mix the solution thoroughly, then spin at 233  g for 5 min. 5. Remove the supernatant; add 2 mL differentiation medium and pipette up and down to resuspend the cells. 6. Count cells and make a cell solution of 1  104 cells/mL in differentiation medium. Transfer 10 mL cell solution into a plastic reservoir. Open the petri dish lid inside up, and use a multichannel pipette to add around 40 drops (20 μl per drop) on it. Cover the lid back to the bottom which contains 10 mL 1 PBS, and put the dishes into 37  C incubator to let the hanging cells to aggregate by gravity. 7. After 2 days, the cells within the hanging drops form the embryoid body (EB). Collect those EBs with 1 mL pipet and put them into a noncoated 10 cm culture dish contains 10 mL differentiation medium, let the EBs grow for another 2 days in suspension. 8. After 2 days, use 1 mL pipet to collect 20 EBs and plate onto a gelatin-coated 6 cm tissue culture dish containing 3 mL differentiation medium. Put the dish into 37  C incubator for differentiation. Change medium every day after day 3. Spontaneously beating foci normally show up at around day 6 or 7. 3.2 Dissection and Culture of Cardiomyocytes

1. Mark the beating areas of embryoid bodies under a microscopy, isolate the beating areas by a microscalpel, and wash them once in 1 PBS at room temperature. Centrifuge them at 1,000 rpm/min for 5 min and discard supernatant. 2. Incubate the collected fragments in enzyme solution for 30 min at 37  C and then centrifuge at 1,000 rpm/min for 3 min. 3. Aspirate supernatant and resuspend the digested tissue fragments in KB solution. The dissociation of the tissue is completed in KB medium by gentle shaking at room temperature for 30 min. 4. 4. Pipette up and down 5–10 times, then seed the isolated cells on sterile gelatin-coated 18 mm coverglass in 6-well plate and put into 37  C incubator. After 30 min, add 1.5 mL differentiation medium. Change into fresh differentiation medium the next morning. The isolated cardiomyocytes are attached to the coverglass surface and begin spontaneous and rhythmical contractions after around 12 h of incubation.

3.3 Ca2+ Measurement Using Confocal Microscopy

1. Put the 18-mm coverglass seeded with cardiomyocytes into the chamber, and wash with 1 HBSS once. 2. Add 200 μL 1 HBSS contains 4 μM Fluo-4 AM and 0.02 % F127 into the chamber, and incubate in dark at room temperature for 30 min.

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Fig. 1 Representative spontaneous Ca2+ transients in beating mouse ES-derived cardiomyocytes. Fluo-4 AM is used as the Ca2+ indicator, F/F0 indicates normalized fluorescence, F is the intensity of fluorescence signal, and F0 is the basal fluorescence

3. Aspirate the dye solution, and wash the cells with 1 HBSS once. Add 200 μL 1 HBSS into the chamber and wait for 10 min before Ca2+ measurement. 4. Turn on the microscopy and the connected computer, mount the chamber on the stage of the microscopy, and use UPlanAPO 20 lens. 5. Turn on the power supply for 488 nm excitation laser and the controller, open the “fluoview” software, and select “fluo 4” in the dye panel. 6. Set the parameters: mode “XT,” resolution “512  512”, pan “X1”. 7. Press “scan,” adjust the focus plane to get the best image, and press “stop scan” to obtain one picture for further recording. 8. Use the ROI select tool in the panel to select 16 cells, change the mode into “XYT,” interval time “3 s,” recording times “300,” select “Timepo” subpanel, and click “show live plot” to show real-time graph. Left click once on the blank graph to activate real-time graph, click “XYT” to start time lapse. When series have been done, export the experiment for further data analysis (Fig. 1). 9. To measure local Ca2+ transients within the spontaneous beating cardiomyocytes, change the mode into “XT,” select “line” scan mode to scan a single line across the cell repeatedly, set interval time to“3 ms,” and click “stop scan and series done” button when enough scans have been recorded. Export and save the scan data (Fig. 2).

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Fig. 2 Representative confocal line scan image of Ca2+ transients in beating mouse ES cell-derived cardiomyocytes. Fluo-4 AM is used as the Ca2+ indicator

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Notes 1. To digest older EB beating clusters completely, 0.1–1 mg/mL pancreatin can be added into the enzyme solution. 2. 10 % Pluronic F-127 may precipitate when stored at room temperature, short time heating in water bath can dissolve it again before use.

Acknowledgement We thank Professor Yang HT and Professor Michal Opas for kindly sharing their protocol for isolating ES cell-derived cardiomyocytes with us. This work was supported by Research Grant Council (RGC) grants (782709M, 785911M, 769912M, 785213M, and 17126614M) to JY. References 1. Martin GR (1981) Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells. Proc Natl Acad Sci U S A 78:7634–7368 2. James A, Thomson J, I-E SS, Shapiro MA, Waknitz JJ, Swiergiel VS, Marshall JMJ (1998) Embryonic stem cell lines derived from human blastocysts. Science 282:1145–1147 3. Doetschman TC, Eistetter H, Katz M, Schmidt W, Kemler R (1985) The in vitro development of blastocyst-derived embryonic stem cell lines: formation of visceral yolk sac, blood islands and myocardium. J Embryol Exp Morphol 87:27–45

4. Klug MG, Soonpaa MH, Koh GY, Field LJ (1996) Genetically selected cardiomyocytes from differentiating embronic stem cells form stable intracardiac grafts. J Clin Invest 98:216–24 5. Kurosawa H (2007) Methods for inducing embryoid body formation: in vitro differentiation system of embryonic stem cells. J Biosci Bioeng 103:389–98 6. Clapham DE (1995) Calcium signaling. Cell 80:259–268 7. Cannell MB, Cheng H, Lederer WJ (1994) Spatial non-uniformities in [Ca2+]i during

Ca2+ Handling in Mouse Embryonic Stem Cell-Derived Cardiomyocytes excitation-contraction coupling in cardiac myocytes. Biophys J 67:1942–1956 8. Takahashi A, Camacho P, Lechleiter JD, Herman B (1999) Measurement of intracellular calcium. Physiol Rev 79:1089–1125 9. Minta A, Kao JP, Tsien RY (1989) Fluorescent indicators for cytosolic calcium based on rhodamine and fluorescein chromophores. J Biol Chem 264:8171–8178

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10. Grynkiewicz G, Poenie M, Tsien RY (1985) A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260:3440–3450 11. Wei W, Sun HY, Ting K, Zhang LH, Lee HC, Li GR, Yue J (2012) Inhibition of cardiomyocyte differentiation of mouse embryonic stem cells by CD38/cADPR/Ca2+ signaling pathway. J Biol Chem 287:35599–35611

Methods in Molecular Biology (2015) 1212: 171–181 DOI 10.1007/7651_2014_98 © Springer Science+Business Media New York 2014 Published online: 02 August 2014

Potential Application of Extracellular Vesicles of Human Adipose Tissue-Derived Mesenchymal Stem Cells in Alzheimer’s Disease Therapeutics Takeshi Katsuda, Katsuyuki Oki, and Takahiro Ochiya Abstract In the last 20 years, extracellular vesicles (EVs) have attracted attention as a versatile cell–cell communication mediator. The biological significance of EVs remains to be fully elucidated, but many reports have suggested that the functions of EVs mirror, at least in part, those of the cells from which they originate. Mesenchymal stem cells (MSCs) are a type of adult stem cell that can be isolated from connective tissue including bone marrow and adipose tissue and have emerged as an attractive candidate for cell therapy applications. Accordingly, an increasing number of reports have shown that EVs derived from MSCs have therapeutic potential in multiple diseases. We recently reported a novel therapeutic potential of EVs secreted from human adipose tissue-derived MSCs (hADSCs) (also known as adipose tissue-derived stem cells; ASCs) against Alzheimer’s disease (AD). We found that hADSCs secrete exosomes carrying enzymatically active neprilysin, the most important β-amyloid peptide (Aβ)-degrading enzyme in the brain. In this chapter, we describe a method by which to evaluate the therapeutic potential of hADSC-derived EVs against AD from the point of view of their Aβ-degrading capacity. Keywords: Adipose tissue-derived stem cell, Alzheimer’s disease, β-amyloid, Extracellular vesicle, Exosome, Mesenchymal stem cell, Microvesicle, Neprilysin

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Introduction Extracellular vesicles (EVs) have recently attracted significant attention as a novel cell–cell communication vehicle. EVs are cellsecreted lipid bilayered vesicles that contain microRNAs, mRNAs, and proteins, which they transfer between cells. In the last two decades, it has been shown that EVs regulate a variety of biological phenomena. The biological functions of EVs as well as their contents vary according to their cellular origins. Notably, however, the functions of EVs mirror, at least in part, those of the cells from which they originate. For example, EVs derived from B-lymphocytes and dendritic cells have an antigen-presentation capacity (1, 2). However, cancer cell-derived EVs exhibit functions associated with malignancy, including the ability to promote proliferation, drug resistance, invasiveness, and metastasis (3). This evidence indicates that cells utilize their EVs to fulfill their

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roles as cells. This suggests that the EVs released from cells that are capable of repairing damaged tissue could have therapeutic effects in a clinical setting. An increasing number of reports have shown that EVs derived from mesenchymal stem cells (MSCs) have therapeutic potential in several diseases (4–7). MSCs are a type of adult stem cell that can be isolated from connective tissue such as bone marrow and adipose tissue; these cells have emerged as an attractive candidate for cell therapy applications (7–10). Although MSCs initially attracted interest for their multipotency, the beneficial effects of MSCs currently appear to be due to other properties. In particular, their paracrine ability, including the secretion of immunomodulatory cytokines and tissue repair-inducing growth factors, are of the most interest among researchers. In addition to such secretory ability, recent studies have revealed that EVs also contribute to the therapeutic effects of MSCs. Evidence of the therapeutic effects of MSC-derived EVs has been reported in studies on kidney injury (11–15), myocardial injury (16–19), middle cerebral artery occlusion (20–22), and lung injury (23, 24). Recently, we reported a novel therapeutic possibility for using MSCs-derived EVs against Alzheimer’s disease (AD) (25). One of the neuropathological hallmarks of AD is the accumulation of βamyloid peptide (Aβ) in the brain because of an imbalance between Aβ production and clearance (26). Among the several proteases involved in Aβ proteolysis, neprilysin (neutral endopeptidase; NEP), a type II membrane-associated metalloendopeptidase, is the most important (26). We found that human adipose tissuederived MSCs (hADSCs) produce EVs that contain enzymatically active NEP. A coculture of neuronal cell line N2a cells with hADSCs led to decreases in the secreted Aβ40 and Aβ42 levels as well as a decrease in the intracellular Aβ42 level. In this chapter, we describe a method to evaluate the therapeutic potential of hADSCderived EVs from the point of view of NEP activity.

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Materials

2.1 Isolation of hADSC-Derived EVs

Reduced serum medium (MesenPRO RS® Medium) (Invitrogen). Antibiotic-antimycotic (Invitrogen). 100 GlutaMAX® (Invitrogen). CellBIND® Surface 100 mm dish (Corning). Dulbecco’s phosphate-buffered saline without calcium and magnesium (PBS( )) Serum-free medium (StemPRO MSC SFM®) (Invitrogen). 0.22-μm filter membrane. Ultra-Clear centrifugation tubes (Beckman Coulter, Inc.). SW41Ti rotor (Beckman Coulter, Inc.).

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2.2 The Routine Culturing of Mouse Neuroblastoma Cell Line Neuro-2A (N2A) Cells

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Fetal bovine serum (FBS) (Invitrogen). Dulbecco’s modified Eagle’s medium (DMEM) (high glucose) (Invitrogen). Antibiotic-antimycotic (Invitrogen). 100-mm cell culture dish (Nunc). 0.5 % Trypsin-EDTA (Invitrogen). PBS( ).

2.3 NEP Enzyme Activity Assay of EVs

Fluorescence microplate reader. Black 96-well plate (Nunc). NEP assay buffer (100 mM Tris, 50 mM NaCl, 10 μM ZnCl2, pH ¼ 7.5). Fluorogenic peptide substrate Mca-RPPGFSAFK(Dnp)-OH (2 mM in dimethyl sulfoxide) (R&D Systems, Inc.). Thiorphan solution (5 mg/mL in ethanol) (Enzo Life Sciences). Recombinant human NEP (rhNEP) (R&D Systems, Inc.). Transparent plate adhesive film (MicroAmp® Optical Adhesive Film) (Applied Biosystems).

2.4 Transfer Assay of hADSC-EVs to N2a Cells

PKH fluorescent cell linker kit (Sigma). Centrifugal ultrafiltration unit 100 kDa (Amicon Ultra-0.5 100 kDa®, Millipore). PBS( ).

2.5 Coculture of N2a Cells with hADSCs 2.6 Determination of Aβ40 and 42 Concentration via ELISA

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Cell culture inserts possessing 0.4-μm pores (BD Falcon). Multiwell plates for cell culture inserts (BD Falcon). Spectrophotometer. Human beta-amyloid (1-40) ELISA Kit wako II (Wako; 298-64601). Human beta-amyloid (1-42) ELISA Kit wako (Wako; 298-62401). Cell lysis buffer (Mammalian Protein Extraction Reagent (M-PER) ®, Thermo).

Methods

3.1 Isolation of hADSC-Derived EVs

The method described here is summarized as a flowchart in Fig. 1. 1. Seed hADSCs to CellBIND plates and culture them in reducedserum medium MesenPRO® until they reach 90–100 % confluence (see Notes 1 and 2). 2. Remove the culture medium and add fresh serum-free medium StemPRO MSC SFM® (see Note 3).

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Fig. 1 Flow chart of the protocol for preparing hADSC-EVs. Culture hADSCs in reduced serum medium (RSM) until they reach 80–90 % confluency. After a medium change with serum-free medium (SFM), culture the cells for 2 days and harvest the conditioned medium (CM) (first harvest). Supplement the cells with fresh SFM and continue the culture for another 2 days, then harvest the CM (second harvest). Combine the first and second CM into one batch, and filtrate it with a 0.22-μm filter unit. Ultracentrifuge the CM at 110,000  g for 70 min. Then, wash the pellet with PBS( ), and after ultracentrifugation resuspend the pellet in the desired volume of PBS( ). The presence of vesicles 100–200 nm in diameter can then be confirmed. A phase-contrast electron microscopy image of hADSC-EVs is shown (scale bar: 100 nm)

3. Culture the cells for 2 days. 4. Collect the conditioned medium (CM) (first harvest), and supplement the cells with fresh StemPRO SFM. 5. Centrifuge the collected CM at 2,000  g for 10 min at 4  C to remove the cells and cellular debris. 6. After the centrifugation, carefully remove the supernatant to a fresh tube, leaving a few mL behind. 7. Remove the cellular debris via filtration through a 0.22-μm filter membrane. Store the filtrated CM at 4  C. 8. Continue culturing the cells for another 2 days in StemPRO SFM, and collect the CM as described above (second harvest). Combine the two harvested batches of CM into one batch. 9. Transfer 11 mL of the obtained CM to an Ultra-Clear tube. 10. Centrifuge at 110,000  g for 70 min at 4  C in Ultra-Clear centrifuge tubes in a SW41Ti rotor. 11. Carefully remove the supernatant.

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12. Add 2 mL of PBS( ) to each tube and vortex vigorously for a few seconds. 13. Add 9.5 mL of PBS( ) to each tube. 14. Centrifuge at 110,000  g for 70 min at 4  C in Ultra-Clear centrifuge tubes in a SW41Ti rotor. 15. Quickly decant the supernatant. 16. Resuspend the pellet (EV fraction) in 50–100 μL of PBS( ) (see Note 4). 17. Use immediately or store at 4  C. 18. Determine the protein concentration of EVs using generally available methods such as the Bradford assay or BCA assay (see Note 5). 19. (Optional) Confirm the presence the EV marker proteins, such as several tetraspanins (CD9, CD63, and CD81), using western blotting. 20. (Optional) Determine the number of exosomes using the Nanosight system (NanoSight) or qNano system (Izon Science, Ltd.). 3.2 NEP Enzyme Activity Assay of EVs

Our laboratory measured the NEP-specific enzyme activity using a fluorogenic peptide substrate, Mca-RPPGFSAFK(Dnp), and a selective NEP inhibitor, thiorphan. This substrate can be cleaved by several endopeptidases, including NEP, endothelin-converting enzyme (ECE)-1, ECE-2, angiotensin-converting enzyme (ACE)-1, ACE2, and insulin-degrading enzyme (IDE) (27). However, at pH 7.5, the use of thiorphan allows the discrimination of NEP enzyme activity from other closely related enzymes (27) (see Note 6). 1. Prepare 2 EV solution by diluting the EV stock solution in NEP assay buffer. We usually prepare 100–200 μg/mL as a 2 EV solution so that 5–10 μg of EVs are used for the reaction in each well. 2. Prepare 2 rhNEP dilution series using NEP assay buffer as a positive control (see Note 7). 3. Apply 50 μL of 2 EV solution or 2 rhNEP dilution series to each well of a black 96-well plate on ice. 4. Prepare 2 substrate solution by diluting the substrate stock solution with assay buffer to 10–20 μM at the final concentration in the presence or absence of thiorphan (2.5 μg/mL in the final solution). 5. Add 50 μL/well of thiorphan (+) or ( ) substrate solution to each well that contains rhNEP or EVs on ice (final reaction volume is 100 μL/well) (see Note 8). 6. Seal the plate with a transparent adhesive film and keep on ice until measurement.

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Fig. 2 NEP enzyme activity assay. NEP enzyme activity can be measured using a fluorogenic peptide substrate, Mca-RPPGFSAFK(Dnp), and the NEP inhibitor thiorphan. NEP-specific activity (a) is calculated by subtracting the residual fluorescence intensity measured in the presence of thiorphan (b) from the total enzyme activity measured in the absence of thiorphan (c). The addition of thiorphan sharply reduces the enzyme activity of hADSC-EVs

7. Set the fluorescent plate reader as follows: the temperature at 37  C; the measurement condition as “Excitation: 320 nm; Emission: 405 nm” via the kinetic mode at 5 min intervals with plate-shaking prior to each fluorescence measurement. 8. Place the plate into the plate reader, and begin measurement. Usually, 60–120 min are sufficient to obtain the kinetics of the NEP enzymatic reaction of 5–10 μg hADSC-EVs (Fig. 2). The reaction reaches saturation after a longer time. 3.3 Transfer the Assay of hADSC-EVs to N2a Cells

Our laboratory determined whether the hADSC-EVs could be taken up by N2a cells by labeling the EVs with a fluorescent membrane linker. 1. Seed the N2a cells prior to EV administration (see Note 9). 2. Prepare 2 (100 μg/mL) EV suspension. As a negative control, prepare EV-free PBS( ). 3. Prepare 2 (4 μM) PKH solution in diluent C. 4. Mix the 2 EV suspension or EV-free PBS( ) and 2 PKH solution and incubate this mixture at room temperature for 5 min. 5. Transfer the mixture to a centrifugal ultrafiltration unit (100 kDa). 6. Centrifuge at 12,000  g at room temperature until the liquid passes the membrane to the collection tubes. 7. Remove the flowthrough and add 250 μL of filtrated PBS( ) (see Note 10). 8. Pipette 30–50 times to liberate EVs that are trapped on the membrane while being careful not to touch the membrane (see Note 11).

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Fig. 3 Transfer of hADSC-EVs into N2a cells. hADSC-EVs or vehicle PBS( ) as a control were labeled with PKH67, a green fluorescent membrane linker, and were added to the N2a cell cultures (approximately 2 μg/mL of hADSC-EVs at the final concentration). At 7 h after incubation, some of the cells were stained green (arrowheads). After 24 h, most of the N2a cells were stained green (arrowheads). Scale bar: 25 μm

9. Centrifuge at 12,000  g at room temperature until the liquid passes the membrane to the collection tubes. 10. Repeat steps (6)–(8) three more times (four washes in total). 11. Add 250 μL of N2a culture medium into the column and suspend the labeled EVs by pipetting it 30–50 times. 12. Place the column upside down onto a new collection tube. 13. Centrifuge at 12,000  g at room temperature for 1 min. 14. Dilute the retrieved EV suspension to a desired concentration with N2a culture medium. 15. Replace the culture medium of the N2a cells with medium containing the labeled EVs. 16. Culture the N2a cells for the desired time and observe them under fluorescent microscopy (Fig. 3). 3.4 Coculture of N2a Cells with hADSCs

We used mouse neuroblastoma N2a cells that were stably expressing both human APP695 with the Swedish mutation and presenilin 2 with the N141I mutation, which were established by Dr. Saido’s group (RIKEN Brain Institute, Japan) (28). These N2a cells over-produce both human Aβ40 and 42, thereby representing an in vitro AD model. The schematic outline for the experiment is shown in Fig. 4a.

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Fig. 4 Coculture of hADSCs and N2a cells. (a) Schematic representation of the coculture experiment. (b) The Aβ42 secretion levels of the N2a cells are decreased by coculturing them with hADSCs; for the control, hADSC-free medium was used. (c) The intracelluar Aβ42 levels in the N2a cells are also decreased in the coculture with hADSCs. These data are obtained by measuring the Aβ42 levels via ELISA followed by normalization with the cellular protein levels in the N2a cells in the corresponding wells

1. Harvest the N2a cells via trypsinization and resuspend them in a N2a cell culture medium at 4  104 cells/mL. 2. Seed the N2a cells to the lower chambers of 24-well plates at 2  104 cells/well (500 μL/well). 3. One day after the N2a cell inoculation, harvest the hADSCs via accutase treatment and resuspend them in MesenPRO at 2  104 cells/mL. 4. Place a cell culture insert possessing 0.4 μm pores in each well (see Note 12). 5. Seed the hADSCs onto the upper chambers at 6  104 cells/ well (300 μL/well). As a negative control, apply 300 μL/well of cell-free MesenPRO to the upper chambers. 6. Coculture the hADSCs and N2a cells for 2 days.

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7. Remove the upper chambers, and replace the medium in the lower chambers with fresh N2a cell culture medium (500 μL/well). 8. Replace the medium in the upper chambers with fresh MesenPRO (300 μL/well), and return them to the corresponding wells containing N2a cells. 9. Continue the coculture for another 8 h. 10. Remove the upper chamber from each well, and collect the culture supernatant from the lower chambers. 11. Centrifuge the collected culture supernatant at 2,000  g for 10 min to remove the cells and cellular debris. 12. Collect the supernatant and use it immediately for Aβ40 and 42 ELISA (see Note 13) or store it until use at 20  C. 13. Wash the cells on the lower chamber once with PBS( ). 14. Add 200 μL/well of M-PER, shake at room temperature for 10 min, collect the cell lysates into microtubes, and completely dissolve the cell lysates via periodic sonication on ice (see Note 14). 15. Determine the protein concentrations of the cell lysates using generally available protocols such as the Bradford assay or BCA assay (see Note 5). 16. Use the cell lysates immediately for Aβ40 and 42 ELISA or store them at 20  C until use. Representative data of the secreted and intracellular Aβ42 levels are shown in Fig. 4b.

4

Notes 1. For isolation, characterization, and routine culturing of hADSCs, see protocols that were previously published (29). 2. Prepare MesenPRO complete medium by supplementing 500 mL of basal medium with 10 mL growth supplement, 5 mL antibiotic-antimycotic, and 5 mL 100 GlutaMAX. 3. Prepare StemPRO MSC SFM complete medium by supplementing 415 mL of basal medium with 75 mL growth supplement, 5 mL antibiotic-antimycotic, and 5 mL 100 GlutaMAX. 4. Even without the addition of PBS( ), 100–150 μL PBS( ) remains in each tube. Thus, if a highly concentrated EV solution is needed for the subsequent experiments, avoid adding PBS( ). 5. Our laboratory routinely uses the BioRad Protein Assay® (BioRad) and Micro BCA Protein Assay® (Thermo) for the Bradford assay and BCA assay, respectively. 6. The presence of NEP protein in the hADSC-EVs can also be confirmed via immunoblotting using a general procedure with a commercially available antibody (Mouse monoclonal to CD10, clone number: 56C6) (25).

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7. The NEP-specific enzyme activity levels of the hADSCs-EVs can be estimated from rhNEP standard curves and represented as the value ng-rhNEP/mg-protein (25). 8. Prepare 2 mM substrate stock solution by dissolving the powder in dimethyl sulfoxide. Prepare 5 mg/mL thiorphan stock solution by dissolving the powder in ethanol. 9. N2a cells are routinely cultured in DMEM supplemented with 10 % FBS and 1 Antibiotic-antimycotic (Invitrogen). 10. To avoid the precipitation of free PKH in association with contaminating dust particles, remove any small dust particles by filtrating the PBS( ) with a 0.22-μm filter membrane. 11. Determine the number of pipettings required so that you do not observe any free PKH-derived stain on the membrane in PBS( ) control. Be consistent in the number of pipettings and pipetting intensity between the EV sample and PBS( ) control. 12. Be careful not to make bubbles under the cell culture inserts. 13. We performed ELISA using the Human beta-Amyloid (1-40) ELISA Kit wako II (Wako; 298-64601) and Human betaAmyloid (1-42) ELISA Kit wako (Wako; 298-62401) according to the manufacturer’s instructions. 14. We used the Ultrasonic Disrupter UR-20P. Sonication is performed on ice at power level 5 for 10 s. Sonication is usually repeated two times.

Acknowledgements This work was supported by a Grant-in-Aid for the Comprehensive Research and Development of a Surgical Instrument for the Early Detection and Rapid Curing of Cancer Project (P10003) of the New Energy and Industrial Technology Development Organization (NEDO). References 1. Raposo G, Nijman HW, Stoorvogel W et al (1996) B lymphocytes secrete antigenpresentingVesicles. J Exp Med 183:1161–1172 2. Zitvogel L, Regnault A, Lozier A et al (1998) Eradication of established murine tumors using a novel cell-free vaccine: dendritic cell derived exosomes. Nat Med 4:594–660 3. Katsuda T, Kosaka N, Ochiya T (2014) The roles of extracellular vesicles in cancer biology: towards the development of novel cancer biomarkers. Proteomics 14:412–425 4. Biancone L, Bruno S, Deregibus MC et al (2012) Therapeutic potential of mesenchymal

stem cell-derived microvesicles. Nephrol Dial Transplant 27:3037–3042 5. Katsuda T, Kosaka N, Takeshita F et al (2013) The therapeutic potential of mesenchymal stem cell-derived extracellular vesicles. Proteomics 13:1637–1653 6. Katsuda T, Ikeda S, Yoshioka Y et al (2014) Physiological and pathological relevance of secretory microRNAs and a perspective on their clinical application. Biol Chem 395:365–373 7. Chamberlain G, Fox J, Ashton B et al (2007) Concise review:mesenchymal stem cells: their phenotype, differentiation capacity,

Potential Application of Extracellular Vesicles of Human Adipose. . . immunological features, and potential for homing. Stem Cells 25:2739–2749 8. Djouad F, Bouffi C, Ghannam S et al (2009) Mesenchymal stem cells: innovative therapeutic tools for rheumatic diseases. Nat Rev Rheumatol 5:392–399 9. Salem HK, Thiemermann C (2010) Mesenchymal stromal cells: current understanding and clinical status. Stem Cells 28:585–596 10. Psaltis PJ, Zannettino ACW, Worthley SG et al (2008) Concise review: mesenchymal stromal cells: potential for cardiovascular repair. Stem Cells 26:2201–2210 11. Bruno S, Grange C, Deregibus MC et al (2009) Mesenchymal stem cell-derived microvesicles protect against acute tubular injury. J Am Soc Nephrol 20:1053–1067 12. Gatti S, Bruno S, Deregibus MC et al (2011) Microvesicles derived from human adult mesenchymal stem cells protect against ischaemiareperfusion-induced acute and chronic kidney injury. Nephrol Dial Transplant 26:1474–1483 13. Bruno S, Grange C, Collino F et al (2012) Microvesicles derived from mesenchymal stem cells enhance survival in a lethal model of acute kidney injury. PLoS One 7:e33115 14. He J, Wang Y, Sun S et al (2012) Bone marrow stem cells-derived microvesicles protect against renal injury in the mouse remnant kidney model. Nephrology (Carlton) 17:493–500 15. Zhou Y, Xu H, Xu W et al (2013) Exosomes released by human umbilical cord mesenchymal stem cells protect against cisplatin-induced renal oxidative stress and apoptosis in vivo and in vitro. Stem Cell Res Ther 4:34 16. Lai RC, Arslan F, Lee MM et al (2010) Exosome secreted by MSC reduces myocardial ischemia/ reperfusion injury. Stem Cell Res 4:214–222 17. Lai RC, Arslan F, Tan SS et al (2010) Derivation and characterization of human fetal MSCs: an alternative cell source for large-scale production of cardioprotective microparticles. J Mol Cell Cardiol 48:1215–1224 18. Arslan F, Lai RC, Smeets MB et al (2013) Mesenchymal stem cell-derived exosomes increase ATP levels, decrease oxidative stress and activate PI3K/Akt pathway to enhance myocardial viability and prevent adverse remodeling after myocardial ischemia/reperfusion injury. Stem Cell Res 10:301–312 19. Xin H, Li Y, Buller B et al (2012) Exosome mediated transfer of miR-133b from

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multipotent mesenchymal stromal cells to neural cells contributes to neurite outgrowth. Stem Cells 30:1556–1564 20. Xin H, Li Y, Liu Z et al (2013) Mir-133b promotes neural plasticity and functional recovery after treatment of stroke with multipotent mesenchymal stromal cells in rats via transfer of exosome-enriched extracellular particles. Stem Cells 31:2737–2746 21. Xin H, Li Y, Cui Y et al (2013) Systemic administration of exosomes released from mesenchymal stromal cells promote functional recovery and neurovascular plasticity after stroke in rats. J Cereb Blood Flow Metab 33:1711–1715 22. Lee C, Mitsialis SA, Aslam M et al (2012) Exosomes mediate the cytoprotective action of mesenchymal stromal cells on hypoxiainduced pulmonary hypertension. Circulation 126:2601–2611 23. Zhu Y, Feng X, Abbott J et al (2014) Human mesenchymal stem cell microvesicles for treatment of Escherichia coli endotoxin‐induced acute lung injury in mice. Stem Cells 32:116–125 24. Islam MN, Das SR, Emin MT et al (2012) Mitochondrial transfer from bone-marrowderived stromal cells to pulmonary alveoli protects against acute lung injury. Nat Med 18:759–765 25. Katsuda T, Tsuchiya R, Kosaka N et al (2013) Human adipose tissue-derived mesenchymal stem cells secrete functional neprilysin-bound exosomes. Sci Rep 3:1197 26. Iwata N, Higuchi M, Saido TC (2005) Metabolism of amyloid-beta peptide and Alzheimer’s disease. Pharmacol Ther 108:129–148 27. Miners JS, Verbeek MM, Rikkert MO et al (2008) Immunocapture-based fluorometric assay for the measurement of neprilysin-specific enzyme activity in brain tissuehomogenates and cerebrospinal fluid. J Neurosci Methods 167:229–236 28. Shirotani K, Tsubuki S, Iwata N et al (2001) Neprilysin degrades both amyloid beta peptides 1-40 and 1-42 most rapidly and efficiently among thiorphan- and phosphoramidonsensitive endopeptidases. J Biol Chem 276:21895–21901 29. Banas A (2012) Purification of adipose tissue mesenchymal stem cells and differentiation toward hepatic-like cells. Methods Mol Biol 826:61–72

Methods in Molecular Biology (2015) 1212: 183–193 DOI 10.1007/7651_2014_95 © Springer Science+Business Media New York 2014 Published online: 26 July 2014

Application of Fluid Mechanical Force to Embryonic Sources of Hemogenic Endothelium and Hematopoietic Stem Cells Nan Li, Miguel F. Diaz, and Pamela L. Wenzel Abstract During embryonic development, hemodynamic forces caused by blood flow support vascular remodeling, arterialization of luminal endothelium, and hematopoietic stem cell (HSC) emergence. Previously, we reported that fluid shear stress plays a key role in stimulating nitric oxide (NO) signaling in the aorta-gonadmesonephros (AGM) and is essential for definitive hematopoiesis. We employed a Dynamic Flow System modified from a cone-and-plate assembly to precisely regulate in vitro exposure of AGM cells to a defined pattern of laminar shear stress. Here, we present the design of a microfluidic platform accessible to any research group that requires small cell numbers and allows for recirculation of paracrine signaling factors with minimal damage to nonadherent hematopoietic progenitors and stem cells. We detail the assembly of the microfluidic platform using commercially available components and provide specific guidance in the use of an emerging standard in the measurement of embryonic HSC potential, intravenous neonatal transplantation. Keywords: Hemogenic endothelium, Hematopoietic stem cells, HSC, Biomechanical forces, Shear stress, AGM

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Introduction Accumulating evidence suggests that definitive hematopoietic stem cells (HSCs) derive from a specialized population of endothelial cells, termed hemogenic endothelium (1–7). These cells transiently appear in specific sites of the embryonic vasculature and undergo endothelial-to-hematopoietic transition (EHT) by specific, but relatively poorly understood, mechanisms (8). Recent studies have suggested that HSC maturation in the aorta-gonads-mesonephros (AGM) region is regulated by hemodynamic forces created by blood flow (9, 10). Fluid mechanical forces induce expression of hematopoietic markers and blood forming activity in explant culture, whereas cardiac mutants with impaired blood flow exhibit significant deficiency in definitive hematopoiesis (9, 10). Nitric oxide (NO) plays an important role in triggering signaling events downstream of biomechanical force, as interruption of NO production impairs

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blood formation and NO donor compounds are capable of partial restoration of hematopoietic activity in cardiac mutants (9, 10). However, how hematopoiesis is regulated by blood flow and whether other mechanosensitive pathways contribute to fate commitment remain critical questions in the field of developmental hematopoiesis. Three mutually perpendicular hemodynamic forces exist within the vasculature (11). First, blood pressure exerts a hydrostatic force against the vessel wall. Second, vessel wall cells experience circumferential stretch imposed by hydrostatic pressure. Finally, blood flow exerts drag on the luminal wall and causes a frictional force between fluid and vessel surface cells, referred to as shear stress. Among these biomechanical forces, shear stress has been shown to promote hematopoiesis in mouse AGM and from embryonic stem cells (9, 10, 12). Studies of cellular response to shear stress can be challenging due to the specialized instrumentation required for generation of defined fluid flow intensities. Further, recirculation of culture medium is necessary for capture of paracrine signaling mechanisms stimulated by shear stress and translates to a substantial cost savings in reagents with prolonged culture. Peristaltic pumps fall short of meeting the demands of fluid management as they introduce refractory pulsatile patterns in shear stress and damage nonadherent cells that circulate through the roller mechanism. Here we describe a microfluidic laminar shear stress system capable of long-term recirculation of culture medium, accessible to any laboratory and reliant upon commercially available channel slides and one syringe pump. We describe use of the platform for several outcome measures and provide specific guidance in the use of neonatal transplantation, an emerging standard in functional assessment of embryonic HSCs.

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Materials The products and vendors listed here have been employed in our research; however, suitable alternatives may exist for microfluidic culture platforms and syringe pumps. Custom microfluidics could easily be adapted to the system with special attention to prevention of bubble formation.

2.1

AGM Culture

1. Mouse embryos. 2. Accutase cell detachment solution (Stem Cell Research, catalog # 07920). 3. Trypan Blue Solution, 0.4 % (Invitrogen/Life Technologies, catalog # 15250-061).

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4. Sodium pyruvate, 100 mM (Invitrogen/Life Technologies, catalog # 11360-070). 5. MEM nonessential amino acids solution (Invitrogen/Life Technologies, catalog # 11140-050). 6. HEPES buffer solution, 1 M (Invitrogen/Life Technologies, catalog # 15630-080). 7. Penicillin/Streptomycin solution (VWR, catalog # 45000-652). 8. MyeloCult M5300 cell culture medium (Stem Cell Research, catalog # 05350). Add sodium pyruvate (5 ml), MEM nonessential amino acids (5 ml), HEPES (12.5 ml), and Penicillin/ Streptomycin (5 ml) before use. 9. BD Falcon cell strainer, 70 μm mesh (Fisher Scientific, catalog # 08-771-2). 2.2 Laminar Shear Stress Assembly

1. IBIDI VI 0.4 six channel slide (IBIDI, catalog # 80606). 2. Matrigel Basement Membrane Matrix (BD Biosciences, catalog # 354234). 3. Fibronectin human protein, plasma (Invitrogen/Life Technologies, catalog # 33016-015). 4. PBS buffer, 500 ml (Invitrogen/Life Technologies, catalog # 10010-023). 5. Recombinant mouse SCF (mSCF), 10 μg/ml (PeproTech, catalog # 250-03). 6. Check valve 0.27 PSI CP (Qosina, catalog # 80184). 7. Three way stop cock, three female luer lock (Qosina, catalog # 99699). 8. Silicone tubing, 1/16 ID (Fisher Scientific, catalog # 1118915G). 9. Barbed T-connector, polypropylene, 1/16-in. (Cole Parmer, catalog # YO-06365-77). 10. Barbed Y-connector, PVDF (Cole Parmer, catalog # T30703-90). 11. Female Luer x 1/16 hose barb, PVDF (Cole Parmer, catalog # T45512-00). 12. Elbow Luer connector (IBIDI, catalog # 10802). 13. Syringe, 10 ml (BD, catalog # 309604). 14. Syringe pump PHD ULTRA 4400 with Remote (Harvard Apparatus, catalog # 703310).

2.3 Neonatal Transplantation

Hamilton syringe, TLL, 50 ml (FISHER, catalog # 14815-57). 30-G needle (FISHER, catalog # 14826 F).

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Methods Carry out all procedures at room temperature unless otherwise specified.

3.1 AGM Tissue Dissociation

1. Carefully dissect out the AGM region from early mouse embryos (e.g. E10.5) in ice-cold PBS. 2. Remove excess PBS (see Note 1) and add 2–4 ml Accutase solution for AGM tissues from approximately ten embryos. 3. Incubate tissues on a plate shaker for 20 min (see Note 2). 4. Gently pipette 5 times and shake for 3–5 more min. 5. After incubation, pipette ten more times very gently to obtain a single cell suspension. Add 4 ml Myelocult medium and mix well. 6. Filter suspension through 70-μm nylon cell strainers. 7. Collect 10 μl of cell suspension and count live cells using trypan blue. Calculate total cell number. 8. Centrifuge cells at 900  g for 5 min and resuspend in Myelocult medium at a final concentration of 25  106 cells/ml. 9. Seed cells into microfluidic slides or flow chambers immediately, as described below.

3.2 Laminar Shear Stress Assembly

1. Sterilize tubing and check valves for shearing before the experiment (see Note 3). Setup for one laminar shear stress (LSS) channel requires the following materials: One 8-in. tubing. Two 6-in. tubing. Six 3-in. tubing. Three T or Y connectors. Two elbow connectors. Four check valves. Two 3-way stopcocks. Six female luer  1/16 hose barb connector. One 14 ml round bottom tube with cap. One 10 ml Syringe. One Harvard Apparatus syringe pump. Each low-flow control channel requires the following materials: One 8-in. tubing. One 6-in. tubing. One female luer  1/16 hose barb connector. Two elbow connectors.

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2. Prior to seeding cells, coat IBIDI slide channels with 50 μl of Matrigel matrix, diluted 1:20 in PBS (see Note 4). Incubate for at least 45 min at 37  C or room temperature. Wash channels twice with PBS. 3. Inject 30 μl of cell suspension into each channel of an IBIDI VI 0.4 slide (see Note 5). Place the slide inside a 15 cm dish along with a 35 mm dish (no lid) filled with PBS to maintain humidity (see Note 6). Incubate at 37  C for 8 h to allow attachment to the culture surface. 4. Each channel requires a total volume of 10 ml Myelocult media containing 10 ng/ml mSCF for recirculation during application of LSS. Prepare an additional 3–5 ml of media with mSCF per channel to flush out nonadherent cells prior to starting fluidics. 5. Proceed with flushing of each channel by pipetting the corresponding channel volume into the slide inlet (see Note 7). Remove the excess media from the opposite end of the channel, but avoid removing all media and drying out the channel. Repeat flushing two more times. Afterwards, top off the channel and inlets with the media. When done, place the slide back in the incubator. 6. Set up all sterilized tubing and check valves under the hood while the slide is in the incubator warming (Figs. 1 and 2a). Assembly can be done on the incubator shelf if removed and placed under the hood. The two elbow connectors, which will connect to the slide through the inlets for one channel, must first be assembled to two female luer inlets of a 3-way stopcock (Fig. 2b). The third inlet of the stopcock is set to the “off” position. 7. Fill the reservoir tube with 10 ml myelocult-SCF media. Cut a small hole in the tube cap and insert the tubing ends into the reservoir (Fig. 2c). Apply parafilm around the cap and wrap carefully around the junction with the tubing. Secure the parafilm with tape, but be careful not to constrict the tubing, which would prevent media flow. Infuse Withdraw 3 way stopcock

Si tubing Check valve IBIDI Slide Y / T connector

Reservoir

Fig. 1 Schematic diagram of the microfluidic unidirectional shear stress system using a single syringe

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Fig. 2 Laminar shear stress tube assembly. (a) Overview of the tube and check valve assembly. The second 3-way stopcock (red box) is temporary and serves as a “place-holder” for the future site of the slide. (b) Two elbow connectors (arrows) are assembled to two female luer inlets of a 3-way stopcock before connecting to a slide. The third inlet (red “NO” symbol) is closed. (c) Ensure the tubing in the media reservoir is close to but not in contact with the bottom of the 15 ml tube. (d) To attach slide, first clamp the tubing on either side of the stopcock, then remove the stopcock. (e) Connect the tube assembly to a slide by inserting the elbow connectors (arrows) into the two inlets of a channel (a 6-channel IBIDI VI0.4 slide is depicted)

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8. Hold the syringe by hand in a vertical position and repeatedly infuse and withdraw to fill the tubing assembly. Several repetitions will remove all bubbles from the assembly and syringe. Once this is done, set the syringe plunger to the 5 ml mark (see Note 8). 9. Move the slide into the hood and secure it to a flat surface. Clamp the tubing behind the elbow connectors with binder clips (see Note 9; Fig. 2d). Remove the 3-way stopcock and connect the elbow connectors to the inlets of the slide. Secure the junctions and then remove the clamps. 10. Check the tubing end inside the reservoir. Make sure it is immersed in media and not in contact with the bottom surface of the tube (Fig. 2c). 11. Secure the syringe in the syringe pump (Fig. 2e). Place the entire assembly inside the tissue culture incubator. If the incubator shelf was placed in the tissue culture hood, the entire LSS assembly and syringe pump can be placed on top and transferred into the incubator. 12. Connect the syringe pump to the Harvard Apparatus PHD ULTRA 4400 Remote, if using a remote system. 13. Set up a shearing program. Set the syringe pump force at 50 %. Select the correct syringe characteristics by brand and size of the syringe. The infuse/withdrawal run program is listed as “Autofill.” Set the “Set Mode” to INF/WD. Under “Set Rates,” the INF and WD will be the same rate. For 5 dyn/ cm2 on the IBIDI VI0.4 slide, the rate is 2.84 ml/min. Under “Set Volume per Cycle” enter the same volume for “Set Rates” (e.g. 2.84 ml) (see Note 10). Shearing time can be set to the number of cycles or the period of the time, so under “Set Total Volume of # Cycles” select Cycles or Time. Enter the cycle number or shearing time. 14. A low flow control can be set up by directly connecting a syringe containing medium to one inlet and a flow-out tubing to the other. Insert the tubing into a receiving tube to collect flow-through medium. Set the same pump force and syringe characteristics. Set the “Set Mode” to INF. For 0.001 dyn/cm2 on the IBIDI VI0.4 slide, the rate is 600 nl/min. 15. Press “Start” on the syringe control screen to start shearing and low flow control. Wait 10 min after the run starts and check the tubing assembly to make sure there are no leaks, bubbles, or errors on the pump. 16. Following application of shear stress, collect the cells by Accutase treatment and analyze by standard functional and phenotypic methods such as colony formation in methylcellulose, flow cytometry, gene expression profiling, or transplantation (described below).

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3.3 Neonatal Transplantation

1. Set up the male and female(s) that are to provide the neonates 3 weeks exactly from the day of transplantation (see Note 11). 2. Sublethal neonatal myeloablation is typically done the day after birth (see Note 12). Approximately 3 h before transplantation, remove the mother from neonates, transfer pups into an irradiator dish and irradiate with dosing listed below. Put pups back into the cage, sprinkle with bedding, then reintroduce the mother. Change gloves if handling multiple adults and/or neonatal litters. (a) NSG neonates: 100 rads. (b) Rag2-/- Common Gamma-/- neonates: 250 rads. (c) C57Bl6 and B6.SJL (BoyJ): 300 rads. 3. Dissociate cells from slides by Accutase treatment. 4. Run cells through a 70-μm strainer. 5. Resuspend cells in PBS volume sufficient for 15 μl per pup (see Note 12). Add a dose for an additional pup because volume can be “lost” in the syringe or during handling. 6. Remove the mother into a clean cage (see Note 13). 7. Place one or two pups on ice. Carefully observe them for color and movement. They are ready for injection when they appear slightly purple and are nonresponsive to handling or toe pinch. The facial vein is easiest to target when purple in color (see Note 14). 8. Inject 15 μl via the superior temporal vein of the face with a Hamilton syringe and 30-G needle (Fig. 3). Use care to remove air bubbles from the solution, which could result in immediate death if injected (see Note 15).

Fig. 3 Facial vein injection of a neonatal mouse. The superior temporal vein highlighted by the needle tip serves as the best target for injection of the cell suspension

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9. Transfer pups onto gentle heat. Cold stainless steel under the pups should be insulated by a heating pad (on LOW setting) or other material that can absorb heat from the heat lamp. Pups will regain pink tone and be very mobile. Be careful that they do not move away into a dangerous area (see Note 16). 10. Accumulate all the pups in the recovery area under the heat lamp, and then transfer together back into the cage. Sprinkle the pups with bedding to mask odors from gloves, and move the mother back in to join them. 11. Analysis of peripheral blood chimerism can be performed by flow cytometry beginning 5–6 weeks after transplantation, depending upon size and condition of pups, and should be monitored longterm (12–20 weeks) for assessment of stem cell function.

4

Notes 1. Residual PBS buffer can be left in the tube as it does not have negative impact on dissociation efficiency. 2. Incubation time is important. Do not exceed 25 min, as cells may be damaged with excessive enzymatic digestion. 3. Be sure to sterilize tubing, connectors, 3-way stopcocks, and check valves. Tubing and all connectors are autoclavable up to 121  C for 20 min. The check valves and 3-way stopcocks are sterilized by Ethylene Oxide (EtO) gas. 4. Slide channels and flow chambers need to be coated first and washed with media to encourage cells to quickly adhere. A common coating solution includes matrigel (1: 20 dilution) and fibronectin (100 μg/ml) and must be freshly prepared in PBS on ice. 5. Please refer to IBIDI product manual for all information about the slides, medium volumes, and shearing parameters. 6. This provides moisture for the slides to prevent evaporation and drying of the channel. 7. Flushing the channels aids in removal of floating cells and nonadherent clumps of erythrocytes that should be removed prior to application of flow. 8. It is crucial that all syringes are set at the same mark. 9. This prevents leakage of media from the tubing. 10. Each cycle is equal to one full infusion and withdrawal, and lasts 2 min. Thus, for 1 h of LSS, 30 cycles are required. The rate table for shear stress is available in the IBIDI product manual. 11. Adult transplantation can be used in lieu of neonatal transplantation, though emerging evidence suggests that the neonatal environment may provide a more sensitive readout of early hematopoietic stem and progenitor cells (13).

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12. Transplant is least challenging the day after birth. As the pups age, injections become more difficult. 13. Transplant 0.5–2 million cells per neonate, depending upon the cell source and frequency of engraftable cells anticipated in the cell suspension. Trace amounts of serum are not detrimental but should not be added to the PBS for resuspension, as this could cause inflammatory response. 14. Remove the mother from the area so that she is unaware of the handling of her pups. This includes sight, smell, and sound. 15. If delays are encountered, transfer pups to heat and start over after full recovery. Over-chilled pups may be difficult to inject due to vessel constriction and may not revive. Each pup can tolerate cooling two times, if necessary. If a pup has been on ice over 15 min, transfer to heat. 16. Two aspects are critical during the facial injection procedure: (a) positioning of the pup, and (b) minimal movement of the hand while pushing the plunger. The beveled edge of the needle should face upward away from the pup. Try to secure the skin of the head to resist the needle as it enters the vein. If skin is stretched too tightly, the vein will “disappear.” If the needle is well placed in the vein, the vein will clear (red disappears) as cell suspension is injected. If the needle misses the vein, a fluid bubble will form during injection. Engraftment has been observed even when the vein is missed. 17. Warming the pups after immobilizing/anesthetizing on ice is critical. Care should be taken to avoid scorching pups under the heat lamp, and they should be warmed gently from above and below. Circulating water warming pads are particularly well suited to this application, as they provide gentle, uniform heat.

Acknowledgments This work was funded by grants from the American Society of Hematology, State of Texas Emerging Technology Fund, and National Institutes of Health to P.L.W. References 1. Zovein AC, Hofmann JJ, Lynch M, French WJ, Turlo KA, Yang Y et al (2008) Fate tracing reveals the endothelial origin of hematopoietic stem cells. Cell Stem Cell 3:625–636 2. Eilken HM, Nishikawa S, Schroeder T (2009) Continuous single-cell imaging of blood generation from haemogenic endothelium. Nature 457:896–900 3. Lancrin C, Sroczynska P, Stephenson C, Allen T, Kouskoff V, Lacaud G (2009). The

haemangioblast generates haematopoietic cells through a haemogenic endothelium stage. Nature 457:892–895 4. Bertrand JY, Chi NC, Santoso B, Teng S, Stainier DY, Traver D (2010) Haematopoietic stem cells derive directly from aortic endothelium during development. Nature 464:108–111 5. Boisset JC, van Cappellen W, Andrieu-Soler C, Galjart N, Dzierzak E, Robin C (2010) In vivo imaging of haematopoietic cells emerging

Application of Mechanical Force from the mouse aortic endothelium. Nature 464:116–120 6. Kissa K, Herbomel P (2010) Blood stem cells emerge from aortic endothelium by a novel type of cell transition. Nature 464:112–115 7. Lam EY, Hall CJ, Crosier PS, Crosier KE, Flores MV (2010) Live imaging of Runx1 expression in the dorsal aorta tracks the emergence of blood progenitors from endothelial cells. Blood 116:909–914 8. Swiers G, Rode C, Azzoni E, de Bruijn MF (2013) A short history of hemogenic endothelium. Blood Cells Mol Dis 51:206–212 9. Adamo L, Naveiras O, Wenzel PL, McKinneyFreeman S, Mack PJ, Gracia-Sancho J et al (2009) Biomechanical forces promote embryonic haematopoiesis. Nature 459:1131–1135

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10. North TE, Goessling W, Peeters M, Li P, Ceol C, Lord AM et al (2009) Hematopoietic stem cell development is dependent on blood flow. Cell 137:736–748 11. Culver JC, Dickinson ME (2010) The effects of hemodynamic force on embryonic development. Microcirculation 17:164–178 12. Wolfe RP, Ahsan T (2013) Shear stress during early embryonic stem cell differentiation promotes hematopoietic and endothelial phenotypes. Biotechnol Bioeng 110:1231–1242 13. Yoder MC, Hiatt K, Dutt P, Mukherjee P, Bodine DM, Orlic D (1997) Characterization of definitive lymphohematopoietic stem cells in the day 9 murine yolk sac. Immunity 7:335–344

Methods in Molecular Biology (2015) 1212: 195–200 DOI 10.1007/7651_2014_96 © Springer Science+Business Media New York 2014 Published online: 26 July 2014

Electrophysiological Recordings from Neuroepithelial Stem Cells Masayuki Yamashita Abstract During embryonic development, neuroepithelial cells act as neural stem cells by renewing themselves. These cells are tightly interconnected and make contact with the basement membrane of the neuroepithelium. Under such circumstances, intracellular recording with a fine-tipped microelectrode is a successful method to study the electrophysiological properties of the neuroepithelial stem cell. This chapter describes the detailed techniques of intracellular recording from neuroepithelial stem cells. Keywords: Intracellular recording, Neuroepithelium, Embryo, Membrane potential, Microelectrode, Ion channel

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Introduction Neuroepithelieal cells act as neural stem cells by renewing themselves during embryonic development (1, 2). The central nervous systems, which include brain, retina, and spinal cord are derived from the neuroepithelium. The neuroepithelial cells are tightly interconnected by adherens junctions and electrically coupled through gap junctions at the apical portion of the cell. The apical process of neuroepithelial cells faces the ventricle and the basal process makes contact with the basement membrane, which is essential for the proliferation activity of the cell. The basement membrane is made of collagen and laminin to form a mechanical and electrical boundary of the neuroepithelium, which maintains ionic circumstances around the neuroepithelial cell. Thus, dissociation of the cells for patch clamp recording is not a suitable way of studying physiological properties of neuroepithelial cells. Alternatively, intracellular recording with a fine-tipped microelectrode is a successful method to study the electrophysiological properties of neuroepithelial cells and the ion channel activities in these cells. The intracellular recording was applied to the cells in the neural plate and the neural tube of the amphibian embryo to study the electrical properties of these cells (3).

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This chapter describes the detailed techniques of intracellular recording from neuroepithelial cells. To access to neuroepithelial cells, the isolation of a neuroepithelium from early embryos allows clear visualization of these cells for precise impalement of a microelectrode into the cell. For this purpose, the retinal neuroepithelium is an accessible part of the neural tube; it can be isolated from early embryos and seems to be a suitable model for studying the electrophysiological properties of neuroepithelial cells. We have revealed the response of retinal neuroepithelial cells to the neurotransmitter GABA and the electrical cell–cell communication between them with the intracellular recording technique in the retinal neuroepithelium of early embryonic chicks (4, 5).

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Materials Prepare not only electrical recording systems but also optical and mechanical apparatus, which are essential for clear visualization of neuroepithelial cells and microelectrode tips and for stable and successful recordings from these cells.

2.1 Recording Apparatus

1. Vibration isolator: a heavy table supported by cushions of compressed air or nitrogen gas. 2. Microscope: an upright microscope with a high-magnification water immersion objective (100) and a low-magnification objective (10) (see Note 1). 3. Recording chamber (see Note 2). 4. A pair of tungsten needles and two micromanipulators to position a retinal neuroepithelium on the glass bottom of the recording chamber. 5. Perfusion system including solenoid valves to change bath solutions from the normal bath solution to test solutions (see Note 3). 6. Temperature controller: an inline heater to warm the superfusate, a power controller regulated by a feedback circuit, and a thermistor probe, which is positioned near the specimen with another micromanipulator.

2.2

Amplifier

1. Intracellular recording amplifier (see Note 4). 2. DC amplifier (see Note 5). 3. High cut filter (see Note 6). 4. Digital recording system (see Note 7).

2.3

Microelectrodes

1. Glass capillary (see Note 8). 2. Micropipette puller (see Note 9).

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3. A three-dimensional micromanipulator to position the tip of a microelectrode near the basement membrane of the neuroepithelium (see Note 10). 4. An advancer of a microelectrode (see Note 11). 2.4

Solutions

1. The normal bath solution: 137 mM NaCl, 5 mM KCl, 2.5 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, and 22 mM glucose, buffered to pH 7.3 by adding NaOH. Store at 4  C. 2. Solutions to fill microelectrodes: 3 M KCl or 4 M potassium acetate. Store at 4  C.

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Methods Carry out all the procedures as quickly as possible. If good recordings are not available, discard the specimen and make a new preparation. It takes about one hour to finish one recording session including the preparation.

3.1 Preparation of Retinal Neuroepithelium

1. Pick up an embryo from an egg incubated for 3 days. 2. Put the chick embryo on the bottom of a 35 mm plastic dish containing the normal bath solution. 3. Isolate an optic cup together with the lens vesicle from the embryo with a pair of fine forceps under an operation microscope. 4. Transfer the optic cup with a Pasteur pipette to another 35 mm plastic dish containing the normal bath solution. 5. Take off the lens vesicle with fine forceps. 6. Separate the inner wall of the optic cup, which corresponds to the retinal neuroepithelium, from the outer wall of the optic cup, which becomes the pigment epithelium (see Note 12). 7. Transfer the retinal neuroepithelium with a Pasteur pipette to the recording chamber. 8. Mount the recording chamber on the stage of the microscope. 9. Position the retinal neuroepithelium with the inner side up, so that the inner limiting membrane, which is the basement membrane of the retinal neuroepithelium, is penetrated with a microelectrode under the high-magnification objective. 10. Fix the specimen with a pair of tungsten needles at the center of the recording chamber. 11. Put the reference electrode in the bath solution. 12. Position the thermistor probe near the specimen. 13. Start perfusion of the normal bath solution. 14. Set the temperature of the bath solution at 38  C, if necessary.

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Fig. 1 Nomarski optics (DIC) view of the retinal neuroepithelium. A glass microelectrode is approaching the inner limiting membrane of the retinal neuroepithelium. An arrow indicates the inner limiting membrane (reproduced from (8) with permission from Elsevier)

3.2 Preparation of Microelectrodes

1. Pull a glass capillary with the micropipette puller (see Note 13). 2. Fill the micropipette with an electrode solution (see Note 14). 3. Attach the microelectrode to the micropipette holder.

3.3 Intracellular Recording

1. Position the microelectrode tip near the specimen with the three-dimensional micromanipulator under the lowmagnification objective. 2. Position the microelectrode tip near the inner limiting membrane with the three-dimensional micromanipulator under the high-magnification objective (Fig. 1). 3. Adjust the DC potential level to zero. 4. Adjust the high-frequency compensation appropriately (see Note 15). 5. Penetrate the basement membrane with the microelectrode by advancing it into the neuroepithelium (see Note 16). 6. Insert the microelectrode into a cell, which results in a sudden appearance of the resting membrane potential. 7. Change the membrane potential by changing the amplitude of injection currents (see Note 17). 8. Change bath solutions from the normal bath solution to a test solution. 9. Store the data. 10. If dye injection is necessary to identify the recorded cell, fluorescent dyes can be injected electrophoretically through the microelectrode into the recorded cell (see Note 18).

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Notes 1. Microelectrode impalements are made under a microscope using a long working distance objective (2.5 mm) that allows sufficient space for microelectrode access. The stage of the microscope should be fixed on the table of the vibration isolator. The microscope is mounted on a two-dimensional micromanipulator, which is fixed on the table of the vibration isolator. The microscope is moved in X-Y directions with the two-dimensional micromanipulator. 2. The recording chamber is mounted on the fixed stage of the microscope. The volume of the recording chamber is 0.2 mL. 3. The solution is perfused by a gravity feed at 2 mL/min. The outflow from the recording chamber is aspirated through a drain tube by negative pressure. 4. The gain of the intracellular amplifier is 10. A high-frequency compensation circuit is required to compensate the electrode capacitance. A bridge balance circuit is required to compensate the electrode resistance. Constant currents are injected through the microelectrode into the cell in DC or pulse modes. The amplitude of injection currents ranges from 1,000 pA (hyperpolarizing currents) to 1,000 pA (depolarizing currents). 5. The gain of the DC amplifier is 5 or 10. 6. Cut off frequency: 300 Hz, 500 Hz, 1 KHz, 3 KHz, and 10 KHz. 7. A-D sampling rate: 10 KHz or more. 8. Borosilicate glass capillary is suitable for fine-tipped micropipettes. The outer diameter of the glass capillary is 1.5 mm and the inner diameter is 0.86, and 100 mm in length. The glass capillary should contain a thin filament fused to the inner surface of the glass capillary. An electrode solution flows towards the tip by capillary action of the filament after the solution is injected at the shoulder of the micropipette. Both ends of the capillary should be fire-polished not to scratch the surface of an Ag/AgCl wire, which is inserted into the micropipette. 9. Flaming/Brown micropipette puller (Sutter Instrument Co.) is recommended to make fine-tipped micropipettes for intracellular recording. 10. Mechanical manipulators for laser optics experiments are sturdy enough. The fineness of the X-Y motions depends on the micrometers of the manipulator (6). They should be attached to the table of the vibration isolator by magnetic bases.

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11. Piezo-drive linear actuator provides stable and precise motions of microelectrodes without mechanical drift. 12. It is also possible to record from neuroepithelial cells without the isolation of the outer wall of the optic cup. 13. Prepare microelectrodes just before recordings. The tip and surface of micropipettes may be blurred with air dust. 14. To make a fine tube to fill a micropipette with an electrode solution, heat a 1 mL plastic syringe by fire, pull it and cut. After sucking the electrode solution, insert the fine tube into the micropipette and fill it with the electrode solution. 15. Pass current pulses (2 ms in duration) through the electrode, and adjust the high-frequency compensation so that the voltage change becomes square pulses. 16. If it is difficult to penetrate the basement membrane by advancing the microelectrode, change the specimen or the electrode. 17. Bridge balance should be adjusted to cancel the voltage change due to the electrode resistance, which ranges from 100 to 200 MΩ. 18. Alexa Fluor 488 hydrazide can be injected by passing hyperpolarizing current pulses ( 500 to 1,000 pA in amplitude, 400 ms in duration at 800 ms interval) for 0.5–3 min (5, 7). The microelectrode should be filled with a low-concentration KCl solution (200 mM) containing 1 % Alexa Fluor 488 hydrazide. References 1. Yamashita M (2012) Ion channel activities in neural stem cells of the neuroepithelium. Stem Cells Int 2012:847670. doi:10.1155/2012/ 247670 2. Yamashita M (2013) From neuroepithelial cells to neurons: changes in the physiological properties of neuroepithelial stem cells. Arch Biochem Biophys 534:64–70 3. Warner AE (1973) The electrical properties of the ectoderm in the amphibian embryo during induction and early development of the nervous system. J Physiol 235:267–286 4. Yamashita M, Fukuda Y (1993) Calcium channels and GABA receptors in the early embryonic chick retina. J Neurobiol 24:1600–1614

5. Yamashita M (2008) Synchronous Ca2+ oscillation emerges from voltage fluctuations of Ca2+ stores. FEBS J 275:4022–4032 6. Brown KT, Flaming DG (1986) Advanced micropipette techniques for cell physiology. In: Smith AD (ed) IBRO handbook series: methods in the neurosciences, vol 9. Wiley, New York, NY 7. Mobbs P et al (1994) Techniques for dye injection and cell labelling. In: Ogden D (ed) Microelectrode techniques, 2nd edn, The Plymouth workshop handbook. The company of Biologists Limited, London, pp 361–387 8. Yamashita M (2013) Electric axon guidance in embryonic retina: galvanotropism revisited. Biochem Biophys Res Commun 431:280–283

Methods in Molecular Biology (2015) 1212: 201–207 DOI 10.1007/7651_2014_120 © Springer Science+Business Media New York 2014 Published online: 11 September 2014

In Vivo Stem Cell Transplantation Using Reduced Cell Numbers Takeo W. Tsutsui Abstract Dental pulp stem cell (DPSC) characterization is essential for regeneration of a dentin/pulp like complex in vivo. This is especially important for identifying the potential of DPSCs to function as stem cells. Previously reported DPSC transplantation methods have used with huge numbers of cells, along with hydroxyapatite/tricalcium phosphate (HA/TCP), gelatin and fibrin, and collagen scaffolds. This protocol describe a transplantation protocol that uses fewer cells and a temperature-responsive cell culture dish. Keywords: Dental pulp stem cells (DPSCs), Temperature-responsive cell culture dish, Hydroxyapatite/tricalcium phosphate (HA/TCP), Transplantation

1

Introduction DPSCs differentiate into multiple stromal cell lineages that can be easily recruited and used as a source material for regenerative medicine. The first report of DPSCs revealed their ability to regenerate a dentin/pulp-like structure in immunocompromised mice (1). Several methods for DPSC transplantation were then developed. For example, they were transplanted into subcutaneous pockets on the backs of mice in a three-dimensional gelatin scaffold (2), and into a cranial defect in a collagen (3) or fibrin scaffold (4) for bone differentiation. The first method reported for transplantation required approximately 5.0  106 cells mixed with 40 mg hydroxyapatite/tricalcium phosphate (HA/TCP) (1). The number of cells and amount of scaffold required for transplantation was similar to that described for bone marrow stromal cells (BMSCs) transplantation (5). However, fewer dental pulp stem cells can be isolated for primary culture compared with BMSCs. A temperature-responsive tissue culture dish method has therefore been developed to reduce the cell number requirement for DPSC transplantation. This dish has been used in methods for the regeneration of several tissues and in a number of clinical studies (6–8). The dish enables cell sheets to be packed with cell pellets to form a three-dimensional structure. This protocol enables stem cell

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identification and in vivo transplantation using a small number of cells, HA/TCP particles, and a temperature-responsive cell culture dish.

2

Materials

2.1 Transplantation of DPSCs

1. UpCells®. 2. Ceraform (HA/TCP). 3. CellShifter™. 4. Human dental pulp stem cells (see Note 1). 5. Cellmatrix Type I-A. 6. Minimum Essential Medium Alpha Medium (αMEM). 7. Fetal bovine serum (FBS). 8. 100 μM L-ascorbic acid phosphate magnesium salt n-hydrate. 9. 2 mM L-glutamine. 10. 100 units/ml penicillin 100 μg/ml streptomycin. 11. 0.1–10 μl pipette tips. 12. Centrifuge tubes. 13. Faxitron MX-20 Specimen Radiography System. 14. MIN-R EV film. 15. Crl: NIH-Lyst Foxn1Btk (immunocompromised mouse, see Note 2). 16. Histostain® SP kits. 17. Anti-DMP1 antibody.

3

Methods

3.1 Primary DPSC Culture and UpCell® Culture

Dental pulp tissue was separated from teeth and cultured according to published methods (1), with some modifications. This protocol is for primary culture in a 75-cm2 flask. 1. Plate (2.0  106) primary DPSCs into each 75-cm2 flask (see Note 3). 2. Culture until confluent, changing growth medium three times per week. 3. Seed cells into UpCell® (6-well) plates at 1.0  105 cells/well. 4. After cells reach confluency, remove medium, place the CellShifter™ onto cells, and add 50 μl medium. 5. Incubate the 6-well plate at 4  C for 4 min (Fig. 1a, b; see Note 4). 6. Transfer the CellShifter™ onto collagen gel (see Note 5).

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Fig. 1 DPSC culture in temperature-responsive cell culture dishes. (a, b) Analysis of DPSC culture mineralization on temperature-responsive cell culture dishes by alizarin red staining. DPSCs were treated with growth medium as a control (a) and with mineralization medium for 3 weeks for differentiation (b). H&E staining of paraffin-embedded two-layered DPSC cell sheets made using a temperature-responsive cell culture dish (c, d). Two layers of nuclei can be observed at low (c) and high (d) magnification

Fig. 2 Diagram of the three-dimensional culture procedure. (a) The first DPSC cell sheet is placed onto collagen gel. (b) The DPSC pellet is placed onto the first DPSC sheet and Ceraform (HA/TCP) particles are placed around the pellet. (c) The pellet and Ceraform particles are covered by a second DPSC sheet

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3.2 Preparation of a Three-Dimensional DPSC Structure

A three-dimensional DPSC structure was prepared by placing a cell pellet between two cell sheets. 1. First place cell sheet onto collagen gel (Fig. 1c, d). 2. Prepare the DPSC pellet (see Note 6). 3. Place a cell pellet containing approximately 1.0–1.5  105 cells onto the cell sheet using a 0.1–10 μl pipette tip (see Note 7). 4. Place a few Ceraform particles around the cell pellet. 5. Cover the cell pellet and Ceraform particles with a second cell sheet (see Note 8; Fig. 2). 6. Culture overnight.

3.3 Transplantation of the ThreeDimensional DPSC Structure

1. Immunocompromised female beige mice (Crl: NIH-Lyst Foxn1Btk) were used as subcutaneous transplant recipients. 2. Anesthetize mice with pentobarbital at 30–50 mg/kg body weight. 3. Make a skin incision of about 1 cm in length on the dorsal surface of each mouse. 4. Form subcutaneous pockets in each incision (see Note 9). 5. Transplant a three-dimensional DPSC structure into each subcutaneous pocket (see Note 10). 6. Close incisions with surgical staples.

3.4 Analysis of the Regenerative Ability of Dentin/Pulp-Like Complex

1. Recover transplants 10–12 weeks post-transplantation (see Note 11). 2. Fix overnight in 4 % paraformaldehyde. 3. Decalcify for 2–3 weeks in 10 % EDTA. 4. Transfer transplants into 70 % ethanol and embed into a single paraffin block (see Note 12). 5. Deparaffinize sections, hydrate, and stain with hematoxylin and eosin. 6. Incubate for 1 h with primary antibody against dentin matrix protein 1 (DMP1) at room temperature. Use Histostain® SP kits for immunohistochemistry according to the manufacturer’s instructions (Fig. 3).

4

Notes 1. Lower third molars were obtained from adults at the Nippon Dental University Hospital in Tokyo under approved guidelines of the Ethics Committee of the School of Life Dentistry, Nippon Dental University, Tokyo.

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Fig. 3 Transplantation of the DPSC three-dimensional structure. (a) Faxitron analysis of the box region showing the three-dimensional DPSC structure. (b) Stereomicroscopy image of the three-dimensional DPSC structure (box region). (c) Low magnification image after H&E staining showing a dentin/pulp-like complex. (d, e) High magnification image of the H&E-stained box region showing odontoblasts and odontoblast processes (arrow), vessels (dashed arrow), dentin (star) (d), stained with a DMP1 antibody to show odontoblasts (arrow) (e)

2. This experiment was performed in accordance with guidelines for the care and use of laboratory animals of the School of Life Dentistry, Nippon Dental University, Tokyo. 3. Tooth surfaces were wiped with 70 % ethanol and cut around the cementum-enamel junction using a sterilized dental diamond point. After teeth were cracked, the pulp tissue was removed and digested in phosphate-buffered saline (PBS) containing 3 mg/ml collagenase type I and 4 mg/ml dispase for 1.5 h at 37  C. 4. The cells were incubated for 3–4 weeks in αMEM supplemented with 10 % FBS, 10 mM sodium β-glycerophosphate nhydrate, 10 nM dexamethasone, and the same concentrations of penicillin and streptomycin as in mineralization growth medium. After washing in PBS, cells were fixed with 4 % paraformaldehyde and stained with 2 % Alizarin red S. 5. Before removing the CellShifter™, aspirate to completely remove the medium. Otherwise, the DPSC sheet will not attach to the CellShifter™ surface. 6. Prepare DPSCs in another culture dish or flask to make the cell pellet. To harvest DPSCs, remove medium, wash twice with PBS, and treat with 0.25 % EDTA for 3 min. After harvesting, transfer to a tube and centrifuge (440  g, 5 min, 4  C).

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Remove the supernatant and add 1 ml PBS to make a cell suspension. Transfer to a 1.5 ml tube and centrifuge (600  g 3 min, 4  C). Remove the supernatant and recentrifuge (600  g, 3 min, 4  C). Completely remove supernatant using a micropipette. 7. Transfer the cell pellet onto the cell sheet using a micropipette. Release the cell pellet very gently; otherwise the cell sheet will be destroyed. 8. Before covering with the second cell sheet, remove all medium from the culture dish to ensure correct attachment of the second cell sheet to the first cell sheet, cell pellet, and Ceraform particles. After covering with the second cell sheet, add 500 μl medium and incubate for 30 min. Remove medium before removing the CellShifter™. 9. Form the pockets using curved scissors, with a gentle action. If the procedure is done correctly, almost no bleeding will occur. 10. The three-dimensional DPSC culture is removed from the culture dish using sharp scissors under a stereomicroscope. Sharp tweezers are useful for transplanting the threedimensional culture. 11. Faxitron analysis, stereomicroscope observation, and transplant recovery are usually performed in euthanatized animals. Faxitron analysis and stereomicroscope observation can be done under anesthesia. 12. Paraffin embedding of the cell sheet is performed using warm tweezers for orientation. The paraffin block is cut into 5-μm sections.

Acknowledgment This work was supported by a Research Grant from the Nippon Dental University. References 1. Gronthos S, Mankani M, Brahim J, Robey PG et al (2000) Postnatal human dental pulp stem cells (DPSCs) in vitro and in vivo. Proc Natl Acad Sci U S A 97:13625–13630 2. Li JH, Liu DY, Zhang FM, Wang F et al (2011) Human dental pulp stem cell is a promising autologous seed cell for bone tissue engineering. Chin Med J (Engl) 124:4022–4028 3. Maraldi T, Riccio M, Pisciotta A, Zavatti M et al (2013) Human amniotic fluid-derived and dental pulp-derived stem cells seeded into collagen scaffold repair critical-size bone defects promoting vascularization. Stem Cell Res Ther 4:53

4. Riccio M, Maraldi T, Pisciotta A, La Sala GB et al (2012) Fibroin scaffold repairs critical-size bone defects in vivo supported by human amniotic fluid and dental pulp stem cells. Tissue Eng Part A 18:1006–1013 5. Kuznetsov SA, Krebsbach PH, Satomura K, Kerr J et al (1997) Single-colony derived strains of human marrow stromal fibroblasts form bone after transplantation in vivo. J Bone Miner Res 12:1335–1347 6. Nishida K, Yamato M, Hayashida Y, Watanabe K et al (2004) Corneal reconstruction with tissue-engineered cell sheets composed of

Transplantation of Reduced Cell Number Methods autologous oral mucosal epithelium. N Engl J Med 351:1187–1196 7. Ohki T, Yamato M, Ota M, Takagi R et al (2012) Prevention of esophageal stricture after endoscopic submucosal dissection using tissueengineered cell sheets. Gastroenterology 143:582–588, e1–2

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8. Egami M, Haraguchi Y, Shimizu T, Yamato M et al (2014) Latest status of the clinical and industrial applications of cell sheet engineering and regenerative medicine. Arch Pharm Res 37:96–106

Methods in Molecular Biology (2015) 1212: 209–210 DOI 10.1007/7651_2015 © Springer Science+Business Media New York 2015

INDEX A Adipose tissue-derived stem cell.......................... 171–180 AGM. See Aorta-gonads-mesonephros (AGM) Alzheimer’s disease (AD)...............................87, 171–180 β-amyloid ..................................................... 172, 173, 180 Antibody microarray .................................................41, 47 Aorta-gonads-mesonephros (AGM) ................... 183–186

Endothelial cells ................................................... 113, 123 ESCs. See Embryonic stem cell (ESCs) Exosome ........................................................................ 175 Expansion ................................... 65–71, 77, 87–124, 138 Extracellular matrix .................... 21–36, 39, 66, 143, 157 Extracellular vesicle .............................................. 171–180

F

B

Fluo-4 AM............................................................ 164–168

Biomechanical forces............................................ 183, 184

G

C

Glutaraldehyde (GA) .............................. 5, 22–28, 32–35 Growth factor ....................................... 39–63, 67, 88, 90, 104, 106, 109, 115, 129, 172

Ca2+ ....................................................................... 163–168 Cardiomyocytes...................................113, 123, 163–168 Cardiovascular progenitor cells (CVPCs) ........... 113–124 Chemically defined medium......................................... 114 CM. See Conditioned medium (CM) Coculture........................................ 2, 171, 173, 177–179 Colon ........13, 71, 79, 80, 94, 101, 120, 121, 123, 124, 131, 132, 135–137, 141–159, 165, 189 Conditioned medium (CM)........ 46, 47, 53, 57–60, 174 Confocal microscopy ................. 146–147, 154, 166–168 Cryopreservation................................................ 12, 14, 19 Crypt..................................................................... 141–159 Culture..............................................1–20, 39–43, 46, 52, 57, 59, 60, 62, 65–71, 73–84, 88, 89, 94–96, 98, 99, 101, 102, 104–109, 111, 112, 114–119, 122, 123, 128, 129, 131–137, 141–159, 164–166, 172–174, 177–180, 183–185, 187, 189 Cupromeronic blue............................... 22–24, 27, 34, 35 CVPCs. See Cardiovascular progenitor cells (CVPCs) Cytokine .................................................. 39–63, 152, 172

D Derivation............................................ 73, 77–78, 87–102 Differentiation........................................1, 21, 39, 40, 65, 73–84, 87–124, 127–136, 163–166 Directed differentiation .................................................. 82

E Embryoid bodies (EB)............................... 83, 88, 90, 92, 94, 95, 101, 163, 166, 168 Embryonic/fetal fibroblasts ................................ 2, 17, 18 Embryonic stem cell (ESCs) ........................2, 65, 73–84, 88, 114, 163–168, 184

H Hematopoietic stem cells (HSC) ........................ 183–192 Hemogenic endothelium..................................... 183–192 hiPSCs. See Human induced pluripotent stem cells (hiPSCs) hPSCs. See Human embryonic stem cell (hESC) HSC. See Hematopoietic stem cells (HSC) Human.................................................. 40, 60, 73, 88–90, 106, 114, 128, 135–138, 141–159, 163, 171–180, 185 Human embryonic stem cell (hESC)..................... 65, 88, 113, 114, 123 Human induced pluripotent stem cells (hiPSCs)......... 65, 88, 113, 114, 124 Human pluripotent stem cell (hPSC).....................65–71, 87–102, 113–124

I Imaging........................ 42, 144–146, 151–154, 156, 165 Immunocytochemistry............................. 76, 82–83, 109, 144, 146–148, 152–156, 159 In situ hybridization .................................. 144, 147–148, 154–156, 159 Intercellular communication ........................................ 163 Interstitial interface ............................................ 22, 32–35 Intestine ........................................................................... 41

K Kidney.................................... 22, 23, 27, 28, 30, 33, 172

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STEM CELL RENEWAL 210 Index

AND

CELL-CELL COMMUNICATION

L

R

Labware and equipment ...........................................2, 7, 8 Luminex xMAP ............................................................... 40

Rat...................................... 40, 60, 73–84, 128, 135–137 Ruthenium red ................................22–24, 27–28, 34, 35

M

S

Mesenchymal stem cell (MSC).................... 40, 128, 137, 171–180 Mesenchymal stem cell isolation ......................... 127–139 Microvesicle ................................................................... 171 Migration..................................................... 144, 151, 152 Mitogens...................................................... 105, 110, 143 Mouse embryonic stem cells ............................... 163–168 MSC. See Mesenchymal stem cell (MSC) Multipotency .......................................104, 109–110, 172 Mutiplex immunoassays............................................39–63 Mycoplasmosis ................................2, 3, 8, 10–11, 15–17

Seeding density....................................111, 128, 135, 136 Self-renewal .............................1–20, 65, 66, 80, 84, 103, 104, 109, 127 Serum-free medium (SFM) ........... 46, 57, 172–174, 179 SEZ. See Subependymal zone (SEZ) SFM. See Serum-free medium (SFM) Shear stress ........................................................... 184–191 Signaling ...................................... 74, 81–82, 88, 92, 115, 143, 144, 153, 183, 184 Smooth muscle cells (SMC) ...................... 113, 114, 116, 122–123 Standard operation protocol (SOP).........................42–43 Stem cell characterization .................................... 135, 137 Stem cell culture........................................6–7, 11–14, 17, 40, 123, 137 Stem cells .......................................... 1–20, 39–63, 65–71, 89, 103, 104, 113–124, 127–139, 141–159, 163–168, 171–180, 183–192 Stem/progenitor cell niche ......................................21–36 Subependymal zone (SEZ)........................ 103–105, 107, 108, 111 Subpassage ....................................................................... 16

N Neprilysin....................................................................... 172 Neural progenitor cells (NPC)...............................87–102 Neural stem cells (NSC) ...................................... 103–124 Neurogenic niche ................................................. 103, 104 Neuronal differentiation ...........................................90, 91 NPC. See Neural progenitor cells (NPC) NSC. See Neural stem cells (NSC)

P Porous membrane (PM)...........................................65–71 Progenitor maintenance ...................................... 113–114 Proliferation........................ 2, 21, 39, 74, 103, 113, 137, 144, 151, 171 Proliferative human feeder cell ....................................... 65 Propagation ............................73–84, 119–121, 124, 136 Protein secretion .......................................................39–40 Protocol .........................................................1–20, 22, 23, 42, 54, 56, 74, 81, 88, 89, 92, 93, 114, 120, 123, 128, 144, 156, 158, 159, 164, 174, 179

T Tannic acid (TA) ............................ 22–25, 28, 32, 34, 35 Tendon-derived stem/progenitor cells (TDSC)........128, 129, 131–138 Tissue culture ....................................... 75–77, 80, 81, 83, 89, 103, 116, 117, 165, 166, 189 Tissue renewal ...................................................... 141–159 Transmission electron micros-copy..........................21–36